&EPA
united States
Environmental Protection
Agency
Office of Pesticides &
Toxic Substances
Washington DC 20460
EPA 560/6-82-002
PB82-232992
August 1982
Toxic Substances
Environmental Effects
Test Guidelines
Part Two
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ENVIBONMENTAL EFFECTS TEST GUIDELINES
Part Two
FISH EARLY LIFE STAGE TOXICITY TEST
OFFICE OF TOXIC SUBSTANCES
OFFICE. OF PESTICIDES AND TOXIC SUBSTANCES
U.S. ENVIRONMENTAL PROTECTION AGENCY
WASHINGTON, D.C. 20460
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Office of Toxic Substances EG-11
Guideline for Testing Chemicals August, 1982
FISH EARLY LIFE STAGE TOXICITY TEST
('a) Purpose. This guideline is intended to be used for
assessing the propensity of chemical substances to produce
adverse effects to fish during the early stages of their growth
and development. This guideline describes the conditions and
procedures for the continuous exposure of several representative
species to a chemical substance during egg, fry and early
juvenile life stages. .The Environmental Protection Agency (EPA)
will use data from this test in assessing the potential hazard of
the test substance to the aquatic environment.
(b) Definitions. The definitions in section 3 of the Toxic
Substances Control Act (TSCA) and the definitions in Part 792—
Good Laboratory Practice Standards, apply here. In addition, the
following definitions are applicable to this specific test
guideline:
(1) "Acclimation" physiological or behavioral adaptation of
organisms to one or more environmental conditions associated with
the test method (e.g., temperature, hardness, pH) .
(2) "Carrier" solvent or other agent used to dissolve or
improve the solubility of the test substance in dilution water.
(3) "Conditioning" exposure of construction materials, test
chambers, and testing apparatus to dilution water or to the test
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solution prior to the start of the test in order to minimize the
sorption of test substance onto the test facilities or the
leaching of substances from test facilities into the dilution
water or the test solution.
(4) "Control" an exposure of test; organ isms to dilution
water only or dilution water containing the test solvent or
carrier (no toxic agent is intentionally or inadvertently
added).
(5) "Dilution water" the water used to produce the flow-
through conditions of the test to which the test substance'"-Is"
added and to which the test species is exposed.
(6) "Early life stage toxicity test" a test to determine the
minimum concentration of a substance which produces a
statistically significant observable effect on hatching,
survival, development and/or growth of a fish species
continuously exposed during the period of their early
development. • ;
(7) "Embryo cup" a small glass jar or similar container with
a screened bottom in which the embryos of some species (i.e.,
minnow) are placed during the incubation period and which is
normally oscillated to ensure a flow,of water through the cup.
(8) "Flow through" refers to the continuous or ve^ry frequent
passage of fresh test solution through a test chamber with no
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recycling.
(9) "Hardness" the total concentration of the calcium and
magnesium ;Lons in water expressed as calcium carbonate (mg
CaCOyiiter) .
(10) "Loading" the ratio of biomass (grams of fish, wet
weight) to the volume (liters) of test solution passing through
the test chamber during a specific interval (normally a 24-hr.
period).
(11) "No observed effect concentration (NOEC)" the highest
tested concentration in an acceptable early life stage test: (a)
which did not cause- the occurrance of any specified adverse
effect (statistically different from the control at the 95%
level); and (b) below which no tested concentration caused such
an occurrence.
(12) "Observed effect concentration (OEC)" the lowest tested
concentration in an acceptable early life stage test: (a) which
caused the occurrence of any specified adverse effect
(statistically different from the control at the 95% level); and
(b) above which all tested concentrations caused such an
occurrence.
(13) "Replicate" two or more duplicate tests, samples,
organisms, concentrations, or exposure chambers.
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(14) "Stock solution" the source of the test solution
prepared by dissolving the test substance in dilution water or a
carrier which is then added to dilution water at a specified,
selected concentration by means of the test substance delivery
sys tern.
(15) "Test chamber" the individual containers in which test
organisms are maintained during exposure to test solution.
(16) "Test solution" dilution, water with a test substance
dissolved or suspended in it.
(17) "Test substance" the specific form of a chemical
substance or mixture that is used to develop data.
(c) Test Procedures — (1) Summary of test. (i) The early
life stage toxicity test with fish involves exposure of newly
fertilized embryos to various concentrations of a test
substance. Exposure continues for 28 days post hatch for the
minnows and 60 days post hatch for the trout species. During
this time various observations and measurements are made in a
specific manner and schedule in order to determine the lowest
effect and highest no-effect concentrations of the test
subs tance.
(ii) A minimum of five exposure (treatment) concentrations
of a test substance and one control are required to conduct an
early life stage toxicity test. The concentration of tne test
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substance in each treatment is usually 50% of that in the next
higher treatment level.
(iii) For each exposure concentration of the test substance
and for each control (i.e., regular control and carrier control
if required) there should be:
(A) At least two replicate test chambers, each containing
one Or more embryo incubation trays or cups; and there should be
no water connections between the replicate test chambers;
(B) At least 60 embryos divided equally, through
randomization, between the embryo incubation trays or cups for
each test concentration and control (i.e., 30 per embryo cup with
2 replicates);
(C) All surviving larvae divided equally between the test
chambers for each test concentration and control (e.g., 30 larvae
per test chamber with 2 replicates).
(iv) Duration. (A) For fathead minnow and sheepshead
minnow a test begins when the newly fertilized minnow embryos
(less than 48-hours old) are placed in the embryo cups and are
exposed to the test solution concentrations. The test terminates
following 28 days of post-hatch exposure, i.e., 28 days after the
newly hatched fry are transferred from the embryo cups into the
test chambers.
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(B) For brook trout and rainbow trout a test begins when
newly fertilized trout embryos (less than 96-hours old) are
placed in the embryo trays or cups and are exposed to the test
solution concentrations. The test terminates following 60 days
of post-hatch exposure (for an approximate total exposure period
of 90 days ) .
(2) [Reserved]
(3) Range-finding test. (i) A range finding test is
normally performed with the test substance to determine the test
concentrations to be used in the early life stage toxicity test,
especially when the toxicity is unknown. It is recommended that
the test substance concentrations be selected based on
information gained from a 4- to 10-day flow-through toxicity test
with juveniles of the selected test species.
(ii) The highest concentration selected for the early life
stage toxicity test should approximate the lowest concentration
indicated in any previous testing tc cause a significant
reduction in survival. The range of concentrations selected is
expected to include both observed effect and no-observed- effect
levels. The dilution factor between concentrations is normally
0.50, however, other dilution factors may be used as necessary.
(4) Definitive test—(i) General. (A) A test should not be
initiated until after the test conditions have been met and the
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test substance delivery system has been observed functioning
properly for 48-hours. This includes temperature stability, flow
requirements of dilution water, lighting requirements, and the
function of strainers and air traps included in the water-supply
system, and other conditions as specified previously.
(B) New holding and testing facilities should be tested with
sensitive organisms (i.e., juvenile test species or daphnids)
before use to assure that the facilities or substances possibly
leaching from the equipment will not adversely affect the test
organisms during an actual test.
(C) Embryos should be acclimated for as long as practical to
the test temperature and dilution water prior to the initiation
of the test.
(D) When embryos are received from an outside culture source
(i.e. rainbow and brook trout) at a temperature at variance with
the recommended test temperature they should be acclimated to the
test temperature. When eggs are received, they should be
immediately unpacked and the temperature of the surrounding water
determined. Sudden temperature changes should be avoided.
Acclimation to the appropriate test temperature should be
accomplished within a period of six hours, and should incorporate
the use of dilution water.
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(E) Embryos should be visually inspected prior to placement
in the embryo cups or screen trays. All dead embryos should be
discarded. Dead embryos can be discerned by a change in
coloration from that of living embryos (e.g. trout embryos turn
white when dead). During visual inspection, empty shells, opaque
embryos and embryos with fungus or partial shells attached should
be removed and discarded. If less than 50 percent of the eggs to
be used appear to be healthy, all embryos in such a lot should be
discarded.
(ii) Embryo incubation procedures. (A) Embryos can be
distributed to the embryo cups or screen trays using a pipette
with a large bore or a similar apparatus. Trout embryos can be
distributed by using a small container which has been
precal ibrated to determine the approximate number of embryos it
can hold; embryos are measured volumetr ically in this manner, and
are then poured onto the screen tray (or embryo cup). Trout
embryos should be separated on the screen tray so that they are
not in contact with each other. A final count will ensure the
actual number on the screen tray. After random assignmment, the
screen trays or embryo cups are placed in the test chambers.
(B) Each day until hatch the embryos are visually examined.
Minnow embryos may be examined with the aid of a magnifying
viewer. Trout embryos should not be couched. Trout embryos
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should be maintained in low intensity light or in darkness until
one-week post hatch, and are usually examined with the aid of a
flashlight or under low intensity light. Dead embryos should be
removed and discarded. Live embryos which are heavily infected
with fungus should be discarded, but should be subtracted from
the initial number of embryos used as a basis for the
calculations of percentage hatch.
(C) When embryos begin to hatch they should not be handled.
(iii) Initiation of fry exposure. (A) Forty-eight hours
after the first hatch in each treatment level, or when hatching
is completed, the live young fish should be counted and
transferred from each embryo cup into the appropriate test
chamber. All of the normal and abnormal fry should be gently
released into the test chamber by allowing the fry to swim out of
each embryo cup; nets should not be used. The trout embryos
incubated on screen trays will hatch out in the test chambers,
therefore handling of fish is not necessary.
(B) If necessary, fry can be transferred from one replicate
embryo cup to the other replicate within a test concentration to
achieve equal numbers in each replicate chamber.
(C) The number of live fry, live normal fry, live embryos,
dead embryos and unaccounted for embryos for each cup should be
recorded when hatching is deemed complete. Those fry which are
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visibly (without the use of a dissecting scope or magnifying
viewer) lethargic or grossly abnormal (either in swimming
behavior or physical appearance) should be counted. Late
hatching embryos should be left in the embryo cups to determine
if they will eventually hatch or not. The range of time-to-hatch
(to the nearest day) for each cup should be recorded.
( iv) Time to first feeding. (A) The first feeding for the
fathead and sheepshead minnow fry should begin shortly after
transfer of the fry from the embryo cups to the test chambers.
Trout species initiate feeding at swim-up. The trout fry should
be fed trout starter mash three times a day ad libitum, with
excess food siphoned off daily. The minnow fry should be fed
live newly-hatched brine shrimp nauplii (Artemia salina) at least
three times a day.
(B) For the first seven days, feeding should be done at
minimum intervals of four hours (i.e., 8 am, 12 noon, and 4 pm);
thereafter the fry should be fed as indicated below.
(v) Feeding. (A) The fathead and sheepshead minnow fry
should be fed newly-hatched brine shrimp nauplii for the duration
of the test at approximately 4-hour intervals three times a day
during the week and twice on the weekend after the first week.
Trout fry should be fed at similar intervals, and may receive
live brine shrimp nauplii in addition to the trout starter food
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after the first week.
(B) An identical amount of food should be provided to each
chamber. Fish should be fed ad libitum for 30 minutes with
excess food siphoned off the bottom once daily if necessary.
(C) Fish should not be fed for the last 24 hours prior to
termination of the test.
(vi) Carriers . Water should be used in making up tne test
stock solutions. If carriers other than water are absolutely
necessary, the amount used should be the minimum necessary to
achieve solution of the test substance. Triethylene glycol and
dimethyl formamide are preferred, but etnanol and acetone can be
used if necessary. Carrier concentrations selected should be
kept constant at all treatment levels.
(vii) Controls. Every test requires a control that consists
of the same dilution water, conditions, procedures, and test
organisms from the same group used in the other test chambers,
except that none of the test substance is added. If a carrier
(solvent) is used, a separate carrier control is required in
addition to the regular control. The carrier control should be
identical to the regular control except that the highest amount
of carrier present in any treatment is added to this control. If
the test substance is a mixture, formulation, or commercial
product, none of the ingredients is considered a carrier unless
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an extra amount is used to prepare the stock solution.
(viii) Randomization. The location of all test chambers and
species within the test system should be randomized. A
representative sample of the test embryos should be impartially
distributed by adding to each cup or screen tray no more than 20%
of the number of embryos to be placed in each cup or screen tray
and repeating the process until each cup or screen tray contains
the specified number of embryos. Alternatively, the embryos can
be assigned by random assignment of a small group (e.g., 1-5) of
embryos to each embryo cup or screen tray, followed by random
assignment of a second group of equal number to each cup or tray,
which is continued until the appropriate number of embryos are
contained in each embryo cup or screen tray. The method of
randomization used should be reported in detail.
( ix) Observations. During the embryo exposure period
observations should be made to check for nvortality. During the
exposure period of the fry, observations should be made to check
for mortality and to note the physical appearance and behavior of
the young fish. The biological responses are used in combination
with physical and chemical data in evaluating the overall lethal
and sublethal effects of the test substance. Additional
information on the specific methodology for the data obtained
during the test procedure are discussed in the following
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sections.
(x) Biological data. (A) Death of embryos should be
recorded daily and dead embryos removed when discovered to
prevent the spread of fungal contamination.
(B) When hatching commences, daily records of the number of
embryos remaining in each embryo cup are required. This
information is necessary to quantify the hatching success. A
record of all deformed larvae should be kept throughout the
entire post-hatch exposure. Time to swim-up should be recorded
for; the trout. Upon transfer of fry from the embryo cups to the
test chambers, daily counts of the number of live fish should be
made. At a minimum, live fish should be counted on days 4, 11,
18, 25 and (weekly thereafter for the trout species) finally on
termination of the test.
(C) The criteria for death of young fish is usually
immobility, especially absence of respiratory movement, and lack
of reaction to gentle prodding. Deaths should be recorded daily
and dead fish removed when discovered.
(D) Daily and at termination of the test, the number of fish
that appear (without the use of a magnifying viewer) to be
abnormal in behavior (e.g., swimming erratic or uncoordinated,
obviously lethargic, hyperventilating, or over excited, etc.) or
in physical appearance (e.g., hemorrhaging, producing excessive
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mucous, or are discolored, deformed, etc.) should be recorded and
reported in detail.
(E) All physical abnormalities (e.g., stunted bodies,
scoliosis, etc.) should be photographed and the deformed fish
which die, or are sacrificed at the termination of the test,
should be preserved for possible future pathological examination.
(F) At termination, all surviving fish should be measured
for growth. Standard length measurements should be made directly
with a caliper, but may be measured photographically.
Measurements should be made to the nearest millimeter (O.lmm is
desirable). Weight measurements should also be made for each
fish alive at termination (wet, blotted dry and to the nearest
O.Olg for the minnows and 0. Ig for the trout). If the fish
exposed to the toxicant appear to be edematous compared to
control fish, determination of dry, rather than wet, weight is
recommended.
(G) Special physiological, biochemical and histological
investigations on embryos, fry, and juveniles may be deemed
appropriate and should be performed on a case by case basis.
(5) Test results . (i) Data from toxicity tests are usually
either continuous (e.g. length or weight measurements) or
dichotomous (e.g. number hatching or surviving) in nature.
Several methods are available and acceptable for statistical
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anaylsis of data derived from early life stage toxicity tests;
however, the actual statistical methodology to analyze and
interpret the test results should be reported in detail.
(ii) The significance level for all statistical testing
should be a minimum of P=0.05 (95 percent confidence level).
(A) Example of statistical analysis. (JJ Mortality data
for the embryonic stage, fry stage and for both stages in
replicate exposure chambers should first be analyzed using a two-
way analysis of variance (ANOVA) with interaction model. This
analysis will determine if replicates are significantly different
from each other. If a significant difference between replicates
or a significant interaction exists, cause for the difference
should be determined. Modification should then be made in the
test apparatus or in handling procedures for future toxicity
tests. Further calculations should incorporate the separation of
replicates. If no significant difference is observed, replicates
may be pooled in further analyses.
(_2_) After consideration of replicate responses, mortality
data should then be subjected to one-way ANOVA. The purpose of
this analysis is to determine if a significant difference exists
in the percentage mortality between control fish and those
exposed to the test material.
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(_3_) If the one-way ANOVA results in a F ratio that is
significant, it would be acceptable to perform t-tests on the
control versus each concentration. A second technique is to
identify treatment means that are significantly different; this
method should involve the additional assumption that the true
mean response decreases generally with increasing
concentration. The researcher may also be interested in
determining significant differences between concentrations.
(_4_) Growth data should also be analyzed by one-way ANOVA
with the inclusion of a covariate to account for possible
differences in growth of surviving fry in embryo cup(s) that
contain fewer individuals. This condition can occur in cases
when the same amount of food is given to each test chamber
regardless of the number of survivors.
(B) Test data to be analyzed. Data to be statistically
analyzed are:
(_!_) Percentage of healthy, fertile embryos at 40-48
hours after initiation of the test. Percentage is based upon
initial number used.
(2} Percentage of embryos that produce live fry for
release into test chambers. Percentage is based on number of
embryos remaining after thinning.
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(_3_) Percentage of embryos that produce live, normal
fry for release into test chambers. Percentage is based upon
number of embryos remaining after thinning.
(_4_) Percentage of fry survival at swim-up for trout.
Percentage is based upon number of embryos remaining after
t h i n n i ng .
(J5_) Percentage of embryos that produce live fish at
end of test. Percentage is based upon number of embryos remaining
after thinning.
(_6_) Percentage of embryos that produce live, normal
fish at end of test. Percentage is based upon number of embryos
remaining after thinning.
(_7_) Weights and lengths of individual fish alive at
the end of the test
(C) It is important that fish length and weight measurements
be associated with individual test chambers since the density of
the fish and available food should be considered in the growth of
the organism.
(iii) Acceptability criteria. (A) An early life stage
toxicity test is not acceptable unless at least one of the
following criteria is significantly different (p=0.05) from
control organisms when compared with treated organisms, and the
responses are concentration-dependent: mortality of embryos,
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hatching success, mortality of fry (at swim-up for trout), total
mortality throughout the test, and growth (i.e. weight). If no
significant effects occur, but the concentrations tested were the
highest possible due to solubility or other physio-chemical
limitations, the data will be considered for acceptance.
(B) In addition to obtaining significant effects on the
exposed test species, a measure of acceptablity in the response
of control fish is also required.
(C) A test is not acceptable if the average survival of the
control fish at the end of the test is less than 80 percent or it
survival in any one control chamber is less than 70 percent.
(D) If a carrier is used, the criteria for effect (mortality
of embryos and fry, growth, etc.) used in the comparison of
control and exposed test organisms should also be applied to the
control and control with carrier chambers. For the test to be
considered acceptable, no significant difference should exist
between these criteria.
(E) A test is not acceptable if the relative standard
deviation (RSD=100 times the standard deviation divided by the
mean) of the weights of the fish that were alive at the end of
the test in any control test chamber is greater than 40 percent.
(6) Analytical measurements--( i) Analysis of water
quality. Measurement of certain dilution water quality
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parameters should be performed every 6 months, to determine the
consistency of the dilution water quality. In addition, if data
in 30 day increments are not available to show that freshwater
dilution water is constant, measurements of hardness, alkalinity,
pH, acidity, conductivity, TOG or COD and particulate matter
should be conducted once a week in the control and once a week in
the highest test substance concentration. Measurement of
calcium, magnesium, sodium, potassium, chloride, and sulfate is
des irable.
(ii) Dissolved oxygen measurement. The dissolved oxygen
concentration should be measured in each test chamber at the
beginning of the test and at least once daily thereafter (as long
as live organisms are present) in one replicate of the control
and the high, medium, and low test substance concentrations.
(iii) Temperature measurement. Temperatures should be
recorded in all test chambers at the beginning of the test, once
weekly thereafter and at least hourly in one test chamber. When
possible, the hourly measurement should be alternated between
test chambers and between replicates.
(iv) Test substance measurement. (A) Prior to the addition
of the test substance to the dilution, water, it is recommended
that the test substance stock solution be analyzed to verify the
concentration. After addition of the test substance, the
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concentration of test substance should be measured at the
beginning of the test in each test concentration (including both
replicates) and control(s), and in one replicate at each test
concentration at least once a week thereafter. Replicates should
be alternated each week. If a malfunction in the delivery system
is discovered, water samples should be taken from the affected
test chambers immediately and analyzed.
(B) The measured concentration of test substance in any
chamber should be no more than 30% higher or lower than the
concentration calculated from the composition of the stock
solution and the calibration of the test substance delivery
system. If the difference is more than 30%, the concentration of
test substance in the solution flowing into the exposure chamber
(influent) should be analyzed. These results will indicate
whether the problem is in the stock solution, the test substance
delivery system or in the test chamber. Measurement of
degradation products of the test substance is recommended if a
reduction of the test substance concentration occurs in the test
chamber.
(v) Sampling and analysis methodology. (A) Generally,
total test substance measurements are sufficient; however, the
chemical characteristics of the test substance may require both
dissolved and suspended test substance measurements.
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(B) For measurement of dissolved or suspended test substance
or both, water samples should be taken midway between the top,
bottom, and sides of the test chamber and should not include any
surface scum or material stirred up from the bottom or sides.
For measurement of total test substance, a large volume of the
solution in the test chamber should be collected and used as the
sample. Samples of test solutions should be handled and stored
appropriately to minimize loss of test substance by microbial
degradation, photodegradation, chemical reaction, volatilization,
or sorption.
(C) Chemical and physical analyses should be performed using
standardized methods whenever possible. The analytical method
used to measure the concentration of the test substance in the
test solution should be validated before the beginning of the
test. At a minimum, a measure of the accuracy of the method
should be obtained, on each of two separate days by using the
method of known additions, and using dilution water from a tank
containing test organisms. Three samples should be analyzed at
the next to lowest test substance concentration. It is also
desirable to study the accuracy and precision of the analytical
method for test guideline determination by use of reference
(split) samples, or interlaboratory studies, and by comparsion
with alternative, reference or corroborative methods of analysis.
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(D) An analytical method is not acceptable if likely
degradation products of the test substance, such as hydrolysis
and oxidation products, give positive or negative interferences,
unless it is shown that such degradation products are not present
in the test chambers during the test. In general, atomic
absorption spectrophotometric methods for metals and gas
chromatographic methods for organic compounds are preferable to
colorimetrie methods.
(E) In addition to analyzing samples of test solution, at
least one reagent blank also should be analyzed when a reagent is
used in the analysis. Also, at least one sample for the method
of known additions should be prepared by adding test substance at
the concentration used in the toxicity test.
(d) Test conditions--!1) Test species. (i) One or more of
the recommended test species will be specified in rules under
Part 799 requiring testing of specific chemicals. The
recommended test species are:
(A) Fathead minnow (Pimephales promelas Rafinesque).
(B) Sheepshead minnow (Cyprinodon variegatus).
(C) Brook trout (Salvelinus fontinalis)..
(D) Rainbow trout (Salmo gairdneri).
(ii) Embryos used to initiate the early life stage test
should be less than 48-hours old for the fathead and sheepshead
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minnows, and less than 96-hours old for the brook trout and
rainbow trout. In addition, the following requirements should oe
met:
(A) All embryos used in the test should be from the same
source. Embryos should be obtained from a stock cultured in-
house when possible, and maintained under the same parameters as
specified for the test conditions. When it is necessary to
obtain embryos from an external source, caution should be
exercised to ensure embryo viability and to minimize the
possibility of fungal growth. A description of the brood stock
history or embryo source should be made available to EPA upon
reques t.
(B) Test species should be cared for and handled properly in
order to avoid unnecessary stress. To maintain test species in
good condition and to maximize growth, crowding should be
prevented, and the dissolved oxygen level should be maintained
near saturation.
(C) Embryos and fish should be handled as little as
possible. Embryos should be counted and periodically inspected
until hatching begins. When larvae begin to hatch, they should
not be handled. Transfer of minnow larvae from embryo cups to
test chambers should not involve the use of nets. No handling is
necessary following introduction into the test chambers until
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termination of the test.
(D) If fathead minnow embryos are obtained from in-house
culture units, the embryos should be gently removed from the
spawning substrate. The method for separating the fertilized
eggs from the substrate is important and can affect the viability
of the embryos; therefore the finger-rolling procedure is
recommended.
(E) Disease treatment. Chemical treatments to cure or
prevent diseases should not be used before, and should not be
used during a test. All prior treatments of brood stock should
be reported in detail. Severely diseased organisms should be
des troyed.
( 2 ) Test facilities — ( i ) Construction materials.
Construction materials and equipment that contact stock
solutions, test solutions, or dilution water into which test
embryos or fish are placed should not contain any substances that
can be leached or dissolved into aqueous solutions in quantities
that can affect test results. Materials and equipment that
contact stock or test solutions should be chosen to minimize
sorption of test chemicals from dilution water. Glass, #316
stainless steel, nylon screen and perfluorocarbon plastic (e.g.,
Teflon®) are acceptable materials. Concrete or rigid
(unplas tic ized) plastic may be used for holding and acclimation
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tanks, and for water supply systems, but they should be
thoroughly conditioned before use. If cast iron pipe is used in
freshwater supply systems, colloidal iron may leach into the
dilution water and strainers should be used to remove rust
particles. Natural rubber, copper, brass, galvanized metal,
epoxy glues, and flexible tubing should not come in contact witn
dilution water, stock, solutions, or test solutions.
(ii) Test chambers (exposure chambers). (A) Stainless
steel test chambers should be welded or glued with silicone
adhesive, and not soldered. Glass should be fused or bonded
using clear silicone adhesive. Epoxy glues are not recommended,
but if used ample curing time should be allowed prior to use. .As
Little adhesive as possible should be in contact with the water.
(B) Many different sizes of test chambers have been used
successfully. The size, shape and depth of the test chamber is
acceptable if the specified flow rate and loading requirements
can be achieved.
(C) The actual arrangement of the test chambers can be
important to the statistical analysis of the test data. Test
chambers can be arranged totally on one level (tier) side by
side, or on two levels with each level having one of the
replicate test substance concentrations or controls. Regardless
of the arrangement, it should be reported in detail and
considered in the data analysis.
25
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EG-11
August, 1982
(iii) Embryo incubation apparatus. (A) recommended embryo
incubation apparatus include embryo cups for the minnow species
and screen trays for the trout species, although embryo cups can
be used for the trout species. Embryo cups are normally
constructed from approximately 4-5 cm inside diameter, 7-8 cm
high, glass jars with the end cut off or similar sized sections
of polyethylene tubing. One end of the jar or tubing is covered
with stainless steel or nylon screen (approximately 40 meshes per
inch is recommended). The embryo cups should be appropriately
labeled and then suspended in the test chamber in such a manner
as to ensure that the test solution regularly flows through the
cup and that the embryos are always submerged but are not
agitated too vigorously. Cups may be oscillated by a rocker arm
apparatus with a low rpm motor (e.g., 2 rpm) to maintain the
required flow of test water. The vertical-travel distance of the
rocker arm apparatus during oscillation is normally 2.5 -
4.0cm. The water level in the test chambers may also be varied
by means of a self-s tarting siphon in order to ensure exchange of
water in the embryo cups.
(B) The trout embryo incubation trays can be made from
stainless steel screen (or other acceptable material such as
plastic) of about 3-4 mm mesh. The screen tray should be
supported above the bottom of the test chamber by two folds
26
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EG-11
August, 1982
oE screen or other devices which function as legs or supports.
The edges of the screen tray should be turned-up to prevent bump
spills and to prevent the embryos from rolling off in the event
of excessive turbulence. Suspending or supporting the screen
tray off the bottom ensures adequate water circulation around the
embryos and avoids contact of embryos with possible bottom
debris.
(iv) Test substance delivery system. (A) The choice of a
specific delivery system depends upon the specific properties and
requirements of the test substance. The apparatus used should
accurately and precisely deliver the appropriate amount of stock
solution and dilution water to the test chambers. The system
selected should be calibrated before each test. Calibration
includes determining the flow rate through each chamber, and the
proportion of stock solution to dilution water delivered to each
chamber. The general operation of the test substance delivery
system should be checked twice daily for normal operation
throughout the test. A minimum of five test substance
concentrations and one control should be used for each test.
(B) The proportional diluter and modified proportional
diluter systems and metering pump systems have proven suitable
and have received extensive use.
27
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EG-11
August, 1982
(C) Mixing chambers should be used between the diluter and
the test chamber(s). This may be a small container or flow-
splitting chamber to promote mixing of test substance stock
solution and dilution water, and is positioned between the
diluter and the test chambers for each concentration. If a
proportional diluter is used, separate delivery tubes should run
from the flow-splitting chamber to each replicate test chamber.
Daily checks on this latter system should be made.
(v) Other equipment required. (A) An apparatus for removing
undesirable organisms, particulate matter and air bubbles.
(B) An apparatus for aerating water.
(C) A suitable magnifying viewer for examination of minnow
embryos.
(D) A suitable apparatus for the precise measurement of
growth of the fish, including both length (e.g., with metric or
ruler caliper or photographic equipment) and weight.
(E) Facilities for providing a continuous supply of live
brine shrimp nauplii (Artemia salina).
(F) Facilities (or access to facilities) for performing the
required water chemistry analyses.
(vi) Cleaning of equipment. (A) Test substance delivery
systems and test chambers should be cleaned before use. Test
chambers should be cleaned during the test as needed to maintain
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EG-11
August, 1982
the dissolved oxygen concentration, and to prevent clogging of
the embryo cup screens and narrow flow passages.
(B) Debris can be removed with a rubber bulb and large
pipette or by siphoning with a glass tube attached to a flexible
hose. Debris should be run into a bucket light enough to observe
that no live fish are accidentally discarded.
(vii) Dilution water--(A) General. (JJ A constant supply
of acceptable dilution water should be available for use
throughout the test. Dilution water should be of a minimum
quality such that the test species selected will survive in it
for the duration of testing without showing signs of stress
(e.g., loss of pigmentation, disorientation, poor response to
external stimuli, excessive mucous secretion, lethargy, lack of
feeding or other unusual behavior). A better criterion for an
acceptable dilution water for tests on early life stages should
be such that the species selected for testing will survive, grow
and reproduce satisfactorily in it.
(2] The concentration of dissolved oxygen in the dilution
water (fresh or salt) should be between 90% and 100%
saturation. When necessary, dilution water should be aerated by
means of airstones, surface aerators, or screen tubes before the
introduction of the test substance.
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EG-11
August, 1982
(_3_) Water that is contaminated with undesirable microoganisms
(e.g. fish pathogens) should not be used. If such contamination
is suspected, the water should be passed through a properly
maintained ultraviolet sterilizer equipped with an intensity
meter before use. Efficacy of the sterilizer can be determined
by using standard plate count methods.
(B) Fres hwater. (1) Natural water (clean surface or ground
water) is preferred, however, dechlorinated tap water may be usea
as a last resort. Reconstituted freshwater is not recommended as
a practical dilution water for the early life stage toxicity test
because of the large volume of water required.
(2] Particulate and dissolved substance concentrations should
be measured at least twice a year and should meet the following
specif ic at ions :
Subs tance Concentration Maximum
Particulate matter < 20 mg/liter
Total organic carbon (TOG) < 2 mg/iter
Chemical oxygen demand (COD) < 5 mg/liter
Un-ionized ammonia < 1 ug/liter
Res idual chlorine < 1 ug/liter
Total organoposphorus pesticides < 50 ng/liter
Total organochlorine pesticides
plus polychlorinated biphenyls (PCBs) < 50 ng/liter
Total organic chlorine < 25 ng/liter
30
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cG-11
August, 1982
(3) During any one month, freshwater dilution water should
not vary more than 10% from the respective monthly averages of
hardness, alkalinity and specific conductance; the monthly pH
range should be less than 0.4 pH units.
(C) Saltwater. (_!_) Marine dilution water is considered to
be of constant quality if the minimum salinity is greater than 15
o/oo and the weekly range of the salinity is less than 15 o/oo,
The monthly range of pH should be less than 0.8 pH units.
Saltwater should be filtered to remove larval predators. A pore
size of <_ 2 0 micrometers (urn) is recommended.
(_2_) Artificial sea salts may be added to natural seawater
during periods of low salinity to maintain salinity above 15
o/oo.
(3) Test parameters (i) Dissolved oxygen concentration.
It is recommended that the dissolved oxygen concentration be
maintained between 90 and 100 percent saturation; but it should
be no less than 75 percent saturation at all times for both
minnow species and between 90 and 100 percent saturation for the
trout species in all test chambers. Dilution water in the head
box may be aerated, but the test solution itself should not be
aerated.
(ii) Loading and flow rate. (A) The loading in test
chambers should not exceed 0.1 grams of fish per liter of test
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EG-11
August, 1982
solution passing through the test chamber in 24 hours. The flow
rate to each chamber should be a minimum of 6 tank volumes per 24
hours. During a test, the flow rates should not vary more than
10 percent from any one test chamber to any other.
(B) A lower loading or higher flow rate or both should be
used if necessary to meet the following three criteria at all
times during the test in each chamber containing live test
organisms: (_1_) the concentration of dissolved oxygen should not
fall below 75 percent saturation for the fathead and sheepshead
minnows and 90 percent for the rainbow and brook trout; (2_) the
concentration of un-ionized ammonia should not exceed 1 ug/liter;
and (_3_) the concentration of toxicant should not be lowered
(i.e., caused by uptake by the test organisms and/or materials on
the sides and bottoms of the chambers) more than 20 percent of
the mean measured concentration.
(iii) Temperature. (A) The recommended test temperatures
are:
(_!_) Fathead minnow 25°C for all life stages.
(_2) Sheepshead minnow 30°C for all life stages.
(_3_) Rainbow and brook trout 10°C for embryos.
12°C for fry and alevins.
(B) The actual test temperature during the duration of the
test should remain within 1.5°C of the selected test
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EG-11
August, 1982
temperature. It is recommended that the test system be equipped
with an automatic alarm system to alert staff of instantaneous
temperature changes in excess of 2°C. If the water is heated
(i.e., for minnow species), precautions should be taken to ensure
that supersaturation of dissolved gases is avoided. Temperatures
should be recorded in all test chambers at the beginning of the
test and weekly thereafter. The temperature should be recorded
at least hourly in one test chamber throughout the test.
(xi) Light. (A) Brook and rainbow trout embryos should be
maintained in darkness or very low light intensity through one
week post-hatch, at which time a 14-hour light and 10-hour dark
photoperiod should be provided.
(B) For fathead and sheepshead minnows, a 16-hour light and
8-hour dark (or 12:12) photoperiod should be used throughout the
test period.
(C) A 15-minute to 30-minute transition period between light
and dark is optional.
(D) Light intensities ranging from 30 to 100 lumens at the
water surface should be provided; the intensity selected should
be duplicated as closely as possible for all test chambers.
(e) Reporting. A report of the results of an early life
stage toxicity test should include the following:
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BG-11
August, 1982
(1) Name of test, sponsor, investigator, laboratory, and
dates of test duration.
(2) Detailed description of the test substance including its
source, lot number, composition (identity and concentration of
major ingredients and major impurities), known physical and
chemical properties, and any carriers (solvents) or other
addi tives used.
(3) The source of the dilution water, its chemical
characteristics, and a description of any pretreatment.
(4) Detailed information about the test organisms including
scientific name and how verified and source history, observed
diseases, treatments, acclimation procedure, and concentration of
any contaminants and the method of measurement.
(5) A description of the experimental design and the test
chambers, the depth and volume of the solution in the chambers,
the way the test was begun, the number of organisms per
treatment, the number of replicates, the loading, the lighting, a
description of the test substance delivery system, and the flow
rate as volume additions per 24 hours.
(6) Detailed information on feeding of fish during the
toxicity test, including type of food used, its source, feeding
frequency and results of analysis (i.e., concentrations) for
contaminants.
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EG-11
August, 1982
(7) Number of embryos hatched, number of healthy embryos,
time to hatch, mortality of embryos and fry, measurements of
growth (weight and length), incidence of pathological or
histological effects and observations of other effects or
clinical signs, number of healthy fish at end of test.
(8) Number of organisms that died or showed an effect in the
control and the results of analysis for concentration(s) of any
contaminant in the control(s) should mortality occur.
(9) Methods used for, and the results of (with standard
deviation), all chemical analyses of water quality and test
substance concentration, including validation studies and reagent
blanks; the average and range of the test tempe rature (s ) .
(10) Anything unusual about the test, any deviation from
these procedures, and any other relevant information.
(11) A description of any abnormal effects and the number of
fish which were affected during each period between observations
in each chamber, and the average concentration of test substance
in each test chamber.
(12) Reference to the raw data location.
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ES-8
August, 1982
TECHNICAL SUPPORT DOCUMENT
FOR
FISH EARLY LIFE STAGS TOXICITY TEST
OFFICE OF TOXIC SUBSTANCES
OFFICE OF PESTICIDES AND TOXIC SUBSTANCES
U.S. ENVIRONMENTAL PROTECTION AGENCY
WASHINGTON, D.C. 20460
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TABLE OF CONTENTS
Subject Page
I. Purpose 1
II. Scientific Aspects 1
General 1
Test Procedures 4
General 4
Experimental Design 5
General 5
Controls 8
Carriers 9
Beginning the Test 11
Observations and Measurements 13
Acceptability Criteria 14
Test Results and Analysis 15
General 15
Analys is 17
Test Conditions 19
Tes t Species 19
Selection 19
Sources 25
Maintenance of Test Species 27
Acclimation 27
Feeding 23
Facilities 29
General 29
Construction Materials 30
Tes t Ch ambe rs 31
Embryo Incubation Apparatus 32
Test Substance Delivery System 33
Cleaning 36
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Dilution Water 37
Subject Page
Environmental Conditions 41
Dissolved Oxygen 41
Flow Rates 43
Loading 44
Temperature 45
Light 47
Reporting 49
III. Economic Aspects 50
IV. References 51
11
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Office of Toxic Substances ES-8
August, 1982
TECHNICAL SUPPORT DOCUMENT FOR FISH
EARLY LIFE STAGE TOXICITY TEST
I. Purpose
The purpose of this document is to provide the
scientific background and rationale used in the development
of Test Guideline EG-11 which uses several species of fish
to evaluate the effects of chemical substances on the early
life stages of fish. The Document provides an account of
the scientific evidence and an explanation of the logic used
in the selection of the test methodology, procedures and
conditions prescribed in the Test Guideline. Technical
issues and practical considerations are discussed. In
addition, estimates of the cost of conducting the test are
provided.
of-the-science developments.
II. Scientific Aspects
A. General
Toxicity tests with the early life stages of fish
provide an extremely useful tool for the assessment of a
chemical's potential to produce adverse environmental
effects to aquatic organisms. An early life stage toxicity
test is the logical sequel to an acute toxicity test,
particularly where a high degree of toxicity has been
demonstrated, or where continuous exposure is expected.
Acute toxicity tests primarily measure mortality during a
short duration (i.e. 96 hours) of exposure. The early life
stage toxicity test is conducted to evaluate and estimate
lethal and sublethal effects of low concentrations of a
chemical or mixture over a longer period of exposure (i.e.
32 days or more), and during sensitive and critical stages
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August, 1982
of fish growth and development.
The early life stage toxicity test can be used to assess
the hazard potential to certain freshwater and estuarine
fish resulting from chronic exposure to a chemical substance
or mixture. The test procedures apply to nearly all types
of chemicals that can be measured in freshwater and
saltwater, and with slight modifications can be useful for
testing other potentially toxic agents (e.g. oils,
particulate matter, etc.).
It is generally recognized that fish are resources that
are valuable ecologically, economically, recreationally and
aesthetically. Healthy aquatic environments support a
diversity of fish populations. A reduction in the quality
of an aquatic environment will be indicated by effects upon
the communities contained therein. Fish are considered for
use in toxicity tests because they are sensitive indicators
of chemical pollution, and are important, integral parts of
aquatic communities and can be used as surrogates for other
species in comparative toxicology.
The early life stage toxicity test can produce results
which permit estimates of chronic toxicity normally derived
from longer term, full life-cycle toxicity studies (Macek
and Sleight 1977, McKim 1977). This conclusion was
reflected by several investigators over the past decade who
proposed the use of the early life stage toxicity test to
predict chronic toxicity (Pickering and Thatcher 1970,
Pickering and Cast 1972, McKim et al. 1975, Eaton et al.
1973, Spehar et al. 1977). McKim (1977) analyzed data from
56 life-cycle toxicity tests completed during the last
decade with 34 organic and inorganic chemicals and four
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ES-3
August, L982
species of fish, and concluded that the embryo-larval and
early juvenile life stages were the most, or among the most,
sensitive in their responses to chemical insult. Macek and
Sleight (1977) concluded after a review of much of the
available data that critical life stages (embryos and
developing fry) exposed to toxicants provide estimates of
chronically safe concentrations remarkably similar to those
empirically derived from definitive chronic toxicity
studies. In addition, it was stated that toxicity tests
with critical life stages can provide information on
potential long-term effects in situations where acute
toxicity (i.e. 96-hr. LC50) is not observed.
Macek and Sleight (1977), stated that the concentration
for most toxicants which will not be acutely toxic to the
most sensitive life stages is generally the chronically safe
concentration for fish. Carlson (1971) stated that several
studies (i.e. Mount and Stephan 1967, 1969, Mount 1968,
Brungs 1969, Eaton 1970, Pickering and Thatcher 1970, McKim
and Benoit 1971) have shown significant differences between
acute and long-term toxicity values. Allison and Hermanutz
(1977) stated that freshwater fish populations could be
directly damaged by prolonged exposure to a chemical at
concentrations several thousand times lower than the
concentration which causes acute mortality. The hazard of
Kepone, for example, is greatly underestimated by acute
toxicity tests (Hansen et al. 1977). It is obvious that
assessment of the hazard potential of a chemical substance
would be more accurate when using toxicity values derived
from studies using sensitive, early life stages of fish and
addressing chronic intoxication (Pickering 1974), rather
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August, 1982
than the use of toxicity values derived solely from acute
toxicity studies (i.e. 96-hour LC50 values). McKim (1977)
also acknowledged the sensitivity of the early life stage
toxicity test and proposed its use for screening large
numbers of chemicals rapidly at a far lower cost per test
than with a partial or complete life-cycle toxicity test.
B. Tes t Procedures
1. General
The basic concept behind the early life stage toxicity
test is to estimate chronic toxicity through the use of a
relatively short and inexpensive test. The literature
reviews of McKim (1977) and Macek and Sleight (1977) are the
most comprehensive evaluations of the early life stage
toxicity test for use in evaluating chronic toxicity of
chemicals to fishes.
The current experimental design incorporated into the
early life stage toxicity test guidelines is derived from
the various methods which have evolved historically, and
reflect current state-of-the-science improvements. The
first specific methods for use by government research and
regulatory personnel were proposed in 1971 by the Committee
on Aquatic Bioassays of the US Environmental Protection
Agency and revised in 1972. The Bioassay Committee proposed
a "Recommended Bioassay Procedure for Eggs and Fry Stages of
Freshwater Fish." Another method by the Bioassay Committee
was titled: "Recommended bioassay procedure for fathead
minnow Pimephales promelas Rafinesque Chronic Tests," (USEPA
1972a). The Environmental Research Laboratories at Duluth
(MN), Corvallis (OR) and Gulf Breeze (FlA) continue to play
a major role in methodology development for aquatic toxicity
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August, 1982
tests and have supplied valuable information in the
development of the early life stage test guidelines.
Investigative work reported in the literature at large has
built considerably upon the early bioassay committee
methods. In selecting the most practical and reliable test
methods and procedures £or the early life stage toxicity
test guidelines, the available literature was reviewed to
search for improvements or recommendations to the early
committee methods. In addition, the American Society for
Testing and Materials (ASTM) has been extremely active in
developing a standard method for conducting early life stage
toxicity tests. Many of the most prominent scientists in
the field of aquatic toxicology are ASTM members with
extensive practical experience in conducting early life
stage toxicity tests. EPA has reviewed and considered the
working drafts generated by the ASTM Task Group involved in
the development of a standard practice for conducting
toxicity tests with early life stages of fishes. The Agency
has also solicited methods for conducting early life stage
toxicity tests from several established testing laboratories
for consideration in selection of the test standard
methodology, as well as to ascertain practical contraints
and deviations (in design and method) between commercial
facilities (e.g. ABC protocols #7809 and #7810, EG & G
Bionomics method 1979, and Union Carbide Environmental
Services, March 1979).
2. Experimental Design
a. General
Many test design requirements were established upon the
preponderance of available information supporting specific
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ES-8
August, 1982
methods or conditions. Occasionally the data available
could not support selection of a specific requirement, and
such requirements were selected following careful
deliberation in order to minimize the number of possible
variables which could affect the outcome or reproducibility
of results. While lacking empirical support, such decisions
provide for consistency in the test method which in turn
enables comparison of test results with a higher degree of
confidence. Selection of a testing requirement should
always take into consideration the optimum conditions and
requirements for the test species, based upon available
information and research.
Early life stage exposure is defined by Macek and
Sleight (1977) as exposure of a test species to a test
substance during most, preferably all, of the em'oryogenic
period (incubation of the fertilized eggs) and exposure of
fry (as newly hatched fry through young juveniles) for a
period of 30 days after hatching for warm water fish with
em'oryogenic periods of 1-14 days (fathead and s heaps he ad
minnows), and 60 days exposure after hatching for fish with
longer embryogenic periods (i.e. brook and rainbow trout).
The TSCA early life stage toxicity test guideline is
initiated with newly-fertilized embryos (less than 48-hours
old for the minnow species and less than 96-hours for the
trout species). An attempt was made to ensure uniformity of
exposure periods, and to permit the shipment of embryos to a
testing facility which may lack in-house cultures of the
test species. The 48-hour requirement ensures that warm
water species (i.e. fathead and sheepshead minnows) will
receive a minimum amount of exposure as embryos prior to
hatch ing .
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August, 1932
A minimum o£ five duplicated exposure concentrations of
a test substance and at least one control should oe used in
conducting an early life stage toxicity test. This provides
a minimum number of data responses to ensure statistical
validity, as well as provide the minimum amount of responses
to adequately develop the quantitative data necessary to
establish biological conclusions and significance. The
quantitative data derived from an early life stage toxicity
test is used to determine the lowest toxicant concentrations
which adversely effect the test fish and those which
apparently do not. Five test concentrations with a minimum
of one control have been used exteas ively and are
statistically valid and historically acceptable. The
preponderance of data developed in the field of aquatic
toxicology has been generated using five concentrations witn
a minimum of one control.
The use of 5 test substance concentrations is designed
to produce effects at the higher concentrations, while
producing no observed effects in the lowest concentrations,
with the mid-range concentration(s) contributing to the
degree of certainty in the definition of the concentration-
response curve (Doudoroff et al. 1951). It is obvious,
however, that the desired responses of effect and no-effect
are dependent upon how accurately the test substance
concentrations are selected. Selection of the test
substance concentrations for use in the early life stage
toxicity test is considered on a case by case basis. A
flow-through toxicity test at least 96 hours long using the
larval or juvenile stages of the fish species to be tested
should be performed. The highest concentration for the
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ES-8
August, 1982
early life stage test should be similar to the lowest
concentration in the acute test causing a reduction in
survival. The dilution factor recommended over the past
decade has been primarily 0.5 between treatments. No hard
and fast rule is possible for selecting the proper test
concentrations. Caution should be exercised, because
selection of the wrong concentrations and dilution factor
can negate the acceptability of the test for its intended
purpose by EPA. For example, if all treatment
concentrations elicit a significant effect, the test
probably should have been conducted at lower concentrations;
conversely, if no significant effect occurs, higher
concentrations probably should have been used.
The location of the test chambers and the introduction
of test organisms into the test chambers are randomized to
prevent biases from being introduced, and consequently
affecting the objectivity necessary in the test. Biases can
occur in environmental conditions (temperature, lighting,
disturbances, etc.), embryo selection and distribution,
diluter system function, etc.. Both the test organisms and
test concentrations should be assigned to the exposure
chambers by formal randomization, in order to avoid
introduced biases.
b. Controls
Controls are required for every test to assure that any
effects which are observed are due to the test substance and
not to other factors. Similarly, the carrier (solvent for
test substance) control demonstrates any stress induced by
the carrier, apart from the test substance. Controls are
universally required in toxicological studies and contribute
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August, 1982
an essential value to the interpretation of test results, as
effects observed in the test organisms exposed in test
solutions can be attributed to the test substance only when
evaluated in light of results from the controls.
c_. Carriers
Carriers can affect (i.e. stress) test organisms and can
possibly alter the form of the test substance in water. For
these reasons the preferred standard carrier for a test
substance is water, and the use of other carriers should be
avoided unless absolutely necessary to get the test
substance into water. A safety margin between the acute
toxicity of the carriers and any possible subacute effects
that might influence the experimental outcome is
necessary. Triethylene glycol (TEC) and dimethyl formamide
(DMF) appear to exert the least influence on test organisms
and test substances of the several carriers that have been
tested. Work performed by Rick Cardwell (unpublished data)
indicates the following 96-hour, LC50 values (mg/1):
Fathead minnow Brook trout
TEG
(Triethylene glycol) - 92,500 73,500
DMF
(Dimethyl Formamide) - 10,410 8,366
Acetone 9,100 6,070
EG&G Bionomics (1979) state in their method for
conducting an early life stage test with fathead minnows
that the carrier concentration should not exceed one-one
thousandth (1/1000) of the 96-hour LC50 for the species and
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August, 1982
solvent. This would equate to 92 mg/1 and 73 mg/1 of TEG
for the fathead minnow and brook trout respectively, based
upon the above information.
Due to the low toxicity to aquatic animals, low
volatility and a high ability to dissolve many organic
compounds, dimethyl formamide and triethyleae glycol are the
preferred organic carriers for preparing stock solutions,
but methanol, acetone, and ethanol may also be used. In
order to determine the influence or possible effects caused
by the carrier, a carrier control containing dilution water
and the carrier at the highest concentration used in the
toxicity test should be included and conducted
simultaneously with the toxicity test.
Responses of control and carrier control fish provide a
measure of the acceptability of the test, and it is for this
reason and for use in interpreting the test results, that
the controls are required. If a carrier is used, criteria
for effects (e.g. embryo mortality, larva mortality, growth)
used in the comparison of control and exposed animals should
also be applied to the animals in the control and control
with carrier chambers. If a significant difference exists
in these criteria, the test may not be considered
acceptable, as the data would inaccurately reflect the test
substance toxicity, and thus the overall test results would
be ambiguous. The overall survival of the control animals
should be at least 80%; otherwise the test should be
considered unreliable, because the effects which are
observed cannot be attributed with confidence to the test
substance alone.
10
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August, 1932
3. Beginning the Test
It is important that the quality of the dilution water
and the test substance concentration be constant at the time
that the test is started, and for the duration of the
test. A 48-hour observation period is required prior to
beginning the test in order to ascertain that all equipment
(e.g. temperature control, test substance delivery system,
etc.) is functioning properly and to allow an equilibrium to
be reached in the test system. An equilibrium is achieved
when the test dilution water quality (e.g. dissolved oxygen,
temperature, pH, etc.) is constant, and all phys icochemical
reactions, such as leaching of contaminants and adsorption
of the test substance within the test system are
stabilized. Fluctuation of the test substance concentration
can be caused by adsorption of the test substance by the
test system or by a malfunction in the test substance
delivery system. The 48-hour observation period permits
fine adjustments to the test substance delivery system and
allows for any adsorption which may initially occur.
Consistent test substance concentrations are
critical for valid test results and should be accurately
evaluated and maintained (Lloyd 1978).
Carrnignani and Bennett (1976) demonstrated leaching of
plastics from some closed aquaculture systems, and
recommended that new aquaculture systems using plastics
should be flushed for at least 10 days with fresh water at
temperatures at least 3°C above the operating temperature of
the system before they are put into service. Established
systems should only require flushing for 48 hours to achieve
an equilibrium.
11
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August, 1982
After final calibrations of the test substance delivery
s ys tern and verification of the actual test substance
concentrations in the test water from each chamber, the test
can be initiated with acclimated, normal appearing
embryos. Since one of the measured parameters is hatching
success, use of non-viable embryos would skew the test
results and the data interpretation. In order to keep the
embryos viable an exchange of water through the embryo cups
should be maintained. This ensures a flow of oxygenated
water over the incubating embryos and dislodges particles
and debris which can encourage undesirable fungal growth.
Static conditions contribute to fungal growth, thus
maintaining water flow over incubating eggs has been
standard practice since the earliest chronic studies «?i th
the fathead minnow. Mount (1968) used a rocker arm
apparatus driven by a low speed (>lrpm <3rpm) electric
motor, rfhile other investigators (e.g. Hansen et al . 1977)
have elected to use a self-starting siphon to vary the water
level in the test chambers.
Each day until hatching the embryos are usually
inspected without the use of a scope or magnifying viewer
for the purpose of discarding obviously dead embryos. This
is done in order to prevent fungus, which attacks dead
embryos, from spreading to the surrounding live embryos.
The use of a scope or viewer could unduly stress the
developing embryos because of tne amount of handling
involved and is therefore discouraged. Handling of newly
hatched larvae/fry is also discouraged in order to minimize
stress and avoid injury.
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August, 1982
Transfer of fry from embryo cups into the test chambers
should not involve the use of nets; rather the fry should be
allowed to swim out. This reduces the amount of stress and
injury that could result from directly handling the fry.
Trout incubated on screen trays will hatch out into test
chambers and thus transfer is not necessary.
Daily records should be kept of the number of dead
embryos and fry, and in addition, the number of unaccounted
for or deformed embryos. Daily records are also made for
any lethargic or glossly abnormal fry. This includes
swimming behavior and physical appearance. The range of
time-to-hatch (to the nearest day) for each replicate should
also be recorded. These and other observations are detailed
in Section 2.1.4: Observations and Measurements.
The number of embryos required to begin the test is
flexible and varies upon the statistical power desired and
the test species selected. Minimal standards of control
response (e.g. percentage of embryos to hatch, percentage
larval survival,and percentage overall survival) for each
species dictates the number of embryos used.
Test guidelines for conducting early life stage toxicity
tests specify only the minimum number of embryos required to
enable the use of sensitive statistical analyses of the
biological data.
4. Observations and Measurements
A final aspect of the test procedure pertains to the
measurements and observations which should be made during
the test. Daily observations are made to ascertain the
functioning of the test equipment and to ensure the required
test conditions are maintained (i.e. temperature, dissolved
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oxygen, test substance concentrations, dilution water
quality, etc.). Biological data responses are gathered
through observations of mortality for embryos and fry, and
for abnormal physical appearances and behavior. The
biological responses are used in combination with physical
and chemical data (i.e. test conditions) in evaluating the
overall lethal and sublethal effects observed. The
observations required during the test are the minimum
intervals necessary to produce valid and interpretable data.
Generally, the most important data derived from
observations in tests with the early life stages of fish are
those that show significant differences between control
treatments and experimental treatments based on the
following: percent survival of embryos to hatching, time to
hatching, survival of hatched fish to termination of test,
time to swim-up for trout, survival from beginning to
termination of test, and growth and development of young
fish.
5. Acceptability Criteria
The acceptability criteria specified in the test
guidelines are essential to the interpretation of the test
results. The criteria are used to minimize and detect
possible influences on the test results other than those
produced by the test substance. The acceptability criteria
also delineate the function of test conditions and
parameters, and are based upon what has been normally
achieved and observed in the preponderance of relevant
studies reported in the literature.
Test acceptability is basically dependent upon the test
results being primarily a function of the test substance,
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and not due to the influence of external variables. For
example, since various measured parameters of the test
species (e.g. weight, length) are compared with the
controls, it is important that controls are "normal" as
determined historically. When control survival values are
less than those typically observed in past tests, it can be
assumed that there are problems with some aspect of the test
(e.g. fluctuations in environmental parameters, stressed
test organisms, etc.). Thus, test acceptability is
dependent upon the criteria specified in the test guidelines
which are designed to allow a determination of statistical
validity, in addition to providing quantitative data
necessary to establish biological conclusions and
significance. Variation from the specified criteria
indicates that other aspects of the test may be affected and
final results can not be attributed entirely to the test
substance. Thus, the usefulness of the test in evaluating
the hazard potential of a test substance is of questionable
value unless the acceptability criteria are satisfied.
6. Test Results and Analysis
a. General
In conducting toxicity tests with the early life stages
of fish, data are obtained to determine the effects of the
test substance on a particular test organism during a
sensitive period of its growth and development. As in all
aquatic toxicity tests, statistical hypotheses are developed
and the experiments are designed to test these hypotheses.
Experimental data are gathered, analyzed and statistical
conclusions are drawn. It should be remembered, however,
that statistical hypotheses and subsequent conclusions are
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August, 1982
measurements of broader biological hypotheses and biological
responses and interpretations. Hence, a statistically
significant conclusion based upon a preselected significance
level does not necessarily signify a similar or equivalent
biological significance. The biological significance and
interpretation of a statistically derived conclusion should
be based upon the experience and professional judgment of
the investigator or reviewer.
Toxicity tests with fish early life stages are generally
used to determine the lowest test substance concentration^)
which adversely affect the test species, and those which
apparently do not.
These are usually defined as:
Lowest Observed Effect Concentration - The lowest test
concentration from a valid toxicity test for which the
null hypothesis of no difference (i.e. control and
treatment means were similar) was rejected (P<.05) for
any specified response.
No Observed Effect Concentration - The highest test
concentration from a valid toxicity test for which
insufficient evidence exists to reject the null
hypothesis of no difference (P< 0.05) for any specified
r es po ns e .
The proposed early life stage toxicity test guidelines
are specific only where minimum parameters and conditions
are recognized to be essential for the production of valid
results. The flexibility of the test guidelines allows for
the wide variations among commercial testing facililties.
For example, the number of tanks and fry chambers or embryo
cups is not mandatory. While the majority of facilities
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August, 1932
utilize the minimum number of test substance concentrations
(i.e. 5 plus controls), there is little uniformity in the
number of embryos used to initiate a test (e.g. the typical
range is from 850 to 2800). Each test can produce valid and
acceptable results, yet specific guidance for the analysis
of data derived is difficult. Each different combination of
fry chambers, embryo cups, etc., would require a different
design, and therefore would require a separate analysis.
b. Analysis
Various methods for the analysis of data are available
and acceptable. The following discussion is an example of
an acceptable approach. The key to selecting an appropriate
experimental design and accompanying statistical analyses
lies in the identification of the smallest division of
experimental material such that any two units may receive
different treatments in the actual experiment (Cox 1958).
Data from the toxicity tests are usually either continuous
(e.g. length or weight measurements) or dichotomous (e.g.
number hatching or surviving) in nature. In general,
continuous data should be analyzed using an appropriate
analysis of variance (ANOVA) technique followed by an
appropriate multiple comparison test. Dichotomous data
should be analyzed using a form of 2x2 contingency table.
Several reference sources are available to determine
which ANOVA is the most appropriate (Sokal and Rohlf 1969,
Box et al. 1978). If solvent and dilution water controls
are used in the toxicity test, they should be evaluated for
possible combining. This should be done using a "T-test"
with a large probability level (e.g. P< 0.25). If the null
hypothesis is rejected, the responses in both controls
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August, 1982
should be evaluated and the most appropriate control used in
all further data analyses. As a part of the ANOVA it is
desirable to plot the residual (error) versus concentration,
to determine whether there have been any obvious violations
of the assumption of normality. The following should be
presented when reporting the results from an ANOVA
technique: (a) the ANOVA model; (b) the ANOVA table; (c) the
F statistic; and (d) the level of significance for the F
statistic. It is also appropriate to report any residual
plots generated. The ANOVA is only the first step; assuming
the ANOVA F test was sigif leant (P< 0.05), a comparison is
made of all non-zero concentration responses with tne
control response. The appropriate procedure for this is
normally a multiple comparison test such as Dunnett's
procedure (Dunnett 1955, 1964). Dunnett's procedure is
commonly used and is analagous to performing +/ - tests on
the control with each concentration tested. A second
technique is William's procedure (Williams 1971, 1972).
This procedure also compares the control with each test
concentration, but wi th the assumption that the true mean
response decreases (or increases for certain types of
response) with increasing concentrations. If this
assumption is correct, then William's procedure is the more
powerful statistical technique. An excellent discussion of
these techniques and other multiple comparison procedures is
provided by Chew (1977).
Presentation of results from these tests should include
the critical value of the statistic and its corresponding
level for each comparison of the test statistics.
Dichotomous data should be analyzed using a 2x2 contingency
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August, 1982
table. When attempting to distinguish between a control
response and the same response (e.g. hatchabil i ty) at a low
test substance concentration level, the calculated expected
cell frequencies are likely to be low. Therefore, it is
appropriate to analyze these data using Fisher's exact test
(Sokal and Rohlf 1969, Fisher 1958) rather than with an
approximation test like the Chi-square test. If two control
treatments (e.g. regular and carrier) are used in the
toxicity test, they should be evalulated using a 2x2
contingency table rather than the T-test.
C. Test Conditions
1. Test Species
a. Selection
Four species of fish are proposed for use in conducting
early life stage toxicity tests. These species are:
1) Fathead Minnow (Pimephales promelas Rafinesque)
2) S'neepshead Minnow (Cyprinodon variegatus)
3) Brook Trout ( Salvelinus fontinal is_ Mitchill)
4) Rainbow Trout (Salmo gairdneri)
These species are representative of warm and cold
freshwater and estuarine fish respectively.
There are several criteria to be considered when
selecting a fish species for use in an early life stage
toxicity test:
1) Embryos are required to initiate the early life
stage toxicity test; therefore recently fertilized eggs
or fresh sperm and ova should be readily available. As
such, it is desirable that the species be amenable to
laboratory conditions, demonstrating high survival in
capt ivity.
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August, 1982
2) The species should possess a demonstrated ability to
reproduce readily in close confinement, producing large
numbers of eggs, or they should readily available from
reliable commercial hatcheries.
3) The species should mature rapidly, yet be small
enough in size to enable the use of large (statistically
valid) numbers during the toxicity tests.
4) The species should be relatively widely distributed,
with non-polluted areas included in its geographical
range .
5) The species should be relatively sensitive to toxic
pollutants, and its susceptability to toxic pollutants
should equal or exceed that of other fish species in its
geographical range.
The four species selected for use in conducting early
life stage toxicity tests generally satisfy these criteria,
and can be used individually to address practical problems
which may be restricted to a specific geographical
location. Each species has been used successfully in early
life stage toxicity tests, and in fact, have been used more
often than any other fish species for such test purposes.
In addition, these species have also demonstrated success in
both acute and full chronic toxicity tests. The use of the
same species for many aquatic toxicity tests provides for a
good comparative toxicological data base. Additional
support for the selection of these particular species is
provided in the following discussions.
Fathead Minnow (Pimephales promelas Rafinesque)
The fathead minnow is an important forage fish (Scott
and Grossman 1973), which has significant value as a
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August, L932
converter of algae, organic detritus from bottom deposits
and planktonic organisms. It is widely distributed, small,
highly prolific and has a prolonged spawning period,
assuring availability of recently hatched and juvenile
fatheads to a large variety of predators (Scott and Grossman
1973). A list of predators for the fathead minnow would
require a listing of all of the carnivorous fisn and fish-
eating birds associated with it throughout its geographic
range (Scott and Grossman 1973). Its role as an important
forage species makes it an essential link in the food
economy of some natural ecosystems and its value in this
role is inestimable (Scott and Grossman 1973).
The fathead minnow is distributed widely throughout the
Great Plains region of Canada and the United States, as well
as much of the region east of the Great Plains from the
southern drainage of the Hudson Bay and the Maritime
Provinces of Canada southward through the Ohio and
Cumberland systems to the Tennessee River Basin. Although
apparently absent on the Atlantic slope and the Gulf states
east of the Mississippi River, the fathead is present as far
west as New Mexico and Chihuahua, Mexico in the South. It
has also become established in California (Blair et al .
1968, Scott and Grossman 1973).
The fathead minnow has been established by many
laboratories as an assay fish for the determination of the
toxicity of aquatic pollutants (Manner and DeVJese 1974). It
is considered a good bioassay animal (Mount 1973) and has
been used successfully in many bioassays (Martin 1973, Me Kim
1977). It has a demonstrated ease of spawning and handling,
reaching sexual maturity in five months under tne proper
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August, 1982
conditions (Adelman and Smith 1976). The time to hatching
depends on temperature, but the average is 4.5 - 6 days; at
25°C, 5 days is normal (Scott and Grossman 1973). Newly
hatched fry are about 5 mm long, and reach an average
length of 50-70 mm as adults (Scott and Grossman 1973). The
early embryology of the fathead minnow has been studied by
Manner and DeWese (1974), who also indicated the extensive
use of the fathead in toxicity studies. McKim (1977)
considered the fathead minnow an example of a consistently
sensitive species for use in toxicity studies. These
attributes and characteristics in combination with the
proven ability to easily transport embryos and fry, make the
fathead minnow an appropriate selection for use in early
life stage toxicity tests.
Sheepshead Minnow (Cypr inodon variegatus)
The sheepshead minnow is an omnivorous kiliifish (Family
Cypr inodont idae) which occurs naturally in estuaries from
Cape Cod, Massachusetts to Yucatan, Mexico and in the West
Indies to northern South America (Hildebrand 1917, Blair et
al. 1968). The sheepshead minnow inhabits tidal ponds,
sloughs, saltwater creeks, bayous and bay shores (Simpson
and Gunter 1956) and while prefering estuarine environments
has also been observed in freshwater streams and in strictly
saltwater (Hildebrand 1917). The sheepshead minnow is
considered an ecological dominant in its environment, and is
very hardy, pugnacious, and tolerant of salinity fluctuation
(Simpson and Gunter 1956). It is a very prolific species
and its fecundity helps to explain its great abundance
(Hildebrand 1917, Simpson and Gunter 1956). It has value as
a converter of detritus and vegetable matter and in
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August, 1982
controlling mosquito larvae, but its greatest value probably
lies in the food source it provides for larger fishes and
other predators. These include: spotted seatrout
(Cynoscion nebulosus), yellow bass (Micropogon undulatus ) ,
red drum (Sciaenops ocellata), black drum (Pogonias cromis)
and many others (Gunter 1945, Darnell 1958).
Hildebrand (1917) stated the spawning period for the
oviparous sheepshead minnow was cons ideraole, ranging from
March until October. The incubation period at laboratory
temperatures ranges from 5 to 6 days (Hildebrand 1917), with
5 days at 30°C being normal (Schimmel and Hansen 1974). The
sheepshead minnow spawns readily in laboratory aquaria, is
fecund and has a short life cycle and survives well under
laboratory conditions (Schimmel and Hansen 1974, Hansen and
Parrish 1977). The combination of its size and hardiness
make it a useful fish for laboratory toxicity tests
(Schimmel et al. 1974, Schimmel and Hansen 1974, Hansen and
Parrish 1977, Parrish et al. 1977). It should be noted,
however, that several of these authors have indicated that
the sheepshead minnow is relatively insensitive and that its
reproductive strategy differs from most marine species, both
are drawbacks which make the sheepshead minnow less than
ideal. However, no other marine or estuarine species has a
standardized method available for conducting early life
3 tage tests .
Brook Trout (Salvelinus fontinalis Mitchill)
The brook trout is cons idered an impo>rtant res ident of
many lakes, streams and ponds across the US, as both a
native species and introduced species (Thatcher et al. 1976,
Larson et al. 1977). Under natural conditions this North
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August, 1982
American species occurs along the northeastern North
American Seaboard south to Cape Cod, in the Appalachian
Mountains southward to Georgia, west in the upper
Mississippi and Great Lakes drainages to Minnesota, north to
Hudson Bay (Scott and Grossman 1973). In Canada the brook
trout is widely distributed throughout the Maritime
Provinces, and it has been introduced widely and often
successfully into many parts of the world because of its
appeal as a sport fish (Scott and Grossman 1973). Included
in its introduced range are the higher mountainous parts of
Western North America. These authors indicated that brook
trout occur in clear, cool, well oxygenated streams and
lakes and that a long interest in brook trout as a
hatchery-reared and pond-cultured sport fish has resulted in
an accumulation of information on the species. Thatcher et
al. (1976) stated that its life cycle and laooratory culture
is well documented.
Brook trout are carnivorous and feed upon a very wide
range of organisms, including worms, leeches, crustaceans,
aquatic insects, terrestrial insects, and a number of fish
species (Scott and Grossman 1973). These authors indicate
that the most serious predators are fish-eating birds (i.e.
kingfishers and mergansers) and larger fish such as the rock
b as s .
The brook trout eggs are large, 3.5-5.0 mm in diameter,
and incubation time is dependent upon such factors as
temperature and oxygen; at 41°F (5°C) eggs hatch in about
100 days, at 43°F (6.1°C) in about 75 days, and at 50°F
(10°C) in 50 days (Scott and Grossman 1973). The long
incubation period of this species necessitates that the
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August, 1982
duration of the early life stage life stage toxicity test be
correspondingly longer in comparison to the early life stage
toxicity tests conducted with the warm water minnow species.
Rainbow Trout (Salmo gairdneri Richardson)
The rainbow trout is the most common salmon id used for
bioassay purposes, is the easiest to rear under laboratory
and commercial hatchery conditions, and is easy to spawn
artificially to obtain eggs for testing. Brood stock can be
retained for several years with good success. The eggs are
easily transported from commercial hatcheries, with good
hatching success and survival of young. The rainbow is
native to the West Coast and Rocky Mountains of North
Auerica and is now widespread over most of the United
States. Several early life stage toxicity tests have been
completed with the rainbow trout with very good success.
The relatively rapid embryo and larval growth, along with
its relative resistance to disease compared to other
salmonids, makes it an ideal cold water fish for testing
toxic materials.
b. Sources
It is preferable to maintain in-house brood stock for
obtaining sufficient quantities of test embryos of known
history, quality and age; however, eggs and sperm or newly
fertilized embryos can be obtained from a reliable external
commercial or research facility. This is necessary to
prevent restricting the number of testing facilities to only
those which possess in-house cultures.
Fathead minnow embryos are generally obtained through
natural spawning under laboratory conditions. An artifical
spawning substrate for use in culturing fathead minnows is
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August, 1982
described by Benoit and Carlson (1977). The Duluth
Environmental Research Laboratory has maintained fathead
minnow culture stock successfully for many years.
Sheepshead minnow embryos can be obtained through
natural spawning or artifically through hormone injections
and stripping eggs and excising male testes. These methods
are discussed in Bioassay Procedures for the Ocean Disposal
Permit Program (EPA 600/9-78-010) and by the following
investigators: Schimmel and Hansen (1974) and Hansen and
Parrish (1977).
Rainbow and brook trout embryos can be obtained through
natural spawning (Hokanson et al. 1973, USEPA 1972D) or by
dry artificial methods. Dry artificial techniques are
valuable because brood stock can be artificially stripped
and returned to their ponds or chambers without harm. The
eggs can be immediately fertilized, left for an hour or two
to water harden, and then be transported to the
laboratory. Eggs obtained from dry artificial techniques
can be gathered in quantity, fertilized at the same time and
shipped chilled or iced down with no adverse effect on
viability. The eggs become increasingly sensitive
approximately four days post-fertilization, and should not
be moved between day 5 and day 14. The eggs can be moved
carefully after the eyes become visible, as the sensitive
embryo period occurs prior to day 14. Therefore, testing
should start with eggs which have been fertilized (embryos)
and are less than 4 days old. Natural spawning is less
desirable as it is difficult to obtain a sufficient quantity
of embryos necessary to conduct the toxicity test which are
uniformily less than 96-hours old.
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August, 1932
2. Maintenance of Test Species
a. Acclimation
The health and quality of the test organism is all-
important in any toxicological investigation (Hunn et al.
1958, Brauhn and Schoettger 1975), and acclimation is of
direct importance for correctness of bioassay results
(Sprague 1969). Acclimation of embryos to the test dilution
water and temperature is necessary to minimize or avoid
stress at the initiation of the toxicity test. Results of
tests with stressed organisms will not accurately represent
the response of the test species to the test substance
alone.
In-house cultures for the production of a test species
(i.e. fathead minnow) should be maintained at or very near
the actual test temperature and should utilize test dilution
water. In such cases acclimation is easily accomplished,
and stress producing factors are minimized.
Embryos received from external sources should be
acclimated to the test temperature and dilution water. This
should be accomplished quickly for transported fathead and
sheepshead minnows, because generally they will be received
by the testing facility at close to 48 hours of age and will
commence hatching within a day or two. Acclimation of
rainbow and brook trout embryos may require a longer period
of time, as they are generally transported with ice and
rapid temperature fluctuations should be avoided (Brungs and
Jones 1977). However, the longer incubation period of both
trout species makes the acclimation period prior to the
initiation of the test less critical.
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August, 1982
b. Feeding
The time to first feeding and the amount and type of
food is dependent upon the test species. Feeding of fathead
and sheepshead minnows should coramense when newly hatched
larvae are transferred from embryo cups into the test
chambers. This is because some of tne larvae will be 48
hours old at that time, and feeding is necessary to provide
nourishment for the young fish which were sustained
previously by their egg-sacs. Schimmel and Hansen (1974)
noted that sheepshead minnows accept live brine shrimp
nauplii 48 hours after hatching. The practice of feeding
fathead minnow fry at 43-hours was included in the early EPA
methods for partial and full chronic bioassays (USEPA
1972a,b), and is practiced by the majority of investigators
using both minnow species for early life stage toxicity
tests. Investigators have also supplemented a wide variety
of additional foods such as daphnia, trout pellets, dry
flake and frozen commercial preparations after the first
week. Mehrle et al. (1977) demonstrated the effect and
importance of diet quality on the outcome of toxicologicdl
research. In order to minimize the effect of various diets
on the outcome of tne toxicity test, the recommended food
source is live brine shrimp nauplii Artemia salina. H.T.
Smith et al. (1978) studied the effects of population
density and feeding rate on the final population density,
growth and fecundity of fathead minnows. It was found that
length and weight of fish increased with increased food
availability. As such, the fathead and sheepshead minnow
larvae should be fed at least 3 times a day during the week
and twice on the weekend for the duration of the study.
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August, L982
This recommendation is felt to adequately satisfy the
nutritional requirements of the minnow species, but does not
constitute an unnecessary requirement nor expense in the
test. An automatic brine shrimp feeder is discussed by
Schimmel and Hansen (1975), which could be employed on
weekends .
The brine shrimp should be analyzed for contaminants,
and contaminant levels should not exceed the specified
criteria for dilution water for maximum contaminant
concentrations, in order to minimize the possible effects on
the test species. Contamination of food by pesticides,
PCB's, phthalates , mercury, lead or other ubiquitous
substances would render the test results ambiguous.
Trout feeding requirements are different, in that the
yolk sac of newly hatched trout generally takes between 32
and 33 days to be absorbed depending upon temperature
(Atchison and Johnson 1975, Mauck et al . 1978). Therefore
feeding should commense at swim-up. Trout starter food
should be fed ad libitum five times a day, with the tanks
cleaned daily. This is in accordance with the early EPA
(1972b) partial chronic test method, with indicates trout
alevins and early juveniles should be fed trout starter food
a minimum of 5 times daily.
Fish are not fed for the 24-hour period prior to
termination of the test in order to minimize the effect
consumed food may have on analysis of fish weight.
3 . Facilities
a. General
The facilities needed to perform an early life stage
toxicity test as prescribed in the TSCA test guidelines
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August, 1982
include: (1) flow through test chambers (tanks) for
exposing embryos and fry; (2) embryo cups or screen trays
for holding the embryos until hatching; (3) test substance
delivery system; (4) a mixing chamber to promote mixing of
the test substance and dilution water; (5) a mechanism for
controlling and maintaining water at specified temperatures;
(6) apparatus capable of controlling the photoperiod at
specified regimes; (7) a device for removing particulate
matter and gas bubbles and for aerating the dilution water
as necessary; (3) suitable apparatus (e.g. magnifying
viewer) for examining embryos; (9) apparatus for precise
measurement of fish lengths and weight; and (10) facilities
for providing the necessary food (i.e. live brine shrimp
nauplii); and (11) facilities (or access to facilities) for
performing the required water quality analyses.
b. Construction Material
All pipes, embryo cups, screen trays, mixing chambers,
;netering devices, and test chambers should be made of
materials that minimize the release of chemical contaminants
into the dilution water or the adsorption of test
substances. Chemicals that leach from construction
materials may be toxic to test organisms or they may act
synergistically or antagonistically with test substances
thereby producing inaccurate results. Leaching of
undesirable substances from perfluorocarbon plastic, #316
stainless steel, and glass, and adsorption of test
substances to these materials is minimal. Rubber, copper,
brass, galvanized metal, lead and epoxy resins should not
come in contact with dilution water, stock solution, or test
solutions because toxic substances they contain may leach
into those media.
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August, 1982
All containers and pipes coming in contact with the
dilution water need to be conditioned before use in order to
leach out and wash away undesirable residues that may be
present.
When cast iron is used in freshwater systems, rust may
develop and its removal by strainers is recommended to
prevent fouling of the diluters and delivery systems.
c. Tes t Ch ambe rs
Test chambers, delivery systems, pipes or tanks exposed
to solutions that may come in contact with test organisms
should not be soldered or brazed. The materials used in
soldering or brazing contain lead, tin, copper, or zinc
which may leach into the solution and may be toxic to the
test organisms (Pickering and Vigor, 1965). Instead, .netal
parts should be welded or bonded with clear silicone
adhesive, the preferred bonding agent for all construction
materials. This adhesive is inert, and the acetic acid
which it releases is easily washed away or volatilized from
the system. However, the amount of adhesive which contacts
any test solution should be minimized because it may adsorb
test chemicals. It should be applied to the outs ides of
chambers and apparatus to minimize contact with the dilation
water and test substance. Epoxy glues are not recommended
for use because they may contain unreacted toxic monomers;
if it is necessary to use epoxy glues, allow a considerable
curing time prior to use.
Many different sizes of test chambers; have been used in
early life stage toxicity tests. There is no present
justification for requiring specific dimensions, and the
size of the test chambers may be considered acceptable if
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August, L932
the specified Clow rate and loading requirements can be
ach ieved.
d. Embryo Cups and Screen Trays
Embryos used to initiate early life stage toxicity tests
are exposed to the test substance during incubation in
embryo cups or on screen trays, where they can be carefully
observed and protected, and the optimal conditions to ensure
hatching success can be provided. Embryo cups are usually
glass retaining vessels (or similar sized sections of
polyethylene tubing) with stainless
steel or nylon screen bottoms or petri dishes with nylon
screen or stainless steel sides (Mount and Stephan 1967;
Mount 1968).
Various modifications of embryo cups have been used
successfully to incubate the embryos during toxicity
studies. Acceptable embryo incubator chambers Cor trout
have been described by Me Kim and Benoit (1971). Trays of
stainless steel screen with the sides turned up and
supported above the bottom of the exposure chamber (e.g. by
folded screen legs) are also acceptable, and have been used
successfully at the Environmental Research Laboratory at
Corvallis, Oregon.
Embryo cups should be labeled to facilitate
identification and tracking of embryos with the various test
substance concentrations. The cups should be suspended or
arranged in a test chamber to easure that the test solution
flows regularly into and out of each cup and that the
embryos remain submerged. Test substance delivery systems
or test chambers or both should be constructed so that the
embryos are not stressed by turbulence. As indicated
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August, 1932
previously, embryo cups can be oscillated by a rocker arm
apparatus (Mount 1968), or the water level in the test
chambers varied by means of a self-starting siphon (Hansen
et al . 1977). This oscillation serves to ensure a regular
flow of water through the embryo cup, which provides oxygen
needed by the developing embryos. It also helps to remove
debris, which in combination with stagnant water conditions
contributes to fungal infestation.
e. Test Substance Delivery System
In order to maximize the accuracy and precision of data
developed for use in evaluating the hazard a chemical
presents to the environment, it is necessary to minimize
variability in the testing procedure to the extent that such
reductions in variability are cost effective. To accomplish
this, the quantity of the test substance introduced by the
test substance delivery system should be constant as as
possible from one addition of test substance to the next.
Fluctuation in the quantity of test substance introduced
into the test chamber may affect the validity of the test
results more significantly than fluctuations in other test
conditions. The greater the variation in the quantity of
the test substance introduced, the greater the abnormalities
and spread of the response values; hence, the need to
calibrate and verify the calibration of the test substance
delivery system. Calibration includes determining the flow
rate through each chamber, and the proportion of stock
solution to dilution water delivered to each chamber.
Variations in the quantity of dilution water entering the
test chamber during a given time interval also may create
undesirable differences in test conditions from one test
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August, 1982
chamber to another. The concentrations of dissolved oxygen
and test substance in a test chamber, for example, may
decrease more rapidly in chambers having lower flow rates.
Metabolic products of the test fish, such as ammonia, and
products resulting from the degradation of the test
substance, may also accumulate to a greater extent in
chambers with lower flow rates. Differences from test
chamber to test chamber in the concentration of dissolved
oxygen, test substance, metabolic products and degradation
products, individually or in combination, may also result in
response values for the test organisms which are inaccurate
or which lack an adequate level of precision. Flow rates
through test chambers should not vary by more than 10% from
any one test chamber to any other. The delivery system
operation should be checked daily for normal operation
throughout the test. This is extremely important for
accurate interpretation of data results (Lloyd 1978)
Many test substance delivery systems are referenced in
the literature at large (Lemke et al. 1978) and can be
selected depending upon the specific characteristics and
requirements of the test substance. Proportional diluters
(Mount and Brungs 1967) are suitable for extensive use, and
have the following advantages: (1) timing problems are
minimal; (2) operation is simple and easy to understand; (3)
malfunctions are infrequent; and (4) a series of
concentrations can be delivered, each as much as 90% of each
oreceeding concentration. The main disadvantage is that it
is impractical to deliver a series of concentrations with a
dilution factor greater than 50% between concentrations
(Mount and Brungs 1967). In addition, proportional diluters
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August, 1982
are not commercially available everywhere and may require
ceilings more than eight feet high.
Many modifications to delivery systems have been
recommended by various researches, including: Brungs and
Mount (1970), McAllister et al. (1972), Schimmel et al.
(1974), and DeFoe (1975). Other relevant, systems are
discussed by Freeman (1971), Chandler et al. (1974),
Chandler and Partridge (1975), Smith et al . (1977) and Abram
(1973). Most delivery systems are operated by the
hydrostatic force of dilution water or the test solution in
a system of siphons (with and without float operated
values), or by electro-machanical means. Metering pump
systems (Garton 1980) can be simple, compact, easy to set
up, portable, and relatively accurate if high quality pumps
are used. Such pumps will last a long time with proper
ma intenance.
Proportional diluters and metering pumps have the
capability of delivering consistent quantities of a test
substance and dilution water to the test chambers, and have
been used extensively in aquatic toxicity tests. However,
different delivery systems which incorporate proportional
diluters or metering pumps exhibit individual
idiosyncrasies. Due to possible malfunctions caused by such
idiosyncrasies, it is necessary to calibrate each system to
verify the concentration of the test substance and the
volume of dilution water which each system actually
delivers .
A small chamber to promote mixing of toxicant-bearing
water and dilution water should be used between the test
substance delivery system and the test chambers for each
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August, 1982
concentration. Benoit and Puglisi (1973) describe such a
flow-splitting chamber to promote thorough mixing. When a
single proportional diluter is used, separate delivery tubes
from the mixing chamber to each duplicate test chamber are
necessary to delivery accurate and identical concentrations
of the test substance.
Particulate material and gas bubbles should be removed
from the test system. Test substance concentrations may be
altered by sorption to particulate matter or by
volatilization due to excessive gas bubbles. Both
particulate material and gas bubbles may clog the diluter
system. To avoid this problem, an apparatus capable of
removing particulate matter or gas bubbles or both from the
dilution water is recommended (US EPA 1975, US EPA 1972a).
f. Cleaning
Before use, test systems are cleaned to remove dust,
dirt and other debris and residue that may remain from
previous uses of the system. Any of these substances may
affect the results of a test by sorption of test materials
or by exerting an adverse effect on the test organisms. New
chambers are cleaned to remove any chemical or dirt res idues
remaining from manufacture or accumulated during
construction and storage. Detergent is used to remove
hydrophobic or lipid like substances. Acetone may be used
for the same purpose, and as a final rinse. It is important
to use pesticide-free acetone to prevent the contamination
of the chambers with pesticides which may be toxic to the
test organisms or which might otherwise influence the
outcome of the test. Hypochlorite at 200 mg/liter is useful
for removing organic matter and for disinfection. A
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August, 1982
solution containing 200 mg hypochlorite/liter is made by
adding 5 ml. of household bleach (chlorine) to 1 liter of
water. However, acid and hypochlorite should not be used
together because hazardous fumes may be produced. Acid
(e.g. 5% nitric acid) is useful for removing mineral
deposits, metal residues and bases from the system.
For example, a new system would be typically cleaned by
washing with detergent and rinsing with water, reagent-grade
acetone, water, acid (e.g. 5% nitric acid), and twice with
dilution water.
At the end of a test, if the test substance delivery
system, embryo cups or test chambers are to used again, they
should be: (a) promply emptied; (b) rinsed with water; and
(c) cleaned by a procedure appropriate for removing the
substance tested as well as other debris. It is easiest to
clean the equipment immediately following the test
termination, before chemical residues and organic matter
become embedded or absorbed. Conditioning should be
considered an important part of cleaning.
g. Dilution Water
Variations in water quality parameters have been found
to influence the results of toxicity tests involving aquatic
organisms (Tucker and Leitzke 1979). For instance, Mount
(1968) and Mount and Stephan (1969) conducted experiments to
determine the acute and chronic toxicity of copper to the
fathead minnow in hard and soft water. The dilution water
used in each test was controlled to maintain nearly constant
hardness, alkalinity and pH. In addition, temperature was
maintained within a narrow, optimum range. Results of these
tests indicated that hardness, and to a lesser extent, other
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August, 1982
characteristics such as pH, acidity, and alkalinity,
affected both acute and chronic toxicity of copper. Brungs
et al. (1976) stated that many environmental factors
influence the toxicity of copper and cited several authors
that documented effects of oxygen, temperature, hardness,
turbidity, carbon dioxide, magnesium salts, organic
compounds, nitr ilotriacet ic acid and spent sulfite liquor.
Carroll et al . (1979) stated that many workers have reported
that heavy metals are less acutely toxic to aquatic
organises in hard water than in soft. While data on all the
possible effects of various water chemistry parameters on
the toxicity of chemicals to fish are very incomplete, those
which are available show clearly that variations in certain
water chemistry parameters may cause variation in the
results of toxicity tests with fish. In view of the
Agency's need for accurate data in evaluating the hazard of
a chemical and the risk it poses to the environment, the
dilution water used in performing toxicity tests should show
evidence of consistency in its chemical make-up.
A dependable source of clear surface or ground water
will usually have a greater consistency in its chemical
make-up than a municipal water supply. Municipal water may
originate from several sources which may differ
coas iderably. In addition, municipal water frequently is
treated chemically as part of a purification process. Since
the proportions in which water from different sources are
mixed, and since the chemical treatment given water during
the purification process may be different from time to time,
the chemical make-up of municipal water may vary
coas iderably.
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August, 1982
The long duration of the test, and the large volume of
water required for flow-through conditions economically
prohibit the use of reconstituted water (freshwater or
s altwater) .
Fish culturists do not know all of the conditions
required to maintain fish health, nor do they know all of
the components in water that adversely affect the health of
fish (Brauhn and Schoettger 1975). Nevertheless, to avoid
possible inconsistencies and inaccuracies in test results,
minimal variability and healthy fish are needed for toxicity
tests. There is, therefore, a need to determine that the
dilution water is capable of suporting the fish species to
be tested in a healthy condition for the duration of the
test period.
An appropriate way to make such a determination is to
place young fish of a sensitive species, preferably the one
to be used in subsequent tests, in the dilution water for an
extended period of time and observe their behavior, growth
and development. Ideally, such observations would be made
by experienced fisheries biologists familiar with certain
stress reactions which are difficult for an untrained
observer to identify (Brauhn and Schoettger, 1975). As an
indication of the uniformity of the dilution water, it is
recommended that certain water chemistry parameters be
measured at least twice a year, or more frequently (as
specified) if it is suspected that one or more of those
parameters has changed significantly. The water chemistry
parameters and the maximum acceptable concentrations for
contaminants specified in the test guidelines are those
considered not to adversely affect fish (US EPA 1973, APHP,
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August, 1982
AWWA, WPCF 1975). Recognizing that some variation in water
chemistry is normal in natural freshwater, a 10 percent
fluctuation from month to month in water hardness,
alkalinity and conductivity and a variance of 0.4 pH unit is
considered acceptable. Variations in excess of the
concentrations or values cited in the standard may alter the
value of the data developed in the toxicity test, and render
the results ambiguous.
The quality of dilution water is also important for
toxicity tests with salt water organisms (Bahner et al.
1975). Dilution water for conducting early life stage
toxicity tests with the sheepshead minnow is considered to
be of constant quality if the minimum salinity is greater
than 15°/00 and the weekly range of salinity is less than
15°/00. The monthly pH range should be less than 0.8 unit
(US EPA 1978, Hansen and Parrish 1977, Hansen and Schimmel
1975, Schimmel et al. 1974).
Specially designed systems are usually necessary to
provide seawater from natural sources, and the water should
be filtered through a pore size less than 20 micrometers in
order to remove larval predators (Parrish et al. 1977,
Goodman et al . 1976, Bahner et al . 1975, Schimmel et al.
1974, USEPA 1978).
The dilution water (fresh or salt) should be intensively
aerated by such means as air stones, surface aerators, or
screen tubes (Penrose and Squires 1976, Rucker and Hodgeboom
1953) before introduction of the toxicant. Aeration should
not be employed after the test substance has been introduced
since the process of aeration may result in a loss of the
test substance through volatilization. Adequate aeration of
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August, 1982
the dilation water will help bring the oil and the
concentration of dissolved oxygen and other gases into
equilibrium with air, and minimize oxygen demand and effect
removal of volatile compounds which may be present.
A variety of microorganisms are found in many water
supplies. Their presence in dilution water may be
undesirable as they may be infectious to the test fish.
Dilution water that may be contaminated with such
undesirable microorganisms should be passed through membrane
filters or an ultraviolet sterilizer equipped with an
intensity meter (Bullock and Stuckey 1977). Efficacy of the
sterilizer may be determined by using standard plate count
methods (APHA, AWWA, WPCF 1975).
Additional parameters cannot be specified for freshwater
dilution water quality, because of the wide geographical
distribution of available testing facilities. Specific
requirements for hardness, etc., would preclude those
facilities which failed to meet such requirements.
4. Environmental Conditions
a. Dissolved Oxygen
Fish embryos and developing larvae are particularly
sensitive to deficiencies of oxygen (Doundoroff and Shumway
1970). Exposure of fish to a test substance may increase
the rate at which they consume oxygen, with a resulting
rapid uptake of dissolved oxygen. Some test substances
undergo oxidation when introduced into the test chamber.
This may result in a chemical or biochemical oxygen demand,
depending on the nature of the oxidation which removes
dissolved oxygen from the surrounding water.
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August, 1982
Microorganisms present in the test chamber also may
create a demand for the dissolved oxygen. The normal oxygen
needs of the fish, with possible chemical or biochemical
oxygen demands of the test subs tance, and oxygen consumption
by micoorganisms, individually or in combination, could
reduce the level of dissolved oxygen below that required by
the test fish. Oxygen to replace that which has been
depleted can be provided by increasing dilution water flow
rate.
The concentration of dissolved oxygen in the dilution
water should be between 90% and 100% saturation, the optimal
range for all the test species. A minimum level of 60%
saturation of dissolved oxygen has been recommended by
earlier groups and committees for test solution water ased
in early life stage toxicity studies (APHA, AWWA, WPCF 1975,
USEPA 1972a,b). This minimum level of dissolved oxygen is
required in order to avoid possible variations in test
results due to the effects of anoxia on the test fish.
Doudoroff and Shumway (1970) noted that there is very little
agreement in the literature on the reported findings of the
effects and minimum requirements of dissolved oxygen for
fish. These authors noted that pertinent information on the
effects of low concentrations of dissolved oxygen on
fecundity and embryonic development is also very limited;
but embryos of the fathead minnow appear to be adequately
protected from stress at oxygen concentrations above 5 mg/1
at temperatures normal for this species. Similarly, Brungs
(1971b) studied the chronic exposure of fathead minnows to
continuous reduced levels of dissolved oxygen in order to
evaluate effects on reproduction, fry growth and fry
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August, 1982
survival. Results indicated a statistically significant
reduction in fry growth at all concentrations below the
control (7.26 mg/1), and that time to hatch was increased at
each successively lower dissolved oxygen concentration. No
effects were observed on percentage hatch. The highest
dissolved oxygen concentration in this study (other than the
control) was 5.0 mg/1; this indicates that 60% dissolved
oxygen saturation level appears to be somewhat less than an
ideal minimum concentration for early life stage toxicity
tests. These data support the requirement of a minimum
level of dissolved oxygen for the test species specified in
the early life toxicity test guidelines, especially since
the minimum dissolved oxygen requirements of warm water
species are generally not as critical as those for cold
water species (Doudoroff and Shumway 1970).
b. Flow Rate
The flow of the water through the test chambers
minimizes the accumulation of metabolic products such as
ammonia which, if allowed to accumulate, could reach
concentrations lethal to the fish or alter their seasitivity
to test substances. The build-up of organic matter within
the exposure chambers might provide a nutrient source for
bacteria present in the water. Bacteria using oxygen to
metabolize and decompose the organic matter in the tanks
could reduce the dissolved oxygen concentrations of the
water. Decreased dissolved oxygen concentrations as well as
the accumulation of toxic metabolic products could increase
the likelihood of disease in the test fish (Brauhn and
Schoettger, 1975).
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August, 1982
To avoid the possibility of a build-up of metabolic or
degradation products, the depletion of dissolved oxygen, the
adsorption of the test substance to sediments and the walls
of the test system, and to ensure an adequate mixing of the
test solution, a minimum flow rate equivalent to six tank
volume changes a day is specified. This flow rate appears
to be a practical flow rate based upon past experience of
successful use and satisfies Sprague's (1969) recommendation
for a 90% replacement time of 8-12 hours.
c . Lo ad i ng
The grams of organism per liter of solution in the test
chambers should not be so high that it affects the results
of the test. The most important parameters dfifected by
overloading are the dissolved oxygen concentration, the
waste metabolite accumulation, the concentration of the test
substance, and stress to the test organisms due to
crowding. Hence, loading should be limited to assure
that: 1) the concentration of dissolved oxygen and test
substance do not decrease below acceptable levels; 2) tnat
above acceptable levels; and 3) that organises are not
stressed due to crowding.
The number of fish that can be placed in a given test
chamber depends upon the test solution volume and the rate
of flow. The upper limits are 2 grams of fish per liter of
test solution in the test chamber at any given time, and 0.1
grams of fish per liter of test solution passing through the
test chamber in 24 hours.
It is felt that these loading requirements can be easily
and practically satisfied by all testing facilities without
requiring modification to present equipment because of the
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August, 1982
relatively small size and weight of the embryos and newly
hatched larvae and developing juveniles of the recommended
test species .
The levels of dissolved oxygen and ammonia cited in the
test guideline are recommended for the health of the fish.
The 20 percent limit on the uptake of test substance by the
fish is recommended in order to avoid a reduction in the
concentration of test substance to which the fish are
exposed, such that the results become unsuitable for the
purposes for which the test is run.
d. Temperature
Test temperature is dependent upon the species selected,
but should not deviate instantaneously by naore than 1.5°C
from the selected temperature during the test period. The
effects of sudden temperature changes on fish may range from
death to temporary impairment of physiological function,
depending upon the magnitude of the temperature change, the
tolerance of the species to temperature fluctuations and the
circumstances and duration of the exposure. Differences
between the temperature at which fish embryos are obtained
(i.e. culture or external shipments) and the temperature at
which toxicity tests are conducted could have an adverse
effect on the developing embryos. Therefore, it is
desirable to obtain embryos at temperatures as close as
possible to the temperatures at which the test will be
conducted, and to acclimate embryos as specified. The test
temperatures selected are: fathead minnow - 25°C (Brungs
1971a, USEPA 1972, Brauhn and Schoettger 1975, Brungs and
Jones 1977); sheepshead minnow - 30°C (Schimmel and Hansen
1974, Hansen and Parrish 1977, USEPA 1978); and brook and
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August, 1982
rainbow trout - 10°C for embryos and 12°C for alevins and
fey (McCormick et al. 1972, Hokanson et al . 1973, Brungs and
Jones 1977). These temperatures are generally accepted as
suitable for the fish species indicated and they are the
ones most frequently cited for use in conducting early life
stage toxicity tests and for culturing the species
selected. For example, Thatcher et al. (1976) studied the
effects of temperature and chlorine i;o
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August, 1932
temperatures which have been successfully and widely used in
toxicity tests. Extensive studies have not been reported in
the literature which identify the optimal temperature at
which early life stage toxicity tests with each test species
should be run. There is, however, evidence that suggests
the need to standardize the test temperatures. A number of
investigators have reported toxicity data for several
substances which indicate that the toxicity of those
materials varies with temperature (L.L. Smith et al. 1978,
Brown et al . 1967, Tucker and Leitzke 1979, Alexander and
McClarke 1978). Therefore, even though data are lacking to
unequivocally support selection of an optimal test
temperature, it is recommended that early life stage
toxicity tests be conducted at the test temperatures
specified in the test guidelines. This is particularly
necessary for comparative purposes.
Temperature is normally maintained by preheating and
aerating the dilution water or by placing the test aquaria
in temperature controlled water baths (Syrett and Dawson
1972, McCormick and Syrett 1970, Bahner et al. 1975). It
should be noted that the latter method is not common
practice for flow-through toxicity tests.
e. Light
Light is recognised as an important environmental
variable by most organizations concerned with the
development of uniform testing methods for aquatic
toxicology. Yet, few studies on the effects of a light
regime or light intensity on toxicity have been investigated
and reported. Toxicity tests with pulp mill effluents did
not demonstrate a significant variation in toxicity
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August, 1982
associated with light-dark p'notoperiods of 8-16 hoars and
16-8 hours (McLeay and Gordon 1978). However, in another
study using Daphnia (Crosby et al. 1966), the photoperiod
was shown to have an effect on the toxicity test results.
Although there are insufficient data on which to base
the selection of a photoperiod to be used in fish toxicity
tests, it is generally recognized that a standard
photoperiod should be employed. This serves to minimize the
influence of variations of light conditions on test data, as
does standardization of other test conditions (i.e.
temperature, dissolved oxygen, etc). The source material
cited in Section II.C.I: Test Species, indicates that peak
population levels and spawning activity (with the exception
of trout) occur in the warmer months during which the
photoperiods are naturally the longest. Thus, for the
minnow species, longer photoperiods appear to correspond
with natural conditions which are optimal for growth,
development and survival. Standard Methods for the
Examination of Water and Wastewater (American Public Health
.Association, American Water Works Association, Water
Pollution Control Federation 1975) states that during any
test exposure to light should be based upon what is normal
for and required by the species, and that different light
intensities are required for different organisms and life
stages. One example cited in Standard Methods is the
requirement of darkness (or very low light intensity) for
trout eggs.
The photoperiods recommended by the ASTM (1980) for
conducting early life stage toxicity tests with fishes
are: brook trout 12-14 hours light; sheepshead minnow 16
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August, L982
hours light; and fathead minnow 16 hours light. A
transition period of 15 to 30 minutes between light and dark
is recommended (ASTM 1980) in order to avoid any possible
variation in test results attributable to sudden light-dark
changes. Apparatus which can be incorporated into indoor
lighting systems to provide a programmed ohotoperiod can be
designed according to Drummond and Dawson (1970) or by
Wickham et al . (1971). Durotest vitalite (optima FS) lamps
and wide spectrum Grov/-lux fluorescent tabes are
recommended, based upon past use.
D. Reporting
The reporting requirements specified in the test
guidelines are considered essential to complete a thorough
and proper evaluation of the test results. The required
information is deemed necessary by EPA to: (1) establish
that the test was conducted according to specifications; (2)
evaluate those conditions and procedures that could affect
the results of the test; and (3) supply sufficient
information to accurately interpret results, which include
independent analysis of statistics and conclusions. Due to
the inherent flexibility of the test guidelines, reporting
requirements are necessary to ascertain the conditions,
parameters and observations germane to the performance of
the test.
Additional information may be needed when a concern
exists relative to the results or validity of the test.
Therefore, the location of the raw data storage is needed in
order that additional information can be located
expedi tiously, if necessary, for a detailed evaluation or
for enforcement purposes. It is recognized that some
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August, 1982
chemical companies maintain possession of test data, while
other request the test facility to keep such records.
III. Economic Aspects
Five laboratories were surveyed to obtain estimated
costs for performing the early life stage toxicity test
outlined in the test guideline. A price range of $9,000 to
$40,000 with a "best estimate" of $15,608 was reported. An
additional cost estimate was made by separating the
guideline into components and estimating the cost of each
component, including direct labor cost, overhead cost, other
direct costs, general and administrative costs, and profit
or fee. The protocol best estimate of cost for the minnow
species was $11,348, with an estimated range of $5,674 to
$17,022 based on +_ 50 percent of the best estimate. The
protocol best estimate of cost for the trout species was
$14,354, with an estimated range of $7,177 to $21,531 based
on _+_ 50 percent of the best estimate. The protocol estimate
for the mean of both species was $12,851 with an estimate
range of $6,426 to $19,277.
The test guidelines described for minnows and trout have
basic differences which result in different cost
estimates. For example, the test with minnows requires a
post-hatch exposure time of 28 days, while the test with
trout requires a post-hatch exposure time of 60 days. This
fundamental difference results in a higher cost in the trout
test based on increases in animal care, observation
intervals and analytical monitoring. Various other factors
can affect the cost estimates, including the nature of the
chemical, overhead rates, use of outside consultants and the
degree of effects manifestation. For example, if a majority
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August, 1982
of: test organisms exhibited severe manifestations of
r.'.'xi :i ty, additional photography and detailed observations
with possible pathology would be required. The nature and
number of analytical assays is also a major cost factor.
The above cost estimates were made assuming that all the
requirements of Good Laboratory Practice Standards, as
specified in section (d) of the early life stage toxicity
test guideline, and related considerations are contained in
the cost aialysis report for the ecotoxicity standards by
Enviro Control, Inc. (1930).
In a cost analysis of subpart E, Hazard Evaluation:
Wildlife and Aquatic Organisms, of the pesticide guidelines,
several contract and captive industry laboratories were
surveyed in 1978 and again in 1980 to determine cost
estimates for testing (US EPA 1980a). The cited costs did
not differentiate between species, however, the unit cost
estimate for the fish embryo-larvae (early life stage) test
was $11,500, and can assumed to be for fathead 'iinnow.3.
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August, 1982
IV. References
ABC Protocol No. 7809 (Analytical Bio Chemistry
Laboratories, Inc.) Revised July 16, 1979. Procedure
for conducting early life stage studies with fathead
minnows (Pimephales promelas) in a flow-through system.
ABC Protocol No. 7810 (Analytical Bio Chemistry
Laboratories, Inc.) Revised Sept. 5, 1979, Procedure
for conducting early life stage studies with rainbow
trout (3almo gairdneri) or brook trout (Salvelinus
fontinalis).
Abram FSH. 1973. Apparatus for control of poison
concentration in toxicity studies with fish. Water
Research. 7:1875-1879.
Adelman IR, Smith LL. Jr. 1976. Fathead minnows
(Pimephales orornelas) and goldfish (Carassius auratas)
as standard fish in bioassays and their reaction to
potential reference toxicants.
J. Fish. Res. Board Can. 33:209-214.
Alexander DG, McClarke RV. 1978. The selection and
limitations of phenol as a reference toxicant to detect
differences in sensitivity among groups of rainbow trout
(Salmo gairdneri). Water Research. 12:1085-1090.
Allison DT, Hermanutz RO. 1977. Toxicity of Diazinon
to brook trout and fathead minnows. EPA 600/3-77-060.
79p.
ASTM (American Society for Testing and Materials).
1980. Standard practice for conducting toxicity tests
with the early life stages of fishes. Draft f 4 .
Atchison GJ, Johnson HE. 1975. The degradation of DOT
in brook trout eggs and fry. Trans. Am. Fish. Soc.
4:732-784.
Banner LH, Craft CD, Nimmo DR. 1975. A saltwater flow-
through bioassay method with controlled temperature and
salinity. Prog. Fish. Cult 37:126-129.
Benoit DA, Puglisi FA. 1973. A simplified flow-
splitting chamber and siphon for proportional
diluters. Water Research. 7:1915-1916.
52
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August, 1982
Benoit DA, Carlson RW. 1977. Spawning success of
fathead minnows on selected artificial substrates.
Prog. Fish. Cult. 39(2):67-69.
Blair et al. 1968. Vertebrates of the United States.
2nd Ed. McGraw Hill, New York. 616p.
Box GEP, Hunter WG, Hunter JS. 1978. Statistics for
experimenters. Wiley-Interscience. New York. 653p.
Brauhn JL, Schoettger RA. 1975. Aquisition and culture
of research fish: rainbow trout, fathead minnow, channel
catliish, and bluegills. Ecological Research Series.
EPA 660/3-75-011. U.S. Environmental Protection
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Brungs WA. 1971(b). Chronic effects of low dissolved
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53
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Bullock GL, Stuckey HM.
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Fish. Res. Board Can. 34:1244-
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Chandler JH, Sanders HO, Walsh DF.
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Darnell RM. 1958. Food habits of fishes and larger
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54
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ES-8
1982
DeFoe DL. 1975. Multichannel toxicant injection system
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ES-8
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Larson GL, Hutchins FE, Schlesinger DA. 1977. Acute
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Lemke AE, Brungs WA, Halligan BJ. 1978. Manual for
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56
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ES-8
August, 1982
Lloyd R. 1978. The use of the concentration-response
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Macek KJ, Sleight BH III. 1977. Utility of toxicity
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Manner HW, DeWese CM. 1974. Early embryology of the
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Anat. Rec. 180:99-110.
Martin DM. 1973. Freshwater laboratory bioassays- a
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Acad. Nat. Sciences. Phila., Pa. 3:51 pp.
Mauck WL, Mehrle PM, Mayer FL. 1978. Effects of the
polychlorinated biphenyl Aroclor 1254 on growth,
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McAllister WA, Mauck WL, Mayer FL. 1972. A simplified
device for metering chemicals in intermittent-flow
bioassays. Trans. Amer. Fish. Soc. 3:555-557.
McCormick JH, Syrett RF. 1970. A modular controlled-
temperature apparatus for fish eggs incubation and fry-
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National Water Quality Laboratory Duluth, Minn. p. 1-18.
McCormick JH, Hokansen KEF, Jones BR. 1972. Effects of
temperature on growth and survival of young brook trout,
Salvelinus fontinalis. J. Fish. Res. Board Can.
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McKim JM, Benoit DA. 1971. Effects of long-term
exposures to copper on the survival, growth, and
reproduction of brook trout (Salvelinus fontinalis)
J. Fish. Res. Bd. Can. 28(5):655-662.
57
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ES-3
August, 1982
McKim JW, Arthur JW, Thorslund TW. 1975. Toxicity of a
linear alkylate sulfonate detergent to larvae of four
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McKim JM. 1977. Evaluation of tests with early life
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J. Fish. Res. Board Can. 34:1148-1154.
McLeay DJ, Gordon MR. 1978. Effect of seasonal
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J.Fish. Res. Board Can. 35:1388-1392.
Mehrle PM, Mayer FL, Johnson WW. 1977. Diet quality in
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Mount DI, Stephan CE. 1969. Chronic toxicity of copper
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Mount DI. 1973. Chronic effect of low pH on fathead
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Water Research. 7:987-993.
Parrish PR, Dyar EE, Lindberg MA, Shanika CM, Enos JH.
1977. Chronic toxicity of methoxychlor, malathion and
carbofuran to sheepshead minnows (Cypr inodon
variegatus). EPA 600/3-77-059. 36 p.
58
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ES-8
August, 1982
Penrose WR, Squires WR. 1976. Two devices for removing
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Pickering QH, Cast MH. 1972. Acute and chronic toxicty
of cadmium to the fathead minnow (Pimephales
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59
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ES-8
August, 1982
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Recommended bioassay procedures for fathead minnow
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60
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USEPA 1972(b) U.S. Environmental Protection Agency.
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EPA-660/3-75-009. 67 p.
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SEED GERMINATION/ROOT ELOGATION TOXICITY TEST
OFFICE OF TOXIC SUBSTANCES
OFFICE OF PESTICIDES AND TOXIC SUBSTANCES
U.S. ENVIRONMENTAL PROTECTION AGENCY
WASHINGTON, D.C. 20460
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Office of Toxic Substances EG-12
Guideline for Testing Chemicals August, 1982
SEED GERMINATION/ROOT ELONGATION TEST
(a) Purpose. The guideline in this section is intended for
use in developing data on the acute toxicity of chemical
substances and mixtures ("chemicals") subject to environmental
effects test regulations under the Toxic Substances Control Act
(TSCA) (PUB.L. 94-469, 90 Stat. 2003, 15 U.S.C. 2601 et seg.).
This guideline prescribes test procedures and conditions using
seed of commercially important terrestrial plants to develop data
on the phytotoxicity of chemicals. The United States
Environmental Protection Agency (USEPA) will use data from these
tests in assessing the hazard of a chemical to the environment.
(b) Definitions. The definitions in Section 3 of the Toxic
Substances Control Act (TSCA) and the definitions in Part 792—
Good Laboratory Practice Standards apply to this test
guideline. The following definitions also apply to this
gu ideline:
(1) "ECX" means the experimentally derived chemical
concentration that is calculated to effect X percent of the test
criterion.
(2) "Embryo" means the young sporophytic plant before the
start of germination.
(3) "Germination" means the resumption of active growth by
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an embryo. The primary root should attain a length of 5mm for
the seed to be counted as having germinated.
(4) "Hypocotyl" means that portion of the axis of an embryo
or seedling situated between the cotyledons (seed leaves) and the
radicle.
(5) "Radicle" means that portion of the plant embryo which
develops into the primary root.
(6) "Test solution" means the test chemical and the dilution
water in which the test chemical is dissolved or suspended.
(c) Test procedures — (1) Summary of the test. (i) Seed
should be separated into appropriate size classes, and that size
class containing the most seed used exclusively for the test.
Fresh test solutions should be added to petri dishes that have
been completely filled with either precleaned quartz sand, 200
micron glass beads, or other inert material. The seed should
then be positioned on the substrate allowing adequate roan for
anticipated growth. It is recommended that the radicle end of
the seed be aligned in the direction of this growth. Petri dish
lids should be used to hold the seed in place, and the dishes
sealed with tape. For those chemicals that are insoluble in
water and that should be sorbed to the substrate, deionized or
glass-distilled water should be added to the substrate prior to
positioning the seed.
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(ii) The dishes should be placed in a seed germinator or
other growth facility at a slight angle to facilitate linear root
growth. Seed should be incubated in the dark until at least 65
percent of the control seed have germinated and developed roots
that are at least 20 mm long.
(iii) The number of seed that germinate should be counted,
and root lengths measured. Concentration response curves, EC
10's, and EC 50's for seed germination and root elongation should
be determined and reported for each of the species tested.
(2) Chemical application. (i) Test chemicals that are
soluble in water should be dissolved in deionized or glass
distilled water and added to the substrate in the petri dishes at
the start of the test.
(ii) Test chemicals that are insoluble in water but which
can be placed in aqueous suspension with a carrier should be
suspended in deionized or glass-distilled water with the carrier
and then added to the petri dishes. The carrier should be
soluble in water, relatively non-toxic to plants, and should be
used in the minimum amount required to dissolve or suspend the
test chemical. There are no preferred carriers; however,
acetone, gum arabic, polyethylene glycol, ethanol and others have
extensively been used in testing herbicides, plant growth
regulators, fungicides, and other chemicals that affect plants.
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August, 1982
Tests of the carrier effect should be included in the test
experimental design and conducted simultaneously as controls.
(iii) Water-insoluble chemicals for which no non-toxic
water-soluble carrier is available, should be dissolved in an
appropriate volatile solvent. The solution and substrate should
be placed in a rotary vacuum apparatus, and evaporated, leaving a
uniform coating of test chemical on the substrate. A weighed
portion of the substrate should be extracted with the same
organic solvent and the chemical assayed before the containers
are filled. Solvent controls should be included in the
experimental design and tested simultaneously. Deionized or
glass distilled water should be added to the treated substrate
prior to positioning the seed on the substrate.
(3) Range-Finding Test. (i) A range-finding test should be
conducted to establish (A) if definitive tejting is necessary and
(B) test solution concentrations for the definitive test.
(ii) The seed should be exposed to a chemical concentration
series (e.g., 0.01, 0.1, 1.0, 10, 100, and 1,000 mg/1). The
lowest concentration in the series, exclusive of controls, should
be at the chemical's detection limit. The upper concentration,
for water soluble compounds, should be the saturation
concentration.
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August, 1982
(iii) The test consists of one run for each of the
recommended plant species or selected alternates. A minimum of
15 seed per species should be exposed to each chemical
concentration and control. The test period may be ended when at
least 65 percent of the control seed have germinated and
developed roots that are at least 20 mm long. The exposure
period may be shortened if data suitable to establish the test
solution concentration series for the definitive test can be
obtained in less time and if the definitive test is to be
conducted. No replicates are required; and nominal
concentrations of the chemical are acceptable unless definitive
testing is not required as specified below.
( iv) Definitive testing is not necessary if the highest
chemical concentration tested results in less than a 50 percent
inhibition of germination or reduction in root growth or if the
lowest concentration tested (analytical detection limit) results
in greater than a 50 percent inhibition of germination or
reduction in growth.
(v) Graphical analysis of the range-finding data facilitates
selection of chemical concentrations for the definitive test.
(4) Definitive test. (i) The purpose of the definitive
test is to determine the concentration-response curves, the EC
10's, and EC 50's for seed germination and root elongation for
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August, 1982
each species tested, with the minimum amount of testing beyond
the range-finding test.
(ii) The seed of each species tested should be exposed to at
least 6 concentrations of the chemical chosen in a geometric
series in which the ratio is between 1.5 and 2.0 (e.g.,
2,4,8,16,32 and 64 mg/1). The concentration ranges should be
selected to determine the concentration response curves between
the EC 10 and EC 50 for both germination and root elongation.
Test solutions or substrate extracts should be analyzed to
determine chemical concentration prior to use. Selection of seed
from the size class lot to be exposed to each test concentration
should be unbiased.
(iii) At least three replicates, each with at least 10 seed
per species should be tested for each concentration and control.
(iv) Every test should include controls consisting of the
same dilution water, conditions, procedures and seed from the
same lot used in the exposure group, except that none of the
chemical is added. If a carrier (solvent) is needed to suspend
or disperse the chemical, a separate carrier control should also
be used.
(v) The test period may be ended when at least 65 percent of
the control seed have germinated and developed roots that are at
least 20 mm long. When both conditions are satisfied, the mean
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August, 1982
number of seed germinating and mean root length per treatment
(and control) can be determined. If the test chemical
concentration series does not bracket the EC10 through EC50 for
both germination and root elongation, the test should be repeated
(at a higher or lower concentration series). Concentration
response curves, EClO's and ECSO's for germination and root
elongation should be determined for eace species tested and
reported along with their 95 percent confidence limits.
(vi) Any abnormal seedling development or appearance such as
lesions, enhanced root growth (measured), discoloration,
swelling, loss of turgor, etc., should also be reported.
(vii) A randomized complete block design is recommended for
the definitive test with blocks delineated within the seed
germinator or growth chamber. If, for any reason, blocking is
not feasible total randomization within chambers is acceptable.
(viii) Temperature in the germination facility should be
recorded hourly. The pH of the test solutions should be recorded
at the initiation of the definitive test.
(5) [Reserved]
(6) Analytical measurements — (i) Test chemical. Stock
solutions should be diluted with glass distilled or deionized
water to obtain the test solutions. Standard analytical methods,
if available, should be used to establish concentrations of these
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August, 1982
solutions and should be validated before beginning the test. An
analytical method is not acceptable if likely degradation
products of the chemical, such as hydrolysis and oxidation
products, give positive or negative interference. The pH of
these solutions should also be measured prior to use.
(ii) Numerical. The number of seeds that germinate shall be
counted and root lengths measured for each definitive test
species. All root elongation measurements for a given species
should be made sequentially before proceeding to the next
species. Root length should be measured from the transition
point between the hypocotyl and root to the tip of the root.
Means and standard deviations should be calculated and plotted
for each treatment and control. Appropriate statistical analyses
should provide a goodness-of-fit determination for the
concentration response curves.
(d) Test conditions—(1) Test species. (i) Test plants
recommended for use include:
Lycopers icon esculentum (toma to)
Cucumis sativus (cucumber)
Lactuca sativa (lettuce)
Glycine max (soybean)
Brassica oleracea (cabbage)
Avena sativa (oat)
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August, 1982
Lolium perenne (perennial ryegrass)
Allium cepa (common onion)
Daucus carota (carrot)
Zea mays (corn)
(ii) Other species of economic or ecological importance to
the region of impact, may also be appropriate for testing. A
minimum of ten species should be tested.
(iii) Information on seed lot, the seed year or growing
season collected, and germination percentage should be provided
by the supplier of the seed. Only untreated seed (not treated
with fungicides, repellants, etc.) taken from the same lot, and
year or season of collection should be used in a given test. In
addition, all seed of a species used in a test should be from the
size class which contains the most seed. Damaged seed should be
discarded. Standard seed dockage sieves should be used to size
seed.
(2) Facilities — (i) Apparatus. (A) A seed germinator, or
other controlled environment chamber capable of maintaining a
uniform testing temperature of 25 ± 1°C is required. In
addition, the facilities should include work areas for sizing,
counting, and exposing seed for root measurement. If possible,
these areas should be isolated from other activities. A fume
hood may be needed when testing substances potentially hazardous
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EG-12
August, 1982
to human health. Apparatus for distilling and deionizing water
are needed unless reagent grade water is used. Refrigeration
facilities to hold the seed in cold storage (5°C) in moisture-
proof containers at seed moisture contents of less than 10
percent are also needed.
(B) Disposal facilities should be adequate to accomodate
spent glassware, sand, beads, and test solutions at the end of
each run and any bench covering, lab clothing, or other
contaminated materials.
(ii) Containers and support media. A minimum of 210 petri
dishes and sufficient sand or glass beads, or other inert
substrate to fill them are needed. Large (200 mm) glass petri
dishes are recommended. Perlite, vermiculite, or native soils,
should not be used as substrates.
(iii) Cleaning and sterilization. (A) All glassware and
the substrate should be cleaned following standard good
laboratory practice before each test. The substrate should be
washed in half strength concentrated nitric acid and rinsed with
a mild base followed by washes of glass-distilled or deionized
water. The pH of the washed substrate should be near neutral.
If the glass beads are to be reused, they should be heated to
100°C for 8-12 hours prior to acid washing. A dichromate
solution should not be used for cleaning beads or petri dishes.
10
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August, 1982
The sand and plastic petri dishes should not be reused.
(3) If fungal or other microbial contamination interferes
with seed germination such that germination is less than 65
percent in the controls, glassware should be sterilized and/or
the seed surface sterilized prior to use, e.g., the seed may be
soaked for 10 minutes in a 10 percent sodium hypochlorite
solution, then rinsed and soaked for one hour in glass-distilled
water.
(3) Test parameters. Environmental conditions should be
controlled to maintain incubation temperature at 25± 1°C in
complete darkness. If species other than the ten recommended for
use are tested, incubation conditions may have to be adjusted to
meet germination and root length criteria in the controls.
(e) Reporting. The sponsor should submit to the USEPA all
data developed during the test that are suggestive or predictive
of phytotoxicity. In addition to the general reporting
requirements prescribed in Part 792--Good Laboratory Practice
Standards, the following should be reported:
(1) Information on the source and history of the seed,
germination percentage reported by the supplier, and the seed
size class used for testing.
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August, 1982
(2) The number of seed of each species per treatment, the
number of replicates, carriers, incubation conditions, and seed
sterilization procedures.
(3) The concentration of the chemical added to each
treatment dish and its pH (pH is optional).
(4) If the range-finding test showed that the highest
concentration of the chemical tested (not less than 1,000 mg/1)
had no effect on the test species, report the results by species
and concentration and a statement that the chemical is of minimum
phytotoxic concern.
(5) If the range-finding test showed greater than 50 percent
inhibition of germination or root elongation at a test
concentration at the analytical detection limit, the results by
species and concentration and a statement that the chemical is
phytotoxic below the analytical detection limit.
(6) For each species included in,the definitive test, means
and standard deviations for germination and root length in each
treatment. In addition, concentration response curves with 95
percent confidence limits delineated, goodness-of-fit
determination, and EClO's and ECSO's identified.
(7) Methods and data records of all chemical and numerical
analyses including method validation and reagent blanks.
(8) The data records of the incubation temperature.
germination counts, and root length measurements.
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TECHNICAL SUPPORT DOCUMENT
FOR
SEED GERMINATION/ROOT ELONGATION TOXICITY TEST
OFFICE OF TOXIC SUBSTANCES
OFFICE OF PESTICIDES AND TOXIC SUBSTANCES
U.S. ENVIRONMENTAL PROTECTION AGENCY
WASHINGTON, D.C. 20460
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TABLE OF CONTENTS
Subject Page
I. Purpose 1
II. Scientific Aspects 1
General 1
Test Procedures 5
Chemical Application 5
Range-Finding Test 5
Definitive Test 6
Analytical Measurements 8
Test Conditions 10
Test Species 10
Selection 10
Facilities 13
Apparatus 13
Containers and Support Substrate 14
Cleaning and Sterilization 15
Environmental Conditions 15
Reporting 17
III. Economic Aspects 17
IV. References 19
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Office of Toxic Substances ES-9
August, 1982
TECHNICAL SUPPORT DOCUMENT FOR SEED
GERMINATION/ROOT ELONGATION TOXICITY TEST
I. Purpose
The purpose of this document is to provide the
scientific background and rationale used in the development
of Test Guideline EG-12 which uses the seeds of various
plant species to evaluate the toxicity of chemical
substances on seed germination/root elongation. The
Document provides an account of the scientific evidence and
an explanation of the logic used in the selection of the
test methodology, procedures and conditions prescribed in
the Test Guideline. Technical issues and practical
considerations are discussed. In addition, estimates of the
cost of conducting the test are provided.
II. Scientific Aspects
A. General
Chemicals may influence seedling vigor, a characteristic
of increasing importance with increased mechanization in
agriculture. Some crop plants (e.g., lettuce) with intense
culture practices require high germination rates and
vigorous growth. A single seed should germinate and
establish a plant (planting to stand), and all plants should
reach maturity simultaneously for once-over machine
harvesting (Pollock and Roos 1972). Seedling establishment
in forests also depends on vigorous root growth to survive
environmental stresses and competition with other plants for
light, water, and nutrients. In general, chemicals that
reduce or delay germination and retard maturity of crops
typically result in economic loss.
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In agricultural land with other stresses (e.g., drought,
salinity), the additional presence of chemicals which are
toxic may be the limiting factor for production of crops on
that land. In addition, chemicals may enter human food
chains through processes associated with soil/plant inter-
actions, uptake, translocation, and accumulation in food and
forage crops. In natural systems, affected species are less
competitive and with selection for tolerant species result
in altered species diversity, density, and frequency of
occurrence.
Toxic substances may also cause widely varying and
significant effects on plant community dynamics. Many
chemicals affect plants selectively, with some species
sensitive to and others tolerant of the s ame chemical and
dose. This selectivity may directly affect the successional
replacement of one plant species by another, either by
hastening the departure of an early successional species or
by inhibiting the establishment of a later- stage
successional species (Whittaker 1970). Succession also may
be indirectly affected by inhibition of soil organisms.
Toxic materials may produce high species diversity in some
communities while reducing diversity in others (Brown
1978). Where plant growth and soil organisms are completely
inhibited, soil degradation, instability, and eventually
erosion, may result.
Seed germination and root elongation were selected to
measure phytotoxicity for the following reasons:
o Seed germination is a critical stage in plant
development. It marks the transfer from a period
of dormancy to one of active growth and high
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o metabolic activity. This process involves rapid
cell division, expansion, and formation of the
essential structures of a normal plant (Berlyn
1972).
o Phytotoxicity, as demonstrated by the inhibition of
seed germination and root elongation, has been used
in determining selective toxicities of herbicides
(Behrens 1970, Horowitz 1976, Santelmann 1972),
screening plants for heavy metal tolerances (Imai
and Siegel 1973, Konzak et al. 1976, Siegel 1977,
Whalley et al. 1974, Wilkins 1957), determining
salinity tolerance (Durrant et al. 1974, Neiman and
Poulsen 1971), evaluating chemicals for toxic
effects (Hikino 1978, Rubinstein et. al. 1975), and
studying allelopathic substances (Asplund 1969,
Bode 1958, Garb 1961, Muller 1965).
o Seed germination and root elongation show a
reproducible response in proportion to the chemical
concentration tested. In the literature pertaining
to herbicide, heavy metal, and toxic chemical
effects on plants, root elongation appears to give
a more sensitive response than does seed
germination and is the preferred test endpoint.
Few herbicidal bioassays are based solely on the
lethal concentration required to inhibit seed
germination. More often, bioassays are based on
sublethal concentrations that inhibit root elonga-
tion in which the response is dose-related
(Horowitz 1976). The basis for root elongation
tests can be seen in specific herbicidal studies.
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o or example, the effects of pic.loram on germination
and root elongation in safflower, radish, and
barley were compared by Chang and Foy (1971).
Germination was inhibited at 10,-M whilefiroot
elongation was inhibited at 10 or 10
Similarly, the concentrations of potassium
dichromate required to inhibit germination by 10,0
percent ancUSLO percent of control were 5 x 10
and 1 x 10 , respectively; while a concentration
-5M
of 5 x 10 showed a distinct inhibition of root
elongation (Mukherji and Roy 1977).
o The test measures the inhibition of seed
germination and the stimulation or inhibition of
root growth. Both are easily observed or
measured. In some situations it may not be clear
whether germination or root elongation is
inhibited, since the exact stage at which
germination ends and growth begins is difficult to
define. Some definitions of germination include
the protrusion of the embryo through the seed
coat. For the purpose of this guideline, seeds
with at least a 5 mm protrusion of the embryo are
counted as germinated. Requiring this growth
allows consistent identification of successful
germination. The slanted substrate technique, by
enhancing linear root growth, facilitates root
measurement.
o The test method is relatively rapid, simple, and
inexpens ive.
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B. Test Procedures
1. Chemical Application
Chemicals have different physical and chemical
properties which will influence the mode of application.
Water-soluble chemicals will not present a problem but other
chemicals will require different approaches. In this
section we have attempted to provide general guidance by
suggesting some approaches that have been used successfully
in the past. There is a need for expert scientific
judgement in the choice of solvent. The Agency recognizes
the need to maintain flexibility, yet ensure that any
effects are due to the test chemical and not the carrier.
The suggested use of carrier controls to distinguish between
test chemical and carrier effects is considered a standard
laboratory practice.
2. Range-Find ing Test
It is recommended that a range-finding test be conducted
prior to the definitive test in those instances where no
information is available on the phytotoxicity of the test
chemical. This approach should reduce the risk of using an
inappropriate concentration series in the definitive test.
Under certain circumstances the range-finding test may
preclude the need to conduct the definitive test. In order
to minimize the cost and time required to obtain the
requisite data, nominal concentrations are permitted, test
duration may be shortened, replicates are not required, and
other test procedures and conditions are relaxed.
If test results indicate that the chemical is non-toxic
or very toxic to plants and if definitive testing is not
conducted, it is necessary to ascertain that at least 65
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percent of the control seed have germinated, the control
root lengths have reached 2.0mra, and that the test was
conducted at the specified incubation temperature. These
verifications establish that the seed tested were viable and
that the test was properly conducted.
In some situations there may be enough information
available on toxicity to select the appropriate
concentrations without a range-finding test. The range-
finding test (or other available information) needs to be
accurate enough to ensure that concentration levels in the
definitive test are spaced above and below the EC10 and EC50
values for germination and root elongation. If the chemical
has no measurable effect at the saturation concentration (at
least 1000 mg/1), it is considered relatively non-toxic to
seed germination/root elongation, and definitive testing for
effects on these processes is deemed unnecessary. In all
cases, the range-finding test is conducted to reduce the
expense involved with having to repeat a definitive test
because of inappropriate test chemical concentrations.
3. Definitive Test
The specific requirements of the definitive test are the
analytical determinations of chemical concentrations, the
unbiased selection of seed for each treatment, the use of
controls, the assessment of test validity, and the
recording, analysis, and presentation of data. These
requirements assure that the chemical concentration - plant
response relationship is accurately known, that chemical
effects are not confounded by differential seed vigor, and
that the relationships are clearly presented. Reporting the
occurrence of such abnormal effects as lesions,
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August, 1982
discoloration, or lack of turgor provides qualitative data
that further assist the assessment of phytotoxicity.
The purpose of the definitive test is to determine the
EC10, EC50, and concentration-response curves for seed
germination and root elongation for each species tested with
a minimum of testing beyond the range-finding test. It is
probable that each of the species tested may have a
different response curve for a given chemical based on the
range-finding test and that more than six concentrations of
a test substance in a geometric series may be needed to
properly describe the dose-response relationships for all
species being tested. By testing a minimum of six
concentrations in a series per species, partial effects
(lack of germination and/or reduced root length) will be
probable and the dose response relationship will be better
defined. The slope and shape of the dose-response curve
will allow estimation of the effects of lower concentrations
on the test plants.
The primary observations - number of seed per species
per chemical dose which germinate, measurement of root
growth, and determination of the actual chemical
concentrations employed in the definitive test - are all
needed to accurately describe the dose response curve from
which the EC10 and EC50 are calculated.
The recommended experimental design is the randomized
complete block. As discussed by Hammer and Urquhart (1979),
it is essential that the investigator randomly assign petri
dishes to treatments to assure that each sample of seed has
the same chance of receiving any of the treatments (exposure
level of test chemical). To account for variation within
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the germination chamber and to increase the sensitivity for
detecting treatment differences, small square blocks should
be delineated in the germination chamber with randomization
of treatments within blocks. Replication should occur over
chambers (of the same type) as, in many cases, a within-
chamber estimate of residual variance badly underestimates
the between-chamber estimate (Hammer and Urquhart 1979).
This means that differences between chambers are often
greater than differences in growth and environmental
conditions within chambers. In the event that blocking
within chambers is impossible, total randomization is
acceptable.
In order to substantiate that temperature was maintained
within specified limits, it will be necessary to measure and
record temperature throughout the test. Requisite
instrumentation is readily available, easy to maintain, and
should not increase complexity or costs of the test.
Temperatures should be recorded hourly to prevent any severe
fluctuations that might affect growth processes and/or
chemical uptake.
4. An al y t ic al Me as ur erne n ts
The actual chemical concentration used in the definitive
test should be determined with the best available analytical
precision. Analysis of stock solutions and test solutions
just prior to use will minimize problems with storage (e.g.,
formation of degradation products, adsorption,
transformation, etc.). Nominal concentra-tions are not
adequate for the purposes of the definitive tests. If
definitive testing is not required because the chemical
elicits an insufficient response at the 1000 mg/1 level in
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the range-finding test, the concentration of chemical in the
test solution should be determined to confirm the actual
exposure level.
The pH of the test solution should be measured prior to
testing to determine it if lies outside of the species'
optimal range. While it is recognized that seeds of crop
plants germinate over a broad range of hydrogen-ion
concentrations and typically exhibit a pH optima for
germination, this test guideline does not include pH
adjustment for the following reasons: the use of acid or
base to adjust pH may chemically alter the test chemical
making it more or less toxic; the amount of acid or base
needed to adjust the pH may vary from one test solution
concentration to the next, and the effect the test chemical
has on pH may indirectly affect the growth and development
of the test plant. Therefore, the pH of each test solution
should be determined and compared with the acceptable range
for growth and development of the test species.
To reduce variability resulting from continual root
elongation during the measurement period, it is recommended
that measurements be made sequentially (by concentration)
within a given species.
The data obtained in bioassays are usually expressed as
standard response curves in which growth response of the
test species is plotted against the concentration of the
test chemical. The manner of expressing plant response
varies considerably. For this guideline, plant growth
responses are expressed as direct measurements of number of
seed that germinate and root growth. The statistical
analysis (goodness-of-fit determination) facilitates
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accurate calculations of EC10 and EC50 as well as providing
confidence limits for the concentration- response curves.
C. Test Conditions
1. Test Species
a. Selection
The ten terrestrial plant species recommended for the
seed germination/root elongation, early seedling growth, and
plant uptake test guidelines are as follows:
Lycopersicon esculentum (tomato)
Cucumis sativus (cucumber)
Lactuca sativa (lettuce)
Glycine max (soybean)
Brassica oleracea (cabbage)
Avena sativa (oat)
Lolium perenne (perennial ryegrass)
Allium cepa (common onion)
Daucus carota (carrot)
Zea mays (corn)
In addition, other species of economic or ecological
importance to the region of impact may also be tested in
lieu of these species.
These ten species have been selected for the following
reasons:
o As food, forage, or ornamentals, they are
economically important and constitute major cash
crops .
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August, 1982
Their distribution, abundance, and taxonomic
representation suggest broad coverage of the plant
kingdom.
They are also specified for phytotoxicity testing
of pesticides (Subpart J, Pesticide Registration
Guidelines). Additional justification for
selection of these test species is provided in
these guidelines (see FR 45(214): 72948-72978).
They are sensitive to many toxic compounds and have
been used to some degree in previous bioassays.
Their use in herbicidal bioassays, heavy metal
screening, salinity and mineral stress tests, and
allelopathic studies indicates a sensitivity to a
wide variety of stressors (Guenzi and McCalla 1966,
Geronimo et al. 1973, Puerner and Seigel 1972,
Wiedman and Appleby 1972, Reynolds 1978, Chang and
Foy 1971).
They are compatible with the environmental growth
conditions and time constraints of the test
method. Seed from the selected species germinate
quickly and easily. Root growth is rapid and
uniform. The seed contain no natural inhibitors
and require no special pretreatment to germinate
(such as soaking, chilling, prewashing, light, or
scarification). Seed size is compatible with the
test system; large enough to be easily manipulated
with forceps but not so large to be space limiting
in the petri dishes. All the seed grow rapidly
under the prescribed environmental conditions and
most have measurable root lengths within a one week
period.
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August, 1982
Other species may be substituted for any or all of these
ten species when appropriate. For example, forest or desert
species may constitute the population at greatest risk. If
so, those of most value to man or of ecological dominance to
the affected ecosystem should be selected. The rationale
for selection of alternative species should be discussed
with the Agency and/or supported in the report of findings.
No single plant will always be the most or least
sensitive to all chemicals which may be tested. The use of
different types of plants ensures that variations in plant
response will be evident. In a seedling growth test, Hikino
(1978) used concentrations of 0.01 to 1,000 ppm for eight
chemicals. Rice, turnip, and soybean seed were placed in
petri dishes with test chemical and agar medium and
incubated in the dark. Root and shoot.development were
measured each day and at the end of the test period. Six of
the eight chemicals inhibited root growth at 100 or 1,000
ppm for each species. The other two chemicals inhibited
root growth in at least one of the species. It is important
to note that none of the three species tested was
consistently the "most sensitive". These results further
support the requirements of testing several species.
The definitive test requires that seed of the same size
class be used throughout the test. Germination rate and
percentage, seedling vigor, initial growth rate, and
sensitivity to chemical stress are related to seed size.
For example, Anderson (1969) found a significant correlation
between soybean seed size and sensivitity to the herbicide
atrazine, with the smaller seed being more sensitive than
the larger seed. Also, the larger or heavier the seed, the
12
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August, 1982
greater the percent germination and the more vigorous the
seedling (Pollock and Roos 1972, Smith et al. 1973, Sharpies
1970, Whalley et al. 1966, Carleton and Cooper 1972). The
differential response to chemical stress suggests that seed
of the smallest size class be used for testing to take
advantage of the increased sensitivity. However, as use of
seed from the smallest size class carries with it a
reduction in viability and a probable increase in
variability, it is recommended that larger seed be used. By
selecting seed from the size class containing the most seed,
assurance is provided that sufficient seed are available for
testing, that the percent viability is sufficiently high for
valid testing, and that seed sensitivity to the test
chemical is representative of the species.
Information provided by the seed supplier provides
additional assurance that the seed are viable. Use of seed
produced during one growing season minimizes problems
associated with differential viability between lots. By
using untreated seed, possibilities of confounding test
results with fungicides, repellants, etc., are eliminated.
2. Facilities
a. Apparatus
The test requires a germination chamber or temperature-
controlled enclosure capable of maintaining a uniform
temperature of 25° ± i°c. Other facilities typically needed
for conducting seed germination/root elongation tests
include standard laboratory glassware, petri dishes, work
areas to clean and prepare equipment and to measure chemical
concentrations and plant responses, refrigeration to hold
the seed until needed for testing, and proper disposal
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facilities. Without these facilities, the testing cannot be
adequately conducted.
b. Containers and Support Substrate
Petri dishes should be composed of inert materials such
as borosilicate glass in order to minimize reaction of the
test chemical and/or carrier with the container. The growth
support substrate should also be composed of inert material
for these same reasons. Quartz sand and glass beads are
suitable inert materials and minimally sorb substances,
ensuring that the chemical will be maximally available to
the inbibing seed and developing embryo. The chemical would
not be as readily available if it were sorbed to the
substrate or container. Sand or glass beads are used,
rather than vermiculite, perlite, or soil, to avoid
complications associated with variable physical and chemical
properties and microbial populations indigenous to native
soils. Native soils are undesirable because of the varying
clay, sand, and humus components, the types and proportions
of which vary within the same soil type. Microbial
populations also vary between soil types. These variables
alter moisture-holding capacity, chemical-binding capacity,
aeration, and nutrient and trace element content (Audus
1964, Beetsman et al. 1969, Beall and Nash 1969). In
addition to the variations in these physical factors, there
will also be variation in such chemical properties as pH and
red ox potential. Because of the impossibility of
controlling physical and chemical properties of native
soils, inert material is required to support the plants with
the only variables being the presence and concentration of
test chemical. The purpose of using glass beads or sand
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August, 1982
instead of native soils is not to make test results more
directly applicable to natural systems, rather it is to
eliminate sources of variation in the test.
c. Cleaning and Sterilization
Standard good laboratory practices are recommended to
remove dust, dirt, other debris, and organic and inorganic
residues from the test containers and support media which
might confound test results. Residues could enter the test
solution and be taken up by the developing embryos,
affecting their growth and/or their metabolic activity,
causing misleading plant response. Bichromate solution
should not be used for washing glassware or the glass beads
since dichromate inhibits germination and reduces root
elongation (Mukherji and Roy 1977). Sand or plastic petri
dishes should not be reused to insure against residue carry-
over.
Since untreated seed are used for all testing, fungal
contamination during testing may reduce germination and root
growth. If fungal contamination is a problem, the testing
laboratory should evaluate glassware cleaning procedures,
laboratory ventilation, and other possible contributory
sources and correct them first. If these actions do not
control the problem, then the seed sterilization procedure
described in the test guideline or another appropriate
technique should be utilized. The sterilization procedure
selected, however, should not bias testing results.
3. Environmental Conditions
Controlled environmental conditions are necessary to
maintain uniform growth and ensure reliable data. The
essential environmental factors for germination are an
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August, 1982
adequate supply of water, a suitable temperature, an
adequate supply of oxygen, and in some species, light.
Temperature is the one single variable that could most
readily affect metabolic respiration and thereby growth rate
and chemical effect. The temperature range suitable for
germination is quite broad for most species. Within this
range there is usually an optimal temperature at which the
highest percentage of germination is attained in the
shortest time. A single temperature was selected which was
near the optimal range for germination and root elongation
of all test species and commonly used in seed viability
s tudies .
The seed of the selected species will germinate in
continuous dark. Incubators, seed germinators, or
controlled environment chambers that provide temperature
control are available commercially. Without the need for
controlled light intensity or photoperiod, a constant
temperature is easily maiatained at 25 +_ 1°C and cost is
reduced compared with costs of lighted chambers.
Both seed germination and root elongation are dependent
upon an adequate supply of water. Adequate uniform moisture
for germination and root elongation is provided by
maintaining saturation of the growth substrate (quartz sand,
or glass beads) with test solution in petri dishes. Sealing
the petri dishes with tape or enclosing them in plastic bags
reduces the rate of water loss.
Both seed germination and root elongation depend upon
adequate oxygen. An adequate supply of air may not be
present to allow normal growth and development in petri
dishes if they are completely sealed. It may be necessary
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to leave the petri dish lid partially unsealed in order to
provide sufficient oxygen to support respiration.
D. Reporting
The sponsor should submit to the Agency all data
developed during the test which are suggestive or predictive
of phytotoxicity. In addition, information supplied with
the seed and the size class of seed used for testing are
required because these data have a bearing on the validity
of the test. If testing specifications are followed, the
sponsor should report that specified procedures were
followed and present the results. If alternative procedures
were used instead of those recommended in the test
guideline, then the protocol used should be fully described
and justified.
Test temperature, chemical concentrations, test data,
concentration response curves, and statistical analyses
should all be reported. The justification for this body of
information is contained in this support document. If plant
species other than the ten recommended were used, the
rationale for the selection of the other species should be
provided.
III. Economic Aspects
The Agency awarded a contract to Enviro Control, Inc. to
provide us with an estimate of the cost for performing an
early seedling growth test according to this Guideline.
Enviro Control supplied us with two estimates; a protocol
estimate and a laboratory survey estimate.
The protocol estimate was $1,687. This estimate was
prepared by identifying the major tasks needed to do a test
and estimating the hours to accomplish each task.
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Appropriate hourly rates were then applied to yield a total
direct labor charge. An estimated average overhead rate of
115%, other direct costs of $200, a general and
administrative rate of 10%, and a fee of 20% were then added
to the direct labor charge to yield the final estimate.
Enviro Control estimated that differences in salaries,
equipment, overhead costs and other factors between
laboratories could result in as much as 50% variation from
this estimate. Consequently, they estimated that test costs
could range from $843 to $2,531.
The laboratory survey estimate was $950, the mean of the
estimates received from two laboratories. The estimates
ranged from $700 to $1,200 and were based on the costs to
perform the test according to this Guideline.
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IV. References
Anderson RN. 1969. A search for atrazine resistance in
soybeans. Weed Sci. Soc. Amer. Abstr. No. 157.
Asplund R. 1969. Some quantitative aspects of the
phytotoxicity of monoterpenes. Weed Sci. 17:454-455.
Audus LJ. 1964. Herbicide behavior in the soil. II.
Interactions with soil microorganisms. In: The
physiology and biochemistry of herbicides. New York:
Academic Press, pps. 168-206.
Beall ML Jr and Nash RG. 1969. Crop seedling uptake of
DDT, dieldrin, endrin, and heptachlor from soil. Agron.
J. 61:571-575.
Beetsman GD, Kenney DR, Chesters G. 1969. Dieldrin
uptake by corn as affected by soil properties. Agron.
J. 61:247-250.
Behreas R. 1970. Quantitative determination of
triazine herbicides in soils by bioassay. Residue Rev.
32:355-369.
Berlyn GP. 1972. Seed germination and morphogenesis.
In: Kozlowski TT, ed. Seed Biology. Vol. I. New
York: Academic Press, pp. 223-312.
Bode HR. 1958. Beitrage zur Keuntnis allelopathischer
Erschwinurgen bei inigen Juglandaceen. Planta 51:440-
480.
Brown AWA. 1978. Ecology of pesticides. New York: John
Wiley and Sons, pp. 320-343.
Carleton AE and Cooper CS. 1972. Seed size effects upon
seedling vigor for three forage legumes. Crop Sci.
12:183-186.
Chang I, and Foy CL. 1971. Effect of picloram on
germination and seedling development of four species.
Weed Sci. 19:58-64.
Durrant MJ, Draycott AP, Payne PA. 1974. Some effects
of NaCl on germination and seedling growth of sugar
beet. Ann. Bot. 38:1045-1051.
19
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Garb S. 1961. Differential growth inhibitors produced
by plants. Bot. Rev. 27:422-443.
Geronimo J, Smith LL Jr., Stockdale GD, Goring CAI.
1973. Comparative toxicity of nitrapyrin and its
principle metabolite, 6-chloropicolinic acid. Agron. J.
65:689-692.
Guenzi WD and McCalla TM. 1966. Phenolic acids in
oats, wheat, sorghum and corn residues and their
phytotoxicity. Agron. J. 58: 303-304.
Hammer PA and Urquhart NS. 1979. Precision and
replication: Critique II. In: Tibbitts TW and Kozlowski
TT, eds. Controlled environment guidelines for plant
research. New York: Academic Press, pps. 364-368.
Hikino H. 1978. Study on the development of the test
methods for evaluation of the effects of chemicals on
plants. Chemical Research Report No. 4. Tokyo, Japan:
Office of Health Studies, Environmental Agency Japan.
Horowitz M. 1966. A rapid bioassay for PEBC and its
application in volatilization and adsorption studies.
Weed Res. 6:22-36.
. 1976. Application of bioassay techniques
to herbicide investigations. Weed Res. 16:209-215.
and Hulin N. 1971. A rapid bioassay for
diphenamid and its application in soil studies. Weed
Res. 11:143-149.
Imai I, and Siegel SM. 1973. A specific response to
toxic cadmium levels in kidney bean embryos. Physiol.
Plant. 29:118-120.
Konzak CF, Polle E, Kittrick JA. 1976. Screening
several crops for aluminum tolerance. In: Proc. of
workshop on plant adaptation to mineral stress in
problem soils, Beltsville, MD. Ithaca, New York:
Cornell University Press, pp. 311-327.
Kratky BA and Warren GF. 1971. The use of three
simple, rapid bioassays on forty-two herbicides. Weed
Res. 11:257-262.
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Mayer AM and Pol jakoff -Maybe r A. 1975. The germination
of seeds. 2nd ed . Oxford: Pergamon Press.
Muller WH. 1965. Volatile materials produced by Sal via
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bacteria. Bot. Gaz. 126:195-200.
Mukherji S and Roy BK. 1977. Toxic effects of chromium
on germinating seeds and potato tuber slices. Biochem.
Physiol. Pflanzen. 171:235-238.
Neiman RH and Poulsen LL. 1971. Plant growth
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Bot. Gaz. 132:14-19.
Parker C. 1964. Methods for the rapid bioassay of
herbicides. Proc. Brit. Weed Control Conf . 7:899-902.
Pollock BM and Roos EE. 1972. Seed and seedling vigor.
In: Kozlowski TT, ed. Seed biology. Vol. I. New York:
Academic Press, pp. 313-387.
Puerner NJ and Siegel SM. 1972. The effects of mercury
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Reynolds T. 1978. Comparative effects of aromatic
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Ann. Bot. 42:419-427.
Rubinstein R, Cuirle E, Cole H. 1975. Test methods for
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Santelmann PW. 1972. Herbicide bioassay. In: Truelove
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Smith DW, Welch NC, Little TM. 1973. Studies on
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Whalley RDB, McKell CM, Green LR. 1966. Seedling vigor
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BG-13
August, 1982
EARLY SEEDLING GROWTH TOXICITY TEST
OFFICE OF TOXIC SUBSTANCES
OFFICE OF PESTICIDES AND TOXIC SUBSTANCES
U.S.' ENVIRONMENTAL PROTECTION AGENCY
WASHINGTON, D.C. 20460
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Office of Toxic Substances EG-13
Guideline for Testing Chemicals August, 1982
EARLY SEEDLING GROWTH TEST
(a) Purpose. The guideline in this section is intended for
use in developing data on the toxicity of chemical substances and
mixtures ("chemicals") subject to environmental effects test
regulations under the Toxic Substances Control Act (TSCA) (Pub.L.
94-469, 90 Stat. 2003, 14 U.S.C. 2601 et. seq.). This guideline
prescribes tests using commercially important terrestrial plants
to develop data on the phytotoxicity of chemicals. The United
States Environmental Protection Agency (USEPA) will use data from
these tests in assessing the hazard of a chemical to the
environment.
(b) Definitions . The definitions in section 3 of the Toxic
Substances Control Act (TSCA), and Part 792—Good Laboratory
Practice Standards apply to this test guideline. The following
definitions also apply to this test guideline.
(1) "EC X" means the experimentally derived chemical
concentration that is calculated to effect X percent of the test
criterion.
(2) "Germination" means the resumption of active growth by
an embryo.
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BG-13
August, 1982
(3) "Support media" meant the quartz sand or glass beads
used to support the plant.
(c) Test Procedures — (1) Summary of the Test—(i) Root
exposure. In preparation for the test, seeds are planted in the
potting containers (or in cotton or glass-wool plugs supported in
hydroponic solution) and after germination seedlings are thinned
by pinching the stem at the support medium surface to the ten
(10) most uniform seedlings per pot. This marks the start of the
test and the time of first application of test chemical.
Seedlings emerging after this time are also pinched off at the
surface. Potting mixtures of sand or glass beads are
subirrigated with nutrient solution. Chemicals are applied to
the plants via nutrient solution or are adsorbed to the support
media. Plants are harvested after 14 days and analyzed for
growth.
(ii) Foliar exposure. The foliar exposure test is identical
to the root exposure test except that chemicals are applied to
plants by either spraying or dusting the foliage or by exposing
the plants to gas in a fumigation chamber.
(2) Chemical application—(i) Root exposure. (A)
Chemicals that are soluble in water should be dissolved in the
nutrient solution just prior to the beginning of the test.
Deionized or glass-distilled water should be used in making stock
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BG-13
August, 1982
solutions of the test chemical. Sufficient quantities of each
concentration should be made up as needed to minimize storage
time and disposal volume.
(B) Chemicals that are insoluble in water, but which can be
suspended in an aqueous solution by a carrier, should be added,
with the carrier, to the nutrient solution. The carrier should
be soluble in water, relatively non-toxic to plants, and should
be used in the minimum amount required to dissolve or suspend the
test chemical. There are no preferred carriers; however,
acetone, gum arabic, polyethylene glycol, ethanol, and others
have extensively been used in testing herbicides, plant growth
regulators, fungicides, and other chemicals that affect plants.
Carrier controls should be included in the experimental design of
the test and tested simultaneously.
(C) Water-insoluble chemicals for which no non-toxic, water-
soluble carrier is available, should be dissolved in an
appropriate volatile solvent. The solution should be mixed with
the sand or glass beads which are then placed in a rotary vacuum
apparatus and evaporated leaving a uniform coating of chemical on
the sand or beads. A weighed portion of beads should be
extracted with the same organic solvent and the chemical assayed
before the potting containers are filled. Solvent controls
should be included in the experimental design and tested
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August, 1982
s imultaneously.
(ii) Foliar exposure. (A) Water-soluble chemicals should
be dissolved in deionized or glass-distilled water just prior to
use. Sufficient quantities of each concentration should be made
up as needed. These solutions should be applied daily (during
the normal five-day work week). Plants should be placed in an
exhaust hood and the chemical applied to the foliage. A plastic
sleeve may be fitted over the top of the pot, and the foliage
sprayed with specific quantities of test solution at known
concentrations. The plastic sleeve, confining the chemical to
plant and pot, facilitates expression of chemical dosage to
quantity per pot area (i.e., ug/m^). Shoots of control plants
should be sprayed in an identical manner with deionized or
distilled water. Alternatively, a miniature compressed-air
sprayer mounted on a pendulum and equipped to automatically spray
a plant positioned directly beneath the center of its arc of
swing may be used.
(B) Water-insoluble chemicals, existing as solids, may be
prepared for testing by grinding or other reduction to particles
of <200 urn diameter. Each day (during the normal five-day work
week) plants should be placed in an exhaust hood, a plastic
sleeve fitted over the top of the pot, and specific quantities of
chemical sprinkled uniformly over the potted seedlings. Prior to
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August, 1982
chemical application, plants should be misted with water to
promote foliar retention of the chemical. Control plants should
also be misted with deionized or distilled water at each
treatment date and dusted with an inert material of the same
particle size. Applications are expressed as quantity per unit
pot area (i.e., ug/m2).
(C) Chemicals existing in gaseous form at normal ambient
temperatures and pressures can be generated as needed or stored
under pressure. The bottled gas may be 100 percent chemical or
may be mixed with an inert carrier, such as nitrogen, to known
concentrations. Chemicals of controlled or measured
concentrations should be metered into the exposure chamber,
uniformly mixed about the plants, and exhausted through an outlet
port.
(3) Range-finding test. (i) A range-finding test should be
conducted to establish (A) if definitive testing is necessary and
(B) the concentrations of test substance used in the definitive
test for each species.
(ii) The recommended procedure is to expose newly germinated
seedlings to a series of widely spaced concentrations of test
chemical and assess effect as growth reduction. Seeds
(approximately 30) should be planted directly in containers
filled to within 2.5 cm of the top with quartz sand or glass
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August, 1982
beads. If a hydroponic system is used, the seeds should be
planted in plugs of cotton or glass wool supported at the top of
the solution. When 50 percent of the seeds have germinated the
seedlings should be thinned (by pinching) to the 10 most uniform
per pot and exposed to a widely spaced concentration series
(i.e., 0.01, 0.1, 1.0, 10, 100, 1,000 rag/1) of test chemical.
The lowest concentration in the series, exclusive of controls,
should be at the chemical's detection limit. The upper
concentration, for water-soluble compounds, should be the
saturation concentration. If the anticipated fate of the
chemical is soil or soil water, and the mechanism of concern is
root uptake, the chemical should be applied in nutrient solution
to the root support media (or coated on sand or glass beads for
non-water soluble chemicals). With a chemical whose anticipated
mode of exposure to plants is surface deposition by atmospheric
transport, or irrigation water, the appropriate testing method
may be foliar application allowing subsequent movement into the
rooting zone with watering. Effect is assessed as growth
reduction.
(iii) Alternatively, the seed germination/root elongation
test may be used to establish the appropriate concentration range
for tes ting .
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August, 1982
(iv) No replicates are required and nominal concentrations
are acceptable unless definitive testing is not required.
(v) Definitive testing is not necessary if the highest
chemical concentration tested results in less than a 50 percent
reduction in growth or if the lowest concentration tested
(analytical detection limit) results in greater than a 50 percent
reduction in growth.
(4) Definitive test. (i) The purpose of the definitive test
is to determine the concentration response curves and the EC 10's
and EC 50's for each of the species tested with the minimum
amount of testing beyond the range-finding test.
(ii) At least 5 concentrations of chemical, exclusive of
controls, should be used in the definitive test. For each
species tested the concentration range should be selected to
define the concentration-response curve between the EC 10 and EC
90. Test chemicals should be added to the hydroponic or nutrient
solution or coated on the support media for the root exposure
test; or sprayed, dusted, or gassed directly on the foliage in
the foliage exposure tests.
(iii) Control pots should be included in the experimental
design and should be used in each run. In addition, a carrier
control should also be used for those chemicals that need to be
solubilized.
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August, 1982
(iv) If plants are to be grown hydroponically, seeds should
be planted in plugs of cotton or glass wool supported in the tops
of the containers. When sand or glass beads are used, the
recommended planting procedure is to fill the potting containers
to within 2.5 cm of the top and to sow seeds directly on the
support media. After 50 percent of the seeds have germinated,
the seedlings should be thinned to the 10 most uniform per pot.
(v) Alternate planting methods may be required when the
chemical is highly volatile. An impervious barrier of
polyethylene film, a modification of the double pot method, a
glass plate, or other appropriate apparatus should be used to
prevent volatilization from the root zone. Seeds should be
germinated in the dark at 25°C and seedlings with radicle lengths
in the median range transplanted into the potting containers.
The seedlings should be positioned such that their roots are
exposed to the support media while the shoots pass through holes
in the barrier. A ring of non-toxic, inert, pliable putty should
be used to seal the holes around the stems. Control pots should
be handled identically to the test pots except there is no
exposure to the test chemical. This transplanting procedure,
without the volatilization barrier, is also recommended when the
test chemical is adsorbed to the support medium.
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EG-13
August, 1982
(vi) The test consists of one run for each of the
recommended plant species or selected alternates. The duration
of a run should be at least 14 days from the time that 50 percent
of the seeds have germinated. For a particular chemical, a run
is defined as exposure of the plant species to five
concentrations of the chemical in a minimum of 3 replicate pots
(10 plants per pot), with appropriate controls, followed by
weight and height determinations and analysis.
(vii) All abnormalities (visible effects of the chemicals on
plant growth and morphology including stunting of growth,
discoloration, chlorosis and/or necrosis of the leaves, or
morphological abnormalities) should be recorded. Observations
plants should be made daily (during the normal five-day work
we ek) .
(viii) A randomized complete block design is recommended for
this test with blocks delineated within the chambers or over
greenhouse benches and randomization of treatment occurring
within the blocks. If, because of very large pots, there exists
inadequate space within chambers for blocking, total
randomization within chambers is acceptable.
( ix) Irradiation measurements should be taken at the top of
the plant canopy and the mean, plus a maximum and a minimum
value, determined over the plant-growing area. These
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EG-13
August, 1982
measurements should be taken daily and should be taken at least
at the start and finish of the test. If the test is conducted in
a greenhouse facility, hourly measurements of irradiation should
be recorded and presented as daily total irradiance plus
representative hourly curves for clear sky conditions and cloudy
days .
(x) Temperature and humidity should be measured daily at the
top of the plant canopy during each light and dark period.
(xi) Measurements of carbon dioxide concentration should be
made at the top of the plant canopy (of chamber- grown plants) on
a "continuous basis".
(5) [Reserved]
(6) Analytical measurements — (i) Chemical. Stock solutions
should be diluted with glass distilled or deionized water to
obtain the test solutions. Standard analytical methods, if
available, should be used to establish concentrations of these
solutions and should be validated before beginning the test. An
analytical method is not acceptable if likely degradation
products of the chemical, such as hydrolysis and oxidation
products, give positive or negative interference. The pH of
these solutions should also be measured prior to use.
(ii) Numerical. Mass and length of roots, shoots, and
entire plants (root and shoot) should be measured for the
10
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EG-13
August, 1982
definitive test. Means and standard deviations should be
calculated and plotted for each treatment and control.
Appropriate statistical analyses should provide a goodness-of-fit
determination for the concentration-response curves.
(d) Test conditions — (1) Test Species — (i) Selection. (A)
Test plants recommended for the definitive test include:
Lycopersicon esculentum (tomato)
Cucumis sativus (cucumber)
Lactuca sativa (lettuce)
Glycine max^ (soybean)
Brassica oleracea (cabbage)
Avena sativa (oat)
Lolium perenne (perennial ryegrass)
Allium cepa (common onion)
Daucus carota (carrot)
Zea mays (corn)
(B) Other species, of economic or ecologic importance to the
region of impact, may also be appropriate and selected for
testing .
(ii) Seed selection. Information on seed lot, the seed year
or growing season collected and germination percentage should be
provided by the source of the seed. Only untreated seed (not
treated with fungicides, repellants, etc.) taken from the same
11
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EG-13
August, 1982
lot, and year or season of collection should be used in a given
test. In addition, all seed of a species used in a test should
be of the same size class; and that size class which contains the
most seed should be selected and used in a given test. Any seed
which is damaged should be discarded.
(2) Facilities--(i) Apparatus. (A) Greenhouses or
environmental chambers should provide adequate environmental
controls to meet the carbon dioxide, humidity, irradiation,
photoperiod, and temperature specifications. Chambers should be
designed to prevent escape of internal air into the external
environment other than through appropriate filtering material or
media to prevent contamination of the external environment with
the test chemical.
(B) Laboratory facilities for chemical determinations should
include non-porous floor covering, absorbant bench covering with
non-porous backing, and adequate disposal facilities to
accommodate plant nutrient, test and wash solutions containing
test chemicals at the end of each run, and any bench covering,
lab clothing, or other contaminated materials.
(ii) Containers and support media. For each run, 18
polyethylene pots sufficiently large to grow at least 10 plants
up to 14 days, are required for each species. It is equally
acceptable to use small, individual containers if plants are
12
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BG-13
August, 1982
grown in hydroponic solution. An additional 3 pots will be
needed if a carrier control is needed. Potting containers used
in each experiment should be of equal size and volume and possess
the same configuration. When sand or glass beads are used, the
potting containers should be filled to within 2.5 cm of their
tops. Perlite, vermiculite, native soils, etc., should not be
used for root support.
(iii) Cleaning and sterilization. (A) Potting and receiving
containers, nutrient storage containers, and root support medium
should be cleaned before use. All equipment should be washed
according to good standard laboratory procedures to remove any
residues remaining from manufacturing or prior use. A dichromate
solution should not be used for cleaning beads or pots.
(B) Rooting media other than glass beads should be discarded
at the end of the experiment. Disposal should conform to
existing regulations.
(iv) Nutrient medija. Half-strength modified Hoagland
nutrient solution should be utilized as nutrient media for this
test. When sand or glass beads are used as a support media, the
potting containers should be filled with nutrient solution and
drained periodically. An automated system design is recommended.
(3) Test parameters. Environmental conditions should be
maintained as specified below:
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EG-13
August, 1982
(i) Carbon dioxide concentration at 350 ± 50 ppm.
(ii) Relative humidity should approach 70 ± 5 percent during
light periods and 90 percent during dark periods.
(iii) Irradiation, measured at 1 meter from the source, at
350±50 uE/m2 sec at 400-700 ran.
(iv) Photoperiods of 16 hours light and 8 hours darkness.
(v) Day/night temperatures at 25°/20° ± 3°C.
(e) Reporting. Reporting requirements of Part 792--Good
Laboratory Practice Standards apply to this guideline. The
following data should be reported for each of the species tested
in tabular form:
(i) Concentration of chemical in nutrient solution and in
the root support material when the chemical is soluble in water
or solubilized with a carrier compound; or the concentration of
carrier compound in nutrient solution when carrier is used; or
the quantity of chemical per unit weight of root support material
when it is coated on the material.
(ii) The quantity of chemical, the concentration at <<--hich it
was applied, and the number of applications for those chemicals
applied to the foliage.
(iii) Environmental conditions (day/night temperatures,
relative humidity, light intensity, carbon dioxide concentration,
and photoperiod) .
14
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EG-13
August, 1982
(iv) Mass of above ground (shoot) and below ground (root)
portion of each plant and mass of each whole plant (dry weight at
70°C).
(v) Length of shoot, root, and entire plant.
(vi) Visible effects of chemical, if any, on the intact
plants.
(vii) Means and standard deviations for mass and length of
roots, shoots, and entire plants in each treatment and control.
In addition, concentration- response curves with 95 percent
confidence limits delineated, goodness-of-fit determination, and
EC 10's and EC 50's identified.
15
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ES-10
August, 1982
TECHNICAL SUPPORT DOCUMENT
FOR
EARLY SEEDLING GROWTH TOXICITY TEST
OFFICE OF TOXIC SUBSTANCES
OFFICE OF PESTICIDES AND TOXIC SUBSTANCES
U.S. ENVIRONMENTAL PROTECTION AGENCY
WASHINGTON, D.C. 20460
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TABLE OF CONTENTS
Subject Page
I. Purpose 1
II. ' Scientific Aspects 1
General 1
Test Procedures 3
Chemical Application 3
Range-Finding Test 6
Definitive Test 7
Analytical Measurements 10
Test Conditions 11
Test Species 11
Selection 11
Facilities 14
Apparatus 14
Containers and Support Media 15
Cleaning and Sterilization 16
Nutrient Media 16
Environmental Conditions 18
Carbon Dioxide 19
Irradiation 20
Photoperiod 21
Day/Night Temperatures 22
Relative Humidity 22
Reporting 23
III. Economic Aspects 24
IV. References 25
-------
Office of Toxic Substances ES-10
August, 1982
TECHNICAL SUPPORT DOCUMENT
FOR EARLY SEEDLING GROWTH TOXICITY TEST.
I. Purpose
The purpose of this document is to provide the
scientific background and rationale used in the development
of Test Guideline EG-13 which uses various plant species to
evaluate the toxicity of chemical substances on early
seedling growth in plants. The Document provides an account
of the scientific evidence and an explanation of the logic
used in the selection of the test methodology, procedures
and conditions prescribed in the Test Guideline. Technical
issues and practical considerations are discussed. In
addition, estimates of the cost of conducting the test are
provided.
II. Scientific Aspects
A. General
Chemicals may influence seedling vigor, a characteristic
of increasing importance with increased mechanization in
agriculture. Some crop plants (e.g., lettuce) with intense
culture practices require high germination rates and
vigorous growth. A single seed should germinate and
establish a plant (planting to stand), and all plants should
reach maturity simultaneously for once-over machine
harvesting (Pollock and Roos 1972). Seedling establish-ment
in forests also depends on vigorous root growth to survive
environmental stresses and competition with other plants for
light, water and nutrients. In general, chemicals that
reduce or delay germination and retard maturity of crops
typically result in economic loss.
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ES-10
August, 1982
In agricultural land with other stresses (e.g., drought,
salinity), the additional presence of chemicals which are
toxic may be the limiting factor for production of crops on
that land. In addition, chemicals may enter human food
chains through processes associated with soil/plant
interactions, uptake, translocation, and accumulation in
food and forage crops. In natural systems, affected species
are less competitive and with selection for tolerant species
result in altered species diversity, density, and frequency
of occurrence.
Toxic substances may also cause widely varying and
significant effects on plant community dynamics. Many
chemicals affect plants selectively, with some species
sensitive to and others tolerant of the same chemical and
dose. This selectivity may directly affect the successional
replacement of one plant species by another, either by
hastening the departure of an early successional species or
by inhibiting the establishment of a later stage
successional species (Whittaker 1970). Succession also may
be indirectly affected by inhibition of soil organisms.
Toxic materials may produce high species diversity in some
communities while reducing diversity in others (Brown
1978). Where plant growth and soil organisms are completely
inhibited, soil degradation, instability and eventually
erosion may result.
Early seedling growth was selected to measure
phytotoxicity for the following reasons:
o The early seedling growth phase is a critical stage
in plant development. Occurring immediately after
germination, it constitutes a period of rapid
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ES-10
August, 1982
vegetative growth which is dependent upon such
physiological processes as photosynthesis, water
and nutrient uptake, and metabolic conversions of
photosynthates to structural products (such as cell
walls). Chemical interference with any of these
physiological processes would be evident as reduced
growth. The only physiological processes not
included in this test are the germination of the
seed and the reproductive phases (flowering,
pollination, fruit set).
o The test measures the inhibition of shoot and root
growth. Both are easily observed and measured as
length and as mass.
o The test method is relatively rapid, simple, and
inexpens ive.
The test design is a composite of prior experiments that
investigate the effects of heavy metals, pesticides, or
organic chemicals on plants (Beall and Nash 1971, Cole et
al. 1976, Chou et al. 1978, Fuhr and Mittelstaedt 1980,
Kelly et al. 1980, wickliff et al. 1980). The test is
rapid, simple, and has been successfully used to measure
chemical effects on seedling growth.
B. Test Procedures
1. Chemical Application
Chemicals have different physical and chemical
properties which will influence the mode of application.
Water-soluble chemicals will not present a problem but other
chemicals will require different approaches. In this
section we have attempted to provide general guidance by
suggesting some approaches that have been used successfully
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August, 1982
in the past. There is a need for expert scientific
judgement in the choice of solvent. The Agency recognizes
the need to maintain flexibility, yet insure that any
effects are due to the test chemical and not the carrier.
The suggested use of carrier controls to distinguish between
test chemical and carrier effects is considered a standard
laboratory practice.
Water soluble chemicals are easiest to apply, since
roots may absorb chemicals directly from hydroponic or
nutrient solutions and many air-borne chemicals arrive in
precipitation. Water insoluble chemicals require other
procedures to ensure contact between the test chemical and
the root and shoot systems. Appropriate controls are
necessary and are designed to reduce to the extent possible
the number of variables. One means of incorporating a water
insoluble chemical into the hydroponic solution or the
nutrient medium is to solubilize it in the solution with a
surfactant or carrier compound. When a carrier is used, a
carrier control is included in the procedure to check
carrier's effects on plant growth. A minimum quantity of
carrier should be used to minimize carrier effects.
Foliar applications of powders or sprays should be done
under a hood to ensure worker safety and to minimize growth
chamber contamination. Control plants should also be moved
from chambers to the hood area and sprayed with distilled,
deionized water whenever the chemical is applied to other
plants by spraying because plants are highly sensitive to
mechanical stresses such as handling (Wheeler and Salisbury
1979).
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August, 1982
Attention to maintenance of environmental conditions during
applications of gasses is essential to the success of the
test. Brenann and Leone (1968) observed that plants exposed
to identical concentrations of pollutants at different gas
flow rates exhibited different toxicity symptoms. These
observed variations are understandable when considered in
terms of the effect of environmental physics on mass
transport to the leaves and on the physiological condition
of the plants. In order that each plant in the gas exposure
chamber receive the same dosage, chamber design should
provide a spatially uniform test chemical distribution.
Movement of gaseous substances to the leaf surfaces is
dependent upon air turbulance around the leaves reducing the
boundary layer thickness (Fowler and Unsworth 1974, O'Dell
et al. 1977). Some substances may impact or condense on the
surface of the plants (Fowler and Unsworth 1974) while
others may diffuse through stomatal pores or across the
cuticle to the leaf's interior (Black and Unsworth 1979,
Rich et al. 1970). The cuticle provides an effective
barrier for gaseous substances and several investigators
have shown chemical absorption to be regulated by the same
factors that control stomatal opening including light,
temperature, humidity, carbon dioxide concentration, and the
plant's water status (Bennett et al. 1973, Black and
Unsworth 1979, Rich et al. 1970, Winner and Mooney 1980).
Specified environmental conditions allow for normal stomatal
response and plant growth. These conditions should be
maintained during plant exposure to chemicals. The exposure
chamber designed by Rogers et al. (1977) meets the
environmental criteria and the criteria of uniform gas
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distribution within the chamber.
The growth medium control provides a baseline check of
the effects of hydroponic solution, potting material,
nutrient solution, and the testing environment on plant
growth during each test. This control is simply the plant
growing in the hydroponic solution or root support material
and maintained by the nutrient solution without test
chemical, solvent, or carrier. This control is required in
each run regardless of whether or not the chemical is
soluble in water. It is the only control used when the
chemical is water soluble and applied directly to the root
support media or when it is applied as foliar dust or spray.
A second control is also recommended whenever a carrier
(solubilizer, or non-aqueous solvent) should be used in
order to maximize contact between the chemical and the
plant. The solvent or carrier control is used to check
carrier effects on plant physiology and growth.
2. Range-Finding Test
It is recommended that a range-finding test be conducted
prior to the definitive test in those instances where no
information is available on the phytotoxicity of the test
chemical. This approach should reduce the risk of using an
inappropriate concentration series in the definitive test.
Under certain circumstances the range-finding test may
preclude the need to conduct the definitive test. In order
to minimize the cost and time required to obtain the
requisite data, nominal concentrations are permitted, test
duration may be shortened, replicates are not required and
other test procedures and conditions are relaxed.
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The concentration range for the selected definitive test
is normally based on the results of a range-finding test
that is conducted to reduce the time and expense involved in
having to repeat definitive tests. Results of previously
conducted plant effects tests can be used to this end. For
instance, the seed germination/root elongation test can be
used to establish the appropriate chemical concentration
range for the early seedling growth test. Concentrations
which inhibit seed germination and/or elongation are likely
to be inhibitory to many other metabolic activities of
seedlings and maturing plants. However, for those chemicals
that inhibit photosynthesis, the seed germination/root
elongation test may not provide a suitable range-finding
test.
3. Definitive Test
The specific requirements of the definitive test are the
analytical determinations of chemical concentrations, the
selection of uniformly sized seedlings, the unbiased
selection of seedlings for each treatment, the use of
controls, and the recording, analysis, and presentation of
data. These requirements assure that the chemical
concentration-plant response relationship is accurately
known, that chemical effects are not confounded by
differential seedling vigor, and that the relationships are
clearly presented. Reporting the occurrence of such
abnormalities as stunting, discoloration, chlorosis or
necrosis provides qualitative data that further assist the
assessment of phytotoxicity.
The purpose of the definitive test is to determine the
concentration response curves, EC10, and EC50, for seedling
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growth for each species tested with a minimum of testing
beyond the range-finding test. It is probable that each of
the species tested may have a different response curve for a
given chemical.
The recommended experimental design is the randomized
complete block. As discussed by Hammer and Urquhart (1979),
it is essential that the investigator randomly assign plant
containers to treatments to assure that each plant has the
same chance of receiving any of the treatments (exposure
level of test chemical). To account for variation within
the chamber and to increase the sensitivity for detecting
treatment differences, small square blocks should be
delineated in the growth chamber with randomization of
treatments within blocks. Replication should occur over
chambers (of the same type) as, in many cases, a within-
chamber estimate of residual variance badly underestimates
the between chamber estimate (Hammer and Urquhart 1979).
This means that differences between chambers are often
greater than growth and environmental conditions within
chambers. In the event that blocking within chambers is
impossible, total randomization is acceptable.
In order to substantiate that abiotic test conditions
were maintained within specified limits, it will be
necessary to measure and record irradiation, photoperiod,
temperature, and humidity throughout the test. Requisite
instruments are readily available; easily maintained, and
should not increase complexity or costs of the test. In
fact, these measurements should be made in order to maintain
the specified growing conditions.
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To standardize procedures and provide guidelines for
acceptable data, McFarlane (1979) suggested that radiation
measurements be taken at the top of the plant canopy and
that a mean, plus a maximum and a minimum value, be obtained
over the plant-growing area. These measurements should be
taken daily and should be taken and recorded at least at the
start and finish of the test. If greenhouse facilities are
used, hourly measurements of irradiation should be recorded
and presented as daily total irradiance. Representative
hourly curves of irradiation for clear sky conditions and
cloudy days should also be presented to more fully describe
the light climate.
The photoperiod to which each test organism is exposed
should be reported. If deviations from the recommended
photoperiod occur, they should be recorded so that the
potential impact on plant growth can be assessed.
Day/night temperature measurements should be taken at
least daily at the top of the plant canopy. Data reported
should include the average values for each light and dark
period in the study and the range of variation over the
growing area.
Humidity should be measured at the same time and
location as temperature, and correspondingly reported.
If plants are grown in chambers, it will be necessary to
monitor carbon dioxide concentration in the chamber air to
ensure that excessive depletions do not occur as a result of
rapid photosynthesis rates of the seedlings. These data
should also be reported.
Measurement of solution pH prior to testing is
recommended to determine if it lies outside the species'
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optimal range and contributes to observed effects.
4. Analytical Measurements
The actual chemical concentration used in the definitive
test should be determined with the best available analytical
precision. Analysis of stock solutions and test solutions
just prior to use will minimize problems with storage (e.g.,
formation of degradation products, adsortion, and
transformation). Nominal concentrations are not adequate
for the purposes of the definitive tests. If definitive
testing is not required because the chemical elicits an
insufficient response at the 1000 mg/1 level in the range-
finding test, the concentration of chemical in the test
solution should be determined to confirm the actual exposure
level.
The pH of the test solution should be measured prior to
testing to determine if it lies outside of the species'
optimal range. While it is recognized that plants grow over
a broad range of hydrogen-ion concentration and typically
exhibit a pH optima for growth, this test guideline does not
include pH adjustment for the following reasons: the use of
acid or base may chemically alter the test chemical making
it more or less toxic, the amount of acid or base needed to
adjust the pH may vary from one test solution concentration
to the next, and the effect the test substance has on pH may
indirectly affect the growth and development of the test
plant. Therefore, the pH of each test solution should be
determined and compared with the acceptable range for growth
and development of the test species.
The data obtained in bioassays are usually expressed as
response curves in which growth response of the test species
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is plotted against the concentration of the test chemical.
The manner of expressing plant response varies
considerably. For this guideline, plant growth responses
are expressed as direct measurements of length or mass of
shoots and roots. The statistical analysis (goodness-of-f i t
determination) facilitates accurate determination of the
EC50 as well as providing confidence limits for the
concentration-response curve.
Growth includes both increase in mass (dry weight) and
increase in cell number or size (dimensions). Within a
test, mass and length are frequently closely correlated and
analysis of either produces similar results (Horowitz
1976). Some chemicals, however, induce deformations in
either root or shoot systems. Consequently, it is necessary
to measure length and mass of roots and shoots in order to
adequately assess chemical affects.
C. Test Conditions
1. Test Species
a. Selection
The ten terrestrial plant species recommended for the
seed germination/root elongation, early seedling growth, and
plant uptake test guidelines are as follows:
Lycopersicon esculentum (tomato)
Cucumis sativus (cucumber)
Lactuca sativa (lettuce)
Glycine max (soybean)
Brassica oleracea (cabbage)
Avena sativa (oat)
Lolium perenne (perennial ryegrass)
Allium cepa (common onion)
Daucus carota (carrot)
Zea mays (corn)
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In addition, other species of economic of ecological
importance to the region of impact may also be tested in
lieu of these species.
These ten species have been selected for the following
reasons:
o As food, forage, or ornamentals they are
economically important and constitute major cash
crops .
o Their distribution, abundance and taxonomic
representation suggest broad coverage of the plant
kingdom.
o They are also specified for phytotoxicity testing
of pesticides (Subpart J, Pesticide Registration
Guidelines). Additional justification for
selection of these test species is provided in
these guidelines (see FR 45(214): 72948-72978).
o They are sensitive to many toxic compounds and have
been used to some degree in previous bioassays.
Their use in herbicide bioassays, heavy metal
screening, salinity and mineral stress tests and
allelopathic studies indicates a sensitivity to a
wide variety of stressors (Guenzi and McCalla 1966,
Geronimo et al. 1973, Puerner and Seigel 1972,
Wiedraan and Appleby 1972, Reynolds 1978, Chang and
Foy 1971).
o They are compatible with the environmental growth
conditions and time constraints of the test
method. Seed from the selected species germinate
quickly and easily. Root growth is rapid and
uniform. The seed contain no natural inhibitors
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and require no special pretreatment to germinate
(such as soaking, chilling, prewashing, light, or
scarif ication).
Other species may be substituted for any or all of these
ten species when appropriate. For example, forest or desert
species may constitute the population at greatest risk. If
so, those of most value to man or of ecological dominance to
the affected ecosystem should be selected. The rationale
for selection of alternative species should be discussed
with the Agency and/or supported in the report of findings.
No single plant will always be the most or least
sensitive to all chemicals which may be tested. The use of
different types of plants ensures that variations in plant
responses will be evident. In a seedling growth test,
Hikino (1978) used concentrations of 0.01 to 1,000 ppm for
eight chemicals. Rice, turnip and soybean seed were placed
in petri dishes with test chemical and agar medium and
incubated in the dark. Root and shoot development were
measured each day and at the end of the test period. Six of
the eight chemicals inhibited root growth at 100 or 1,000
ppm for each species. The other two chemicals inhibited
root growth in at least one of the species. It is important
to note that none of the three species tested was
consistently the "most sensitive". These results further
support the requirements of testing several species.
The definitive test requires that seed of the same size
class be used to reduce variability in the size of
seedlings. Generally, the larger or heavier the seed, the
greater the percent germination and the more vigorous the
seedling (Pollock and Roos 1972, Smith et al . 1973, Sharpies
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1970, Whalley et al. 1966, Carleton and Cooper 1972). The
differential response to chemical stress response mandates
uniformity of seed and seedlings in order to minimize
variability. It is recommended that seed be selected from
the size class containing the most seed thus providing
assurance that sufficient seed are available, that the
resulting seedlings will be robust and of uniform size, and
that seedling sensitivity to the test chemical is
representative of the species.
Information provided by the seed supplier provides
additional assurance that the seed are viable. Use of seed
produced during one growing season minimizes problems
associated with differential viability between lots. By
using untreated seed, possibilities of confounding test
results with fungicides, repellants, etc. are eliminated.
2. Facilities
a. Apparatus^
Greenhouse space, growth rooms, or environmental
chambers are equally acceptable as long as the specified
environ-mental conditions are maintained. The environment
affects growth, metabolism, evapotranspiration, and
photosynthesis of plants. Other facilities typically needed
for conducting seedling growth tests include standard
laboratory glassware, a source of distilled water, work
areas to clean and prepare equipment and to measure chemical
concentrations and plant responses, drying ovens,
refrigeration to hold the seed until needed for testing, and
proper disposal facilities. Without these facilities, the
testing cannot be adequately conducted.
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b. Containers and Support Media
Containers for hydroponic solution or root support media
should be composed of inert materials, such as polyethylene,
in order to prevent reaction of test chemicals and/or
carriers with the container. The root support material
should also be composed of inert material for these same
reasons. Quartz sand and glass beads are suitably inert
materials and do not readily adsorb substances. This lack
of adsorption is important because the chemical should be
available for root uptake; it would not be as readily
available if it were adsorbed to the root support material
or container wall.
Sand or glass beads are used, rather than soil, to avoid
complications associated with variable physical and chemical
properties and microbial populations indigenous to native
soils. Native soils are undesirable because of the varying
clay, sand, and humus components, the types and proportions
of which very within the same soil type. Microbial
populations also vary between soil types. These variables
alter moisture holding capacity, chemical binding capacity,
aeration, and nutrient and trace element content (Audus
1964, Beetsman et al. 1969, Beall and Nash 1969). In
addition to the variations in these physical factors, there
will also be variation in such chemical properties as pH and
redox potential. Because of the impossibility of
controlling physical and chemical properties of native
soils, inert material is required to support the plants with
the only variables being the presence and concentration of
test chemical. The purpose of using glass beads or sand
instead of native soils is not to make test results more
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August, 1982
directly applicable to natural systems, rather it is to
eliminate sources of variation in the test.
c. Cleaning and Sterilization
Standard good laboratory practices are recommended to
remove dust, dirt, other debris, and organic and inorganic
residues from the test containers and support media which
might confound test results. Residues could enter the
hydroponic or nutrient solution and be taken up by plants,
affecting their growth and/or other metabolic activity,
resulting in misleading data. A dichromate solution should
not be used for washing containers or beads because
dichromate may enter the nutrient solution, be taken up by
plantss, and affect their growth and metabolic activity.
d. Nutrient Media
The nutrient medium specified in the protocol is
modified half-strength Hoaglund solution (Downs and Helmers
1975). This nutrient solution is recommended because all
the constituent compounds are relatively easy to obtain and
because it works well for the culture of terrestrial plants.
Hydroponic or nutrient culture techniques eliminate
spatial gradients within the growth medium and maintain the
root system at uniform levels of aeration, nutrients, and
water status (Rawlins 1979). The simplest and most
practical method for routine plant growth in controlled
environments is a sand or gravel culture in which thee
plants are grown in containers of sand which are
periodically filled to provide water and nutrients, and
drained, to provide aeration. The frequency of irrigation
depends on the storage capacity of the sand and the rate of
water use by the plant. Use of aerated hydroponic solution
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August, 1982
without a root support media may be preferred by some
laboratories.
Reliability is the primary consideration in the design
of the system which automatically fills and drains the
containers. One of the simplest systems includes a
standpipe connected to an outlet at the bottom of each
container through a manifold. The tops of the containers
are at a uniform elevation. Periodically, the solution
level in the standpipe is raised to the corresponding level
and then lowered below the bottom of the containers. This
system requires only one conduit to each container (for both
filling and draining) and permits variation in the number of
containers. The standpipe is filled by a pump from a lower
storage reservoir. Electrical power to start the pump and
close the drain valve is controlled by a clock timer and,
when the nutrient solution reaches the desired level in the
standpipe, a float switch turns off the power to the pump
and opens the drain allowing the solution to drain back to
the reservoir.
If the rate of uptake for test chemicals applied in the
nutrient solution differs from the rate of nutrient and
water uptake, the exposure concentration will change with
time and differ from that originally specified. In
addition, chemicals applied to foliage as sprays or dust
will enter the rooting zone and accumulate in the nutrient
solution. While foliar application is designed to allow for
chemical movement through, and uptake from, the rooting
zone, excessive concentrations of test chemical should not
be allowed to accumulate in the nutrient solution. The test
solution should be replaced when or if the test chemical
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concentration differs from that specified by ± 20%. When
the chemical is applied to the foliage, the nutrient
solution should be replaced weekly if detectable quantities
move through and accumulate in the solution following
spraying or dusting of foliage.
3. Environmental Conditions
Controlled environmental conditions are necessary to
maintain uniform growth and ensure reliable data.
Maintenance of specified environmental conditions before,
during, and after plant exposure to the chemical is
essential to the successful execution of this test.
Variability of plant response to chemical exposure as a
result of fluctuations in environmental conditions has been
noted (Darwent and Behrens 1972, Dunning and Heck 1973,
Juhren et al. 1957, Leone and Brenann 1970).
Environmental conditions affect growth, metabolism, eva-
potranspiration, and photosynthesis of plants. In addition
to mineral nutrition the conditions that should be
standardized and maintained include: (a) carbon dioxide
concentration; (b) relative humidity; (c) irradiation; (d)
photoperiod; (e) day and night temperatures.
Standardization of environmental conditions is
essential. Several investigators have demonstrated that
differences in environmental conditions influence the
response of plants exposed to chemicals. These include pre-
conditioning by light and humidity (Dunning and Heck 1973),
effects of temperature, photoperiod, and light intensity
during the growth (Juhren et al. 1957), air movement during
exposure (Brenann and Leone 1968), and mineral nutrition
(Leone and Brenann 1970). Environmental conditions between
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August, 1982
growth chambers should be closely maintained, as specified
in this protocol, to ensure common test conditions. If
large growth rooms or greenhouse facilities are used,
comparability of the environment between small groups of
plants is not as critical, and environmental specifications
may be relaxed.
a. Carbon Dioxide Concentration
The carbon dioxide concentration should be high enough
for photosynthesis to occur at a level which will allow
normal plant growth and biomass accumulation.
Photosynthetic rates vary directly with carbon dioxide
partial pressures. In addition, abnormally high levels of
carbon dioxide can affect carbohydrate translocation from
the leaves. Therefore, the concentration of carbon dioxide
in the air surrounding test plants should be kept within
limits conducive to normal plant growth (350 _+_ 50 ppm) . The
carbon dioxide concentration is not expected to be of
concern in greenhouse facilities.
In growth chambers, however, carbon dioxide depletion by
rapidly photosyn thes izing plants may be a real concern. The
extent of depletion is a function of chamber volume, rate of
air exchange between the chambers and the outside, the
number, size, and type of plants, and the growth conditions
maintained in the chambers (Hellmers and Giles 1979, Pallas
1979). Attempts to reduce carbon dioxide depletion by
increasing the air exchange rate between the outside air and
the chamber would probably eliminate temperature and
humidity control within the chambers (Downs and Hellmers
1975). An appropriate system for carbon dioxide control in
up to 5 chambers is described by Hellmers and Giles
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(1979). Alternatively, the use of fewer plants per chamber
may be an appropriate means of avoiding carbon dioxide
depletion problems.
b. Irradiation
The growth chamber irradiation spectrum should be as
similar to natural sunlight as possible in order for test
plants to respond normally. Light drives the photosynthetic
process and specific wavelengths cause particular responses
(e.g., the phytochrome reactions) in plants. Therefore, it
is important that light quality and intensity remain
constant throughout the test. Irradiation in chambers
should be 350 _+_ 50 uE/m2 sec at 400-700 nm measured at the
top of the plant canopy. This corresponds roughly to full,
direct sun plus diffuse radiation. Artificial lighting is
used in growth chambers and as supplemental lighting in
greenhouses. However, lamps may weaken, cease to function,
or function abnormally and fail in the course of long-term
tests. Lamp failures should be corrected as soon as
observed and the changes in light quality and intensity
resulting from the lamp failures should be recorded.
For light quality and intensity to approach that of
natural sunlight, the light source commonly consists of
either a fluorescent-incandescent system, a high intensity
discharge system (HID) composed of metal halide lamps, or a
combination of metal halide and high-pressure sodium
lamps. Combinations of lamps are necessary for artificial
lighting bcause there is no single lamp which is capable of
emanating light of the same quality and intensity as natural
sunlight, although the metal halide lamp closely approaches
ideal lighting conditions.
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August, 1982
HID systems produce intense light that may not be
obtainable in other ways (McFarlane 1978). Because these
lamps emit radiation in discrete line spectra, researchers
frequently combine two different types of HID lamps in
growth chambers to obtain a more balanced spectrum. Metal
halide lamps appear to be the most useful type of HID lamp
because their emission spectra are almost continuous over
the 400 to 700 run wavebands. However, a combination of
metal halide and sodium HID lamps will provide superior
plant growth to either one alone.
A fluorescent-incandescent system should be composed of
70-80 percent input wattage of cool-white fluorescent lamps
and 20-30 percent input wattage of incandescent lamps (Downs
1975). Although incandescent lamps are capable of emanating
light which qualitatively approaches natural sunlight, the
intensity will not approach the 350+50 uE/m2 sec specified
for plant growth. Increasing the number of incandescent
lamps would result in overheating the growth chamber.
Fluorescent lighting alone can approach the intensity of
natural sunlight without overheating the growth chamber,
since very little of the output is in the infrared range
(McFarlane 1978). Incandescent lights should be included to
provide radiation in the red and infrared regions.
Uniformity and intensity of lighting within chambers is
routinely improved by covering the walls with highly
reflective materials.
c. Photoperiod
Plants exhibit three basic photoperiodic responses.
Long-day plants will flower only when the light period is
longer than a certain minimum number of hours in a 24 hour
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day/night cycle. Short-day plants will flower only when the
light period is shorter than a certain maximum number of
hours in a 24 hour day/night cycle. Day-neutral plants will
flower regardless of the length of the light period. In
this test, long day conditions (16 hours light/8 hours
darkness) are recommended in order to maximize biomass.
Continuous light, however, is not recommended as a dark
period is required for the phytochrome interconvers ions
necessary in photosynthetic vascular plants (Salisbury and
Ross 1969).
d . Day/Night Temperature
For any particular plant species, there is an optimal
temperature regime for maximum growth and development. This
regime may differ between the phenophases of germination,
vegetative growth, flowering, and fruit development. The
concept of thermoperiodicity specifies that the temperature
during the light period should be different from that of the
dark period for optimal plant growth and development (Downs
1975). It has been demon-strated that plants grow and
develop better with a day/night temperature differential
(Kramer 1957, Went 1957). While specific temperature optima
can be identified for each species, a regime of 25°/20° _+_
3°C is recommended as it will promote suitable growth and be
cost effective. If growth chambers or rooms are used,
tolerances of +_ 1°C are recommended to ensure environmental
comparability between growth facilities.
e. Relative Humidity
A literature review by Hoffman (1979) indicates that
most plants grow well when the atmospheric saturation
deficit is maintained between 5 and 10 mb. At 25°C, the
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optimum relative humidity range is 65 to 85 percent. Most
environmental control facilities with provision for humidity
control can maintain humidity in this range with an accuracy
at 25°C of _+ 1.0 to 1.6 mb of vapor pressure (+_ 3 to 5
percent relative humidity). Humidity levels have been
observed to affect plant growth and development, rates of
carbon dioxide exchange, flowering, nutrient transport, and
susceptibility to air pollution (Thurtell 1979). However,
these are generally the responses of plants to their total
environments and are not unique to the humidity of the
air. During the daylight period the relative humidity in
the growth chamber should be maintained within the optimal
range of 65 to 85 percent. When several chambers should be
maintained as comparable environments, a relative humidity
of 70 +_ 5 percent is recommended. During the dark period,
relative humidity should approach saturation, as it normally
does in natural environments.
D. Reporting
The sponsor should submit to the Agency all data
developed during the test which are suggestive or predictive
of phytotoxici ty. If. testing specifications are followed,
the sponsor should report that specified procedures were
followed and present the results. If alternative procedures
were used instead of those recommended in the test
guideline, then the protocol used should be fully described
and justified.
Environmental test (growth) conditions, chemical
concentrations, quantity of chemical applied, number of
applications, test data, concentration response curves, and
statistical analyses should all be reported. The
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justification for this body of information is contained in
this support document. If plant species other than the ten
recommended were used, the rationale for the selection of
other species should also be provided.
III. Economic Aspects
The Agency awarded a contract to Enviro Control, Inc. to
provide us with an estimate of the cost for performing an
early seedling growth test according to this guideline.
Enviro Control supplied us with two estimates; a protocol
estimate and a laboratory survey estimate.
The protocol estimate was $1,184. This estimate was
prepared by identifying the major tasks needed to do a test
and estimating the hours to accomplish each task.
Appropriate hourly rates were then applied to yield a total
direct labor charge. An estimated average overhead rate of
115%, other direct costs of $338, a general and
administrative rate of 10%, and a fee of 20% were then added
to the direct labor charge to yield the final estimate.
Enviro Control estimated that differences in salaries,
equipment, overhead costs and other factors between
laboratories could result in as much as 50% variation from
this estimate. Consequently, they estimated that test costs
could range from $592 to $1,776.
The laboratory survey estimate was $12,938, the mean of
the estimates received from two laboratories. The estimates
ranged from $9,600 to $16,000 and were based on the costs to
perform the test according to this Guideline.
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IV. References
Anderson RN. 1969. A search fo-r atrazine resistance in
soybeans. Weed Sci. Soc. Amer. Abstr. No. 157.
Audus LJ. 1964. Herbicide behavior in the soil. II.
Interactions with soil microorganisms. In: The physiology
and biochemistry of herbicides. Chap. 5. New York:
Academic Press, pp. 168-206.
Beall ML Jr. and Nash RG. 1971. Organochlorine insecticide
residues in soybean plant tops: root vs. vapor sorption.
Agron. J. 63:460-464.
Beall ML Jr. and Nash RG. 1969. Crop seedling uptake of
DDT, dieldrin, endrin, and heptachlor from soil. Agron. J.
61:571-575.
Beetsman GD, Kenney DR, Chesters G. 1969. Dieldrin uptake
by corn as affected by soil properties. Agron. J. 61:247-
250.
Behrens R. 1970. Quantitative determination of triazine
herbicides in soils by bioassay. Residue Rev. 32: 355-369.
Bennett JH, Hill C, Gates DM. 1973. A model for gaseous
pollutant sorption by leaves. J. Air Poll. Control Assoc.
23:957-962.
Black VJ and (Jnsworth MH. 1979. Resistance analysis of
sulphur dioxide fluxes to Vicia faba. Nature 282:68-69.
Brenann E and Leone IA. 1968. The response of plants to
sulfur dioxide or ozone-polluted air supplied at varying
flow rates. Phytopathology 58:1661-1669.
Brown AWA. 1978. Ecology of Pesticides. New York: John
Wiley and Sons, pp. 320-343.
Carleton AE and Cooper CS. 1972. Seed size effects upon
seedling vigor or three forage legumes. Crop Sci. 12: 183-
186.
Chou SF, Jacobs LW, Penner D, Tiedje JM. 1978. Absence of
plant uptake and translocation of polybrominated biphenyls
(PBBs). Environ. Health Perspect. 23:9-12.
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Cole LK, Sanborn JR, Me teaIf RL. 1976. Inhibition of corn
growth by aldrin and the insecticide's fate in the soil,
air, crop, and wildlife of a terrestrial model ecosystem.
Environ. Entomol. 5:583-589.
Darwent AL and Behrens R. 1972. Effect of pretreatment
environment on 2,4-D phytotoxicity. Weed Sci. 20:540-544.
Downs RJ. 1975. Controlled Environments for Plant
Research. New York: Columbia University Press.
Downs RJ and Helmers H. 1975. Environment and the
environmental control of plant growth. New York: Academic
Press.
Dunning JA and Heck WW. 1973. Response of pinto bean and
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Fowler D and Unsworth MH. 1974. Dry deposition of sulphur
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Fuhr F and Mittelstaedt W. 1980. Plant experiments on the
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methabenzthiazuron residue from soil. J. Agric. Food
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Hammer PA and Uquhart NS. 1979. Precision and
replication: Critique II. In: Tibbitts TW and Kozlowski
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research. New York: Academic Press, pp. 364-368.
Hellmers H and Giles LJ. 1979. Carbon dioxide: Critique
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Hoffman GL. 1979. Humidity. In: Tibbitts TW and Kozlowski
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Horowitz M. 1976. Application of bioassay techniques to
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2 resistance:
deciduous and
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PLANT UPTAKE AND TRANSLOCATION TEST
OFFICE OF TOXIC SUBSTANCES
OFFICE OF PESTICIDES AND TOXIC SUBSTANCES
U.S. ENVIRONMENTAL PROTECTION AGENCY
WASHINGTON, D.C. 20460
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Office of Toxic Substances EG-14
Guideline for Testing Chemicals August, 1982
PLANT UPTAKE AND TRANSLOCATION TEST
(a) Purpose. The guideline in this section is intended for
use in developing data on the uptake and translocation of
chemical substances and mixtures ("chemicals") by terrestrial
plants subject to environmental effects test regulations under
the Toxic Substances Control Act (TSCA) (Pub.L. 94-469, 90 Stat.
2003, 14 U..S.C. 2601 et. s eg.) . This guideline prescribes tests
using commercially important terrestrial plants to develop data
on the quantity of chemical substances incorporated in plant
tissues and the potential for entry into food chains with
resultant indirect human exposure. The United States
Environmental Protection Agency (USEPA) will use data from these
tests in assessing the hazard of a chemical to the environment.
(b) Definitions. The definitions in section 3 of the
Toxic Substances Control Act (TSCA), and Part 792--Good
Laboratory Practice Standards apply to this test guideline. The
following definitions also apply to this guideline:
(1) "EC X" means the experimentally derived chemical
concentration that is calculated to effect X percent of the test
criterion.
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(2) "Mass balance" means a quantitative accounting of the
distributions of chemical in plant components, support medium,
and test solutions. It also means a quantitative determination
of uptake as the difference between the quantity of gas entering
an exposure chamber, the quantity leaving the chamber, and the
quantity adsorbed to the chamber walls.
(3) "Support media" means the sand or glass beads used to
support the plant.
(4) "Translocation" means the transference or transport of
chemical from the site of uptake to other plant components.
(c) Test procedures — (1) Summary of the test—(i) Root
exposure. In preparation for the test, seeds are planted in the
potting containers (or in cotton or glass-wool plugs supported in
hydroponic solution) and, after germination, seedlings thinned,
by pinching the stem at the support surface. Potting mixtures of
sand or glass beads should be sub-irrigated with nutrient
solution. Chemicals are applied to the plants via nutrient
solution or adsorbed to the support media. Carrot, lettuce,
onion, cabbage, and ryegrass may be harvested whenever there is
adequate plant material for chemical analysis. Cucumber, corn,
soybean, tomato, and oat should be grown until fruit or seed are
mature.
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(ii) Foliar exposure. The foliar exposure test is identical
to the root exposure test except that chemicals are applied to
plants by either spraying or dusting the foliage or exposing the
plants to gas in a fumigation chamber. If plants are fumigated,
either rates of uptake and surface adsorption should be
calculated, or the plants may be harvested and analyzed for test
chemical arid residues.
(2) Chemical Application—(i) Root exposure. (A) Chemicals
that are soluble in water should be dissolved in the nutrient
solution just prior to the beginning of the test. Deionized or
glass distilled water should be used in making stock solutions of
the test chemical. Sufficient quantities of each concentration
should be made up as needed to minimize storage time and disposal
volume.
(B) Chemicals that are insoluble in water but which can be
placed in aqueous suspension with a carrier should be added, with
the carrier, to the nutrient solution. The carrier should be
soluble in water, relatively non-toxic to plants, and should be
used in the minimum amount required to dissolve or suspend the
test chemical. There are no preferred carriers; however,
acetone, gum arable, polyethylene glycol, ethanol, and others
have been used extensively in testing herbicides, plant growth
regulators, fungicides, and other chemicals that affect plants.
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August, 1982
Carrier controls should be included in the experimental design
and tested simultaneously.
(C) Water-insoluble chemicals for which no non-toxic, water-
soluble carrier is available, should be dissolved in an
appropriate volatile solvent. The solution should be mixed with
the sand or glass beads which are then placed in a rotary vacuum
apparatus and evaporated, leaving a uniform coating of chemical
on the sand or beads. A weighed portion of beads should be
extracted with the same organic solvent and the chemical assayed
before the potting containers are filled. Solvent controls
should be included in the experimental design and tested
s imultaneously.
(ii) Foliar exposure. (A) Water soluble chemicals should
be dissolved in deionized or glass distilled water just prior to
use. Sufficient quantities of each concentration should be made
up as needed. These solutions should be applied at weekly
intervals. Plants should be placed in an exhaust hood and the
chemical applied to the foliage. A plastic sleeve may be fitted
over the top of the pot, and the foliage sprayed with specific
quantities of test solution at known concentrations. The plastic
sleeve, confining the chemical to plant and pot, facilitates
expression of chemical dosage as quantity per pot area (i.e.,
ug/m^). Shoots of control plants should be sprayed in an
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August, 1982
identical manner with deionized or distilled water.
Alternatively, a miniature compressed-air sprayer may be mounted
on a pendulum and equipped to automatically spray a plant
positioned directly beneath the center of: its arc of swing. When
radioisotope-labelled chemicals are applied, health and safety
considerations prohibit spray application. Instead, specific
quantities of labelled chemical should be applied directly to
leaves in single drops.
(B) Water-insoluble chemicals, existing as solids, may be
prepared for testing by grinding or other reduction to particles
of <200 urn diameter. These chemicals should be applied at weekly
intervals. Plants should be placed in an exhaust hood, a plastic
sleeve fitted over the top of the pot, and a specific quantity of
chemical sprinkled uniformly over them. Prior to chemical
application, plants should be misted with water to promote foliar
retention of the chemical. Control plants also should be misted
with deionized or distilled water at each treatment date and
dusted with an inert material of the same particle size.
Applications should be expressed as quantity per unit pot area
2
(i.e., ug/m ).
(C) Chemicals existing in gaseous form at normal ambient
temperatures and pressures should be generated for use as needed
or stored under pressure. The bottled gas may be 100 percent
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August, 1982
pure chemical or mixed with an inert carrier, such as nitrogen,
to known concentrations. Chemicals of controlled or measured
concentrations should be metered into the exposure chamber,
uniformly mixed about the plants, and exhausted through the
outlet port where the flow rate and concentration are again
measured. Use of this system design provides an alternate method
of analysis if the quantity of chemical sorbed by plants is less
than that required for chemical analysis. Plants should be
fumigated whenever they have reached sufficient size for
measurement of photosynthesis and transpiration rates, assuming
equivalent detection sensitivity of carbon dioxide, water vapor,
and chemical analyzers. The appropriate size is a function of
the gas exchange system and constitutes an area of expert
judgement.
(3) Range-finding test. (i) A range-finding test should be
conducted to establish the chemical concentrations used in the
uptake and translocation test.
(ii) Because of the different mechanisms involved in root
and leaf uptake, and to more closely define the chemical
concentrations to be used in the uptake test, the definitive
early seedling growth test is recommended as the range-finding
test. Seeds should be germinated directly in containers filled
with sand or glass beads or in cotton or glass-wool plugs
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August, 1982
supported in hydroponic solution. When 50 percent of the
seedlings have germinated, the seedlings should be thinned (by
pinching) to the 10 most uniform per container and exposed to a
concentration series of test chemical. The lowest concentration
in the series, exclusive of controls, should be at or below the
EC 10 while the upper concentration should be at or above the EC
90. If the anticipated fate of the chemical is soil or soil-
water, and the mechanism of concern is root uptake, the chemical
should be applied in nutrient solution to the root support media
(or coated on sand or glass beads for non-water soluble
chemicals). With a chemical whose anticipated mode of exposure
to plants is surface deposition by atmospheric transport or
irrigation water, the appropriate testing method may be foliar
application allowing subsequent movement into the rooting zone
with watering. Effect is assessed as growth reduction. The
concentration selected as the upper limit for the uptake and
translocation test should be near the threshold of visible
injury. Short exposure periods to gas in fumigation chambers are
not expected to promote visible injury or gross reductions in
growth but may alter stomatal resistance, transpiration, or
photosynthesis. Absorption and adsorption rates may be
calculated and gas concentrations for definitive testing selected
based on the calculated sorption rates.
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August, 1982
(iii) Alternatively, the seed germination/root elongation
test or other appropriate phytotoxicity test may be used to
establish the appropriate upper concentration for testing.
(4) Definitive test. (i) The purpose of the uptake and
translocation test is to determine the propensity for a
chemical's accumulation in plants or plant parts.
(ii) At least 3 concentrations of chemical, exclusive of
controls, should be used in the uptake test. Recommended
concentrations would be a descending geometric progression from
the upper concentration tested (i.e. 100, 50, 25 mg/1). A
minimum of 6 replicate pots per concentration, each containing
from one to four seedlings, should be used. If techniques other
than radioisotopes are used to determine uptake, more replicates
may be required to provide sufficient plant materials for
analysis. Test chemicals should be added to the hydroponic or
nutrient solution or coated on glass beads for the root uptake
test; or sprayed, dusted, or gassed directly on the foliage in
the foliage uptake tests. Only untreated seed (not treated with
fungicides, repellants, etc.) taken from the same lot, and year
or season of collection should be used in a given test.
(iii) Control pots should be included in the experimental
design and should be used in each run. In addition, a carrier
control should be used for those chemicals that need to be
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August, 1982
solubilized.
(iv) If plants are to be grown hydroponically, seeds should
be planted in plugs of cotton or glass-wool supported in the tops
of the containers. When sand or glass beads are used, the
recommended planting procedure is to fill potting containers with
sand or glass beads to within 2.5 cm of the top and to sow seeds
directly. After germination, the seedlings should be thinned by
pinching the stem at the support surface. From one to four
seedlings per potting container are required depending on species
tested, the size of the containers, and the size to which the
plants will grow. When plants are grown hydroponically, one
plant per pot will probably be the preferred method. The number
of plants selected should provide sufficient biomass for
analytical procedures. A greater number of plants may be
required depending on species tested, duration of test, and
analytical procedures. Too many plants in a container may
actually reduce the growth and biomass.
(v) Alternate planting methods may be required when the
chemical is highly volatile. An impervious barrier of
polyethylene film, a modification of the double pot method, a
glass plate, or other appropriate apparatus should be used to
prevent volatilization from the root zone. Seeds should be
germinated in the dark at 25°C and seedlings with radicle length
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August, 1982
in the median range transplanted into the potting containers.
The seedlings should be positioned such that their roots are
exposed to the support media while the shoots pass through holes
in the barrier. A ring of inert, non-phytotoxic, pliable putty
should be used to seal the holes around the stems. Control pots
should be handled identically except there is no exposure to the
test chemical. This transplanting procedure, without the
volatilization barrier, is also recommended when the test
chemical is adsorbed to the support medium.
(vi) Hydroponic solutions should be aerated and sand or
glass filled potting containers should be periodically filled
with nutrient solution and drained to provide aeration. For root
exposure tests, the test chemical should be added to the nutrient
solution or directly to substrate. The entire test solution
should be replaced weekly, or earlier if the concentration of
chemical in the test or nutrient solution varies by more than 20
percent of that specified. The volume of solution added should
be recorded .
(vii) The test consists of one run for each of two specified
plant species. The duration of a run, for solid and liquid
chemicals, should be equal to the length of time required for the
particular test variety to achieve sufficient biomass for
testing. The duration of a run for gasseous chemicals
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August, 1982
should the length of time required to make the specified gas
exchange measurements. For a particular chemical, a run is
defined as exposure of the plant species to three concentrations
of test chemical with a minimum of 6 replicate pots and
appropriate: controls. Exposure is followed by extraction and
analysis for parent compound, metabolites, and bound residues in
plant tissues, and in the whole plants for solids, liquids, and
gasses or by calculating rates of absorption and adsorption of
gasses.
(viii) Visible effects (stunting of growth, discoloration,
chlorosis and/or necrosis of the leaves, decreased moisture
content, or morphological abnormalities, etc.) should be
recorded.
( ix) A randomized complete block design is recommended for
this test, with blocks delineated within the chambers or over
greenhouse benches and randomization of treatments occurring
within the blocks. If, because of very large pots and plants,
there exists inadequate space within chambers for blocking, total
randomization within chambers is acceptable. This design is also
appropriate for the growth of plants to be used for foliar
exposure with gas.
(x) Irradiation measurements should be taken at the top of
the plant canopy and the mean, plus a maximum and a minimum
value, determined over the plant-growing area. These
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August, 1982
measurements should be taken at the start of the test, at
biweekly intervals during the test, and at test termination. If
the test is conducted in a greenhouse facility, hourly
measurements of irradiation should be recorded and presented as
daily total irradiance plus representative hourly curves for
clear sky conditions and cloudy days.
(xi) Temperature and humidity measurements should be
measured daily at the top of the plant canopy during each light
and dark period.
(xii) Measurements of carbon dioxide concentration should be
made at the top of the plant canopy (of chamber-growth plants) on
a "continuous" basis.
(xiii) The amount of water and nutrient solution depleted
each week should be recorded, to observe changes in
evapotranspiration rates which may indicate stress. Furthermore,
these data will be used to compute chemical uptake per volume of
water transpired for the uptake test.
(5) [Reserved]
(6) Analytical measurements — (i) Solid or liquid test
chemicals^. (A) Stock solutions should be diluted with glass-
distilled or deionized water to obtain the test solutions.
Standard analytical methods, if available, should be used to
establish concentrations of these solutions and should be
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August, 1982
validated before beginning the test. An analytical method is not
acceptable if likely degradation products of the chemical, such
as hydrolysis and oxidation products, give positive or negative
interference. The pH of these solutions should also be measured
prior to use.
(B) The entire plant should be harvested, rinsed with a
minimum amount of water (which is returned to the nutrient
solution), and separated into its respective organs as follows:
carrot - root peels, peeled roots, and tops; cucumber - fruit,
vines plus leaves, and roots; corn - kernels, husk plus cob,
stalk plus leaves, and roots; lettuce - tops and roots; onion -
bulb and tops; ryegrass - tops and roots; soybean - grain, chaff
plus tops, and roots; oats - grain, chaff plus tops, and roots;
tomato - fruit, vines, and roots; cabbage - head and roots.
Plants from two pots in each treatment may be pooled, giving 3
replicate sample pools per treatment. After the fresh weights of
the plant organs are obtained, each pool of organs should be
subsampled for percent moisture determinations by drying, at 70°C
for 24 hours in a forced-air drying oven, and weighing. Percent
moisture determined from these subsamples is used to correct for
dry weight of the fresh samples which should then be homogenized
and extracted in organic and aqueous solvents. If radioisotopes
are used, the amount of test chemical in each extract should be
13
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August, 1982
determined by liquid or solid scintillation depending on the type
of radiation; otherwise, the amount of chemical should be
determined by standard methods. At test completion, the root
support material should be washed in organic and then aqueous
solvent and analyzed for test chemical before discarding.
(C) A suggested extraction procedure appropriate for many
organic chemicals is as follows: One gram of plant material
should be homogenized with one gram of solvent-washed anhydrous
sodium sulfate in 4 ml of hexane or acetonitrile. The homogenate
should be filtered or centrifuged, the solid residue rinsed with
an appropriate organic solvent, and the filtrate or supernatant
combined with the rinse. The solid residue should be extracted
by sequentially (_!_) homogenizing in water, (2) centrifuging and
decanting the supernatant, (_3_) extracting of the pellet with 6N
hydrochloric acid at 60°C for 10 hours, (_4_) subsequently
digesting with ION potassium hydroxide, and (_5_) combining
supernatants. The resulting solution should be analyzed by
liquid scintillation spectrometry or GLC methodology. The
organic extract should be evaporated under vacuum to a
sufficiently small volume for thin layer chroma tog raphy (TLC) and
co-chromatographed on silica gel plates with known standards of
the parent chemical. If radioisotopes were used, the
chromatographs could be scanned for radioactive substances on a
radiochromatogram scanner. Alternatively, zones may be
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August, 1982
removed from the plates, extracted, and the quantity of chemical
from each zone determined by liquid scintillation spectrometry or
GLC methodology. The unextractable chemical in the remaining
residue may be determined by oxidizing the residue in a complete
combustion oxidizer.
(ii) Ciaseous test chemicals. (A) A gas exposure system
yields requisite data for a direct calculation of uptake rates.
At steady state, chemical uptake may be determined by a mass
balance calculation. Correction for adsorption to surfaces of
the exposure chamber should be made by operating the system
without plants. Pots filled with hydroponic solution or support
media should be included in the system adsorption calibration.
Consequently, chemical analyses of plant tissues exposed to
gaseous chemicals may not be required in order to demonstrate and
quantitate uptake rates.
(B) Altered rates of net photosynthesis, transpiration, and
stomatal conductance are anticipated as a result of chemical
uptake. Rates of these physiological processes before, during,
and after exposure to the gaseous chemical should be determined.
Data required for these calculations are available as a
consequence of maintaining the specified environmental conditions
within the fumigation chamber.
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August, 1982
(iii) Numerical. Mass of pooled plant organs and pooled
whole plants should be measured for the uptake and translocation
test and subjected to chemical analyses (above) to quantify free
parent test chemical, its metabolites and soluble and bound
residues. Mass balance of the test chemical and
evapotranspiration rates of the plants are also determined.
Means and standard deviations should be calculated and plotted
for each of the above for every treatment and control. The data
should also be subjected to an analysis of variance.
(d) Test conditions — (1) Test species. (i) Test plants
recommended for the uptake test include:
Lycopers icon esculentum (toma to)
Cucumis sativus (cucumber)
Lactuca sativa (lettuce)
Glycine max (soybean)
Brass ica oleracea (cabbage)
Avena sativa (oat)
Lolium perenne (perennial ryegrass)
Allium cepa (common onion)
Daucus carota (carrot)
Zea mays (corn)
(ii) Other species of economic or ecologic importance to the
region of impact, may also be appropriate and selected for
16
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August, 1982
testing. Two species of potentially differing sensitivity should
be selected such as a monocotyledonous and a dicotyledonous
species. It is further suggested that the test plants selected
should be of different growth forms, e.g., a root crop and a leaf
crop.
(2) Facilities — (i) Apparatus. Greenhouses, environmental
chambers, or growth rooms should provide adequate environmental
control to meet the carbon dioxide, humidity, irradiation,
photoperiocl, and temperature specifications. Chambers should be
designed to prevent escape of internal air into the external
environment other than through appropriate filtering material or
media to prevent contamination of the external environment with
radioactive and/or test substances. Laboratory facilities for
plant extractions and chemical determinations should include non-
porous floor covering, absorbant bench covering with non-porous
backing, and adequate disposal facilities to accommodate plant
nutrient, test, and wash solutions containing radioisotope and/or
test chemical at the end of each run, and any bench covering, lab
clothing, or other contaminated materials.
(ii) Containers and support media. For testing purposes, at
least 24 polyethylene pots sufficiently large to grow at least 5
plants up to 28 days or one to three plants to maturity are
required. If plants are grown hydroponically, one plant per pot
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August, 1982
may be the preferred method. If a carrier control is needed, 30
pots are used. Potting containers used in each experiment should
be of equal size and volume and possess the same configuration.
When sand or glass beads are used the potting containers should
be filled to within 2.5 cm of their tops with sand or glass
beads. Perlite, verraiculite, native soils, etc., should not be
used for root support. Potting containers should be covered with
opaque polyethylene bags to exclude light and minimize
volatilization of test chemical.
(iii) Cleaning and sterilization. Potting containers,
nutrient storage containers, and root support medium should be
cleaned before use. All equipment should be washed according to
good standard laboratory procedures to remove any residues
remaining from manufacturing or use. A dichromate solution
should not be used for cleaning beads or pots. Rooting media
other than glass beads should be discarded at the end of the
experiment. Disposal should conform to existing regulations.
(iv) Nutrient media. Half-strength modified Hoagland
nutrient solution should be utilized as nutrient media for this
test. Hydroponic solution should be aerated and sand or glass
beads potting containers should be filled with nutrient solution
and drained periodically. An automated system design is
recommended.
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August, 1982
(3) Test parameters. Environmental conditions should be
maintained as specified below:
(i) Carbon dioxide concentrations at 350 ± 50 ppm.
(ii) Relative humidity approaching 70 ± 5 percent during
light periods and 90 percent during dark periods.
(iii) Irradiation, measured at 1 meter from the source, at
350±50 uE/rr.2 sec at 400-700 nm.
(iv) Fhotoperiod of 16 hours light and 8 hours darkness for
all species except soybean which should be provided with 11 hours
light and 13 hours darkness prior to flowering.
(v) Day/night temperatures at 25°/20° ± 3°C.
(e) Reporting . Reporting requirements of Part 792--Good
Laboratory Practice Standards apply to this guideline.
Concentrations should be expressed in appropriate weight units
per grams of dry plant material and of water lost by
evapotranspiration. Data should also include initial and final
total concentration of the test chemical in the growth media.
These data will be used to compute mass balance. The following
should be reported for each of the species tested in tabular
form:
(1) Solid and liquid test chemicals, (i) Concentration of
chemical in nutrient solution and root support material when
chemical is soluble in water or solubilized with a carrier
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August, 1982
compound, as well as the concentration of carrier compound in
nutrient solution when carrier is used, or the quantity of
chemical per unit weight of root support material when it is
coated on the material.
(ii) The quantity of chemical, the concentration at which it
was applied, and the number of applications for those chemicals
applied to the foliage.
(iii) Environmental conditions (day/night temperatures,
relative humidity, light intensity, carbon dioxide concentration,
and photoperiod) and the occurrence and extent of any disruption
of environmental control facilities.
(iv) Mass of each pool of plant organs and by summation, the
mass of whole plants (dry weight after 24 hours at 70°C).
(v) Concentration of free parent test chemical, metabolites
and soluble residues, and bound residues in pooled plant organs
and pooled whole plants.
(vi) Mass balance of chemical.
(vii) Mean evapotranspiration rate per plant.
(viii) Visible effects of chemical, if any, on the intact
plants.
( ix) Analysis of variance, F-test, means, and standard
deviation about the mean are calculated under paragraph (e) (1)
(iv), (v) (vi), and (vii) of this section.
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(2) Gaseous test chemicals. (i) Concentration of gaseous
test chemical at inflow and outflow ports.
(ii) Environmental conditions within gas exposure
system (air temperature, dew point temperature or water vapor
pressure of incoming and outgoing air streams, light intensity,
air speed within chamber, carbon dioxide concentration at inflow
and outflow ports, gas flow rate into and out of exposure
sys tern) .
(iii) Mass (dry weight after 24 hours at 70°C) of leaves and
stems and surface area (one side of leaves) in the exposure
sys tern.
(iv) Calculated measurements of photosynthesis,
transpiration, and stomatal conductance before, during, and after
exposure to test chemical.
(v) Visible effects of chemical, if any, on the plants.
(vi) Analysis of variance, F-test, means, and standard
deviation about the mean are calculated for each of the
following: (A) Steady state rates of photosynthesis,
transpiration, and chemical uptake before, during, and after
fumigation.
(B) Stomatal conductance or leaf diffusion resistance
before, during, and after fumigation.
(vii) If uptake is determined by direct chemical analysis of
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plant tissues, then the reporting requirements also include:
(A) Concentration of free parent test chemical, metabolites and
soluble residues, and bound residues in pooled plant organs and
pooled whole plants.
(B) Mass balance of the chemical.
(C) Analysis of variance, F-test, means and standard
deviation about the mean under paragraph (e)(2)(A) and (B).
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TECHNICAL SUPPORT DOCUMENT
FOR
PLANT UPTAKE AND TRANSLOCATION TEST
OFFICE OF TOXIC SUBSTANCES
OFFICE OF PESTICIDES AND TOXIC SUBSTANCES
U.S. ENVIRONMENTAL PROTECTION AGENCY
WASHINGTON, D.C. 20460
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TABLE OF CONTENTS
Paqe
I. Purpose 1
II. Scientific Aspects 1
General 1
Test Procedures 2
Chemical Application 2
Range-Finding Test 5
Definitive Test 6
Analytical Measurements 9
Test Conditions 13
Test Species 13
Selection 13
Facilities 15
Apparatus 15
Containers and Support Media 15
Cleaning and Sterilization 16
Nutrient Media 17
Environmental Conditions 18
Carbon Dioxide Concentration 19
Relative Humidity 20
Irradiation 21
Photoperiod 23
Day/Night Temperature 23
Reporting 24
III. Economic Aspects 27
IV. References 28
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Office of Toxic Substances ES-11
August, 1982
TECHNICAL SUPPORT DOCUMENT
FOR PLANT UPTAKE AND TRANSLOCATION TEST
I. Purpose
The purpose of this document is to provide the
scientific background and rationale used in the development
of Test Guideline EG-14 which uses various plant species to
evaluate the uptake and translocation of chemical substances
in plants. The Document provides an account of the
scientific evidence and an explanation of the logic used in
the selection of the test methodology, procedures and
conditions prescribed in the Test Guideline. Technical
issues and practical considerations are discussed. In
addition, estimates of the cost of conducting the test are
provided.
11. Scientific Aspects
A. General
Chemicals of concern may be transported to various sites
as gasses or dust, solubilized in precipitation or in
irrigation water, or may be encountered in soils. It is
well known that plants readily take up, translocate,
accumulate, and metabolize chemicals that are nonessential
for plant growth and development. Such uptake and
incorporation often represents the first step in the
transport of these chemicals within terrestrial food webs.
The uptake and translocation test addresses these concerns
of entry into food webs. This test is not concerned with
phytotoxicity; it deals with hazard to animals and the
potential for indirect human exposure.
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The plant uptake test was selected for the following
reasons:
o The incorporation of chemicals in plant tissues, at
concentrations not evidencing visible injury to the
plant, may provide an unrecognized hazard to humans
and other animals.
o The test measures the quantity of chemical adsorbed
to and absorbed by and translocated within
terrestrial plants.
o The test can be used to evaluate the potential for
human exposure to toxic compounds through
terrestrial food webs.
o The data developed, in conjunction with toxicity
data and information on chemical fate and
transport, provide an important segment of the
scientific basis for risk assessment.
The test design is a composite of prior plant
experiments in which uptake of heavy metals, pesticides, or
organic chemicals have been investigated (Beall and Nash
1971, Cole et al. 1976, Chou et al. 1978, Fuhr and
Mittelstaedt 1980, Kelly et al. 1980, Wickliff et al. 1980).
B. Test Procedures
1. Chemical Application
Chemicals have different physical and chemical
properties which will influence the mode of application.
Water-soluble chemicals will not present a problem but other
chemicals will require different approaches. In this
section we have attempted to provide general guidance by
suggesting some approaches that have been used successfully
in the past. There is a need for expert scientific judgment
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in the choice of solvent. The Agency recognizes the need to
maintain flexibility, yet ensure that any effects are due to
the test chemical and not the carrier. The suggested use of
carrier controls to distinguish between test chemical and
carrier effects is considered a standard laboratory
practice.
Water-soluble chemicals are easiest to apply since roots
may absorb chemicals directly from hydroponic or nutrient
solutions and since many air-borne chemicals arrive in
precipitation. Water-insoluble chemicals require other
procedures to insure contact between the test chemical and
the root and shoot systems. Appropriate controls are
necessary and are designed to reduce as much as possible,
the number of variables in the system. One means of
incorporating a water-insoluble chemical into the hydroponic
solution or the nutrient medium is to solubilize it in the
solution with a surfactant or carrier compound. When a
carrier is used, a carrier control is included in the
procedure to check the carrier's effects on plant growth. A
minimal quantity of carrier should be used to minimize
carrier effects.
Foliar applications of powders or sprays should be done
under a hood to ensure worker safety and to minimize growth
chamber contamination. Control plants should also be moved
from chambers to the hood area and sprayed with distilled,
deionized water whenever the chemical is applied to other
plants by spraying because plants are highly sensitive to
mechanical stresses such as handling (Wheeler and Salisbury
1979) .
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Attention to maintenance of environmental conditions
during applications of gasses is essential to success of the
test. Brenann and Leone (1968) observed that plants exposed
to identical concentrations of pollutants at different gas
flow rates exhibited different toxicity symptoms. These
observed variations are understandable when considered in
terms of the effect of environmental physics on mass
transport to the leaves and on the physiological condition
of the plants. In order that each plant in the gas exposure
chamber receive the same dosage, chamber design should
provide a spatially uniform test chemical distribution.
Movement of gaseous substances to the leaf surfaces is
dependent upon air turbulance around the leaves reducing the
boundary layer thickness (Fowler and Unsworth 1974, O'Dell
et al. 1977). Some substances may impact or condense on the
surface of the plants (Fowler and Unsworth 1974) while
others may diffuse through stomatal pores or across the
cuticle to the leaf's interior (Black and Unsworth 1979,
Rich et al. 1970). The cuticle provides an effective
barrier for gaseous substances and several investigators
have shown chemical absorption to be regulated by the same
factors that control stomatal opening including light,
temperature, humidity, carbon dioxide concentration, and
plant water status (Bennett et al. 1973, Black and Unsworth
1979, Rich et al. 1970, Winner and Mooney 1980). Specified
environmental conditions allow for normal stomatal response
and plant growth. These conditions should be maintained
during plant exposure to chemicals. The exposure chamber
designed by Rogers et al. (1977) meets the environmental
criteria and the criteria of uniform gas distribution within
the chamber.
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The growth medium control provides a baseline check of
the effects of hydroponic solution, potting material,
nutrient solution, and the testing environment on plant
growth during each test. This control is simply the plant
growing in the hydroponic solution or root support material
and maintained by the nutrient solution without test
chemical, solvent, or carrier. This control is required in
each run regardless of whether or not the chemical is
soluble in water. It is the only control used when the
chemical is water-soluble and is applied directly to the
support media or is applied as foliar dust or spray.
A second control is also recommended whenever a carrier
(solubilizer, or non-aqueous solvent) is used to maximize
contact between the chemical and the plant. The solvent or
carrier control is used to check carrier effects on plant
physiology and growth and on processes affecting chemical
uptake and translocation.
If chemical volatilization is anticipated, a
modification of the double pot method is used in which
germinated seedlings of uniform size are transplanted into
the potting containers; an impervious barrier of
polyethylene film is placed over the root support material
and seedlings are positioned such that their roots are
exposed to the support material and are sealed in place with
a ring of non-toxic, inert, pliable putty.
2., Range-Finding Test
It is recommended that a range-finding test be conducted
prior to the uptake and translocation test to reduce the
risk of using an inappropriate concentration series in the
definitive test. In order to minimize the costs and time
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required to obtain the requisite data, nominal
concentrations are permitted, replicates are not required,
and other test procedures and conditions are relaxed.
It may be possible to use results of previously
conducted plant effects tests rather than conducting a
range-finding test. For instance, results of the seed
germination/root elongation test, the early seedling growth
test, or other appropriate phytotoxicity tests may be used
to establish the appropriate upper concentration for
t es t i ng .
3. Definitive Test
The specific requirements of the uptake and
translocation test are the analytical determinations of
chemical con-centrations, the selection of uniformly sized
seedlings, the unbiased selection of seedlings for each
treatment, the use of controls, and the recording, analysis,
and presentation of data. These requirements assure that
the propensity for a chemical to accumulate in plants or
plant parts is accurately elucidated, that uptake and
translocation phenomena are not confounded by differential
plant vigor, and that the results are clearly presented.
Reporting the occurrence of such abnormalities as stunting,
discoloration, chlorosis and/or necrosis of leaves, etc.
provides qualitative data that may help explain deviations
from expected uptake trends.
The purpose of the uptake and translocation test is to
determine the propensity of a chemical to accumulate in
plants or plant parts for each species tested with a minimum
of testing beyond the range-finding test. The concentration
range for the uptake and translocation test is based upon
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the results of the range-finding test for that species. It
is probable that each species tested will require a
different exposure concentration series and will exhibit
different uptake and translocation rates.
Plants from 2 of the 6 pots per treatments are pooled,
giving 3 replicate sample pools per treatment. The purpose
of pooling the plants is to insure that there is sufficient
sample biomass for all the specified analyses. Since the
reported results should include a statistical summary of the
data, plants from al1 6 pots per treatment cannot be pooled
into one sample pool. However, if chemical recovery is
insufficient to permit measurement (not due to sensitivity
of the analytical method), a single analysis of pooled
material from all six pots may be required to confirm lack
of uptake.
The randomized complete block design is the recommended
experimental design for the uptake and translocation test.
If growth chambers are used, it is essential that the
investigator randomly assign plant containers to treatments
to assure that each plant has the same chance of receiving
any of the treatments (exposure level of test chemical)
(Hammer and Urquhart 1979). To account for variation within
the chamber and to increase the sensitivity for detecting
treatment differences, small square blocks should be
delineated with randomization of treatments within blocks.
Replication should occur over chambers (of the same type)
as, in many cases, a with in-chamber estimate of residual
variance badly underestimates the between chamber estimate
(Hammer and Urquhart 1979). This means that differences
between chcimbers are often greater than differences in
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growth and environmental conditions within chambers. In the
event that blocking within chambers is impossible, total
randomization is acceptable.
In order to substantiate that abiotic test conditions
were maintained within specified limits, it will be
necessary to measure and record irradiation, photoperiod,
temperature, and humidity throughout the test. Requisite
instruments are readily available, easily maintained, and
should not increase complexity or costs of the test. In
fact, these measurements should be made in order to maintain
the specified growing conditions.
To standardize procedures and provide acceptable data,
McFarlane (1979) suggested that radiation measurements be
taken at the top of the plant canopy and that a mean, plus a
maximum and a minimum value, be obtained over the plant-
growing area. These measurements should be taken daily and
should be taken and recorded at least at the start and
finish of the test. If greenhouse facilities are used,
hourly measurements of irradiation should be recorded and
presented as daily total irradiance. Representative hourly
curves of irradiation for clear sky conditions and cloudy
days should also be presented to more fully describe the
light climate.
The photoperiod to which each test organism is exposed
should be reported. If deviations from the recommended
photoperiod occur, they should be recorded so that the
potential impact on plant growth can be assessed.
Day/night temperature measurements should be taken at
least daily at the top of the plant canopy. Data reported
should include the average values for each light and dark
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period in the study and the range of variation over the
growing area.
Humidity should be measured at the same time and
location as temperature, and correspondingly reported.
If plants are grown in chambers, it will be necessary to
monitor carbon dioxide concentration in the chamber air to
ensure that excessive depletions do not occur as a result of
rapid photosynthesis rates of the seedlings. These data
should also be reported.
4. Analytical Measurements
It is imperative that the actual concentration of
chemical to which plants are exposed be determined with the
best available analytical precision. Analysis of stock
solutions and test solutions just prior to use will minimize
problems with storage such as formation of degradation
products, adsorption to container walls, or chemical
transformations. Nominal concentrations are not adequate
for the purposes of the definitive tests.
The pH of the test solution should be measured prior to
testing to determine if it lies outside of the species'
optimal range. While it is recognized that plants grow over
a broad range of hydrogen-ion concentration and typically
exhibit a pH optima for growth, this test guideline does not
include pH adjustment for the following reasons: the use of
acid or base may chemically alter the test chemical making
it more or less toxic, the amount of acid or base needed to
adjust the pH may vary from one test solution concentration
to the text, and the effect the test substance has on pH may
indirectly affect the growth, development, and uptake
processes of the test plant. Therefore, the pH of each test
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solution should be determined and compared with the
acceptable range for growth and development of the test
species.
Harvested plants are rinsed with water to remove any
surface depositions of the test chemical. This rinse water
is retained for purposes of the mass balance calculations.
The plants are separated into component parts representative
of use. Plant component parts are pooled to provide
sufficient material for chemical analysis. Subsamples of
the pooled plants are oven dried and weighed; and dry weight
equivalents are calculated for each component. This
procedure avoids the potential volatilization of organic
chemicals from the plant tissues during the drying process.
Because the chemical may be differentially soluble in
organic and aqueous solvents, both are included in the
extraction procedure. The nature of the organic solvent
used will depend upon the particular chemical properties.
Hexane or acetonitrite is suggested in the homogenization
procedure. They are commonly used organic solvents and most
organic compounds will dissolve in them. Anhydrous sodium
sulfate is used to absorb water from the mixture and
minimize moisture complications in the procedure. The
organic solution is separated from the homogenate and may be
analyzed by liquid scintillation spectrometry in order to
determine the amount of radioactive compound in the
extractable organic portion.
If the chemical is present in the organic extract, it
can be identified and quantified by thin-layer
chromatography followed by autoradiography of the thin-layer
plates and/or counting with a radiochromatogram scanner.
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A water soluble chemical would be expected to appear
primarily in the aqueous extract of the plant sample
homogenate. Therefore, chemical analysis of the aqueous
extract is also conducted as above.
Metabolites and soluble residues of the chemical may
also be present in the organic and aqueous extracts of plant
homogenates. Liquid scintillation counting of the extracts
followed by evaporation and thin-layer chromatography
provides a measurement of metabolites and soluble residues
of appropriately labeled chemicals but does not allow for
differentiation between chemical forms.
Bound residues are those forms of the chemical in the
plants that are not extractable in either organic or aqueous
solvents. Plant tissues should be digested in some manner
before bound residues can be measured. Three methods are
available: (1) digestion with a tissue solubilizer, (2)
complete combustion oxidation and collection of carbon
dioxide and (3) wet digestion. When the test chemical is
labeled with 3H or 14C, either digestion with a tissue
solubilizer or complete combustion oxidation is
applicable., Of the two, combustion oxidation is preferred,
because tissue solubilization does not dissolve all material
at the same rate and may result in quenching and
interference in the scintillation solution. This problem
does not appear in the carbon dioxide solution resulting
from combustion oxidation. When the test chemical is
labeled with 32P or 35S, either digestion with a tissue
solubilizer or wet digestion is appropriate. Wet digestion
is preferred for 32P or 35S determination because it results
in a more complete digestion of plant tissue than occurs
with a tissue solubilizer.
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Although it may be possible to determine plant uptake of
gases by analytical chemistry, a direct mass balance
calculation during plant exposure may be preferred. This
would completely eliminate the need to do chemical analyses
of plant tissues and be very time and cost effective.
Measured rates of photosynthesis and transpiration prior to
fumigation are essential in order to establish that
environmental conditions in the exposure chamber are
conducive to normal physiological processes and that the
stomates are open. In addition, this establishes a base
level rate (zero chemical concentration) of important
physiological processes. Rates of photosynthesis during and
after fumigation provide indications of chronic
phytotoxicity and, with data on stomatal conductivity, also
allow considerations of mechanism of chemical action and of
plant resistance to the chemical (Winner and Mooney 1980).
Details of equipment requirements and calculations for gas
exchange/fumigation studies can be found in Bennett et al.
(1973), Black and Unsworth (1979), Keller (1980), O'Dell et
al. (1977), Rogers et al. (1979), Thompson et al. (1979),
and Winner and Mooney (1980).
The mass of pooled plant organs and pooled whole plants,
the mass of chemical in each plant pool, rates of trans-
piration and photosyn thetis, and values of stomatal
conductivity (when determined) are analyzed. Means and
standard deviations are calculated and plotted for each
treatment and control and the data are subjected to analysis
of variance.
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C. Test Conditions
1. Test Species
a. Selection
The ten terrestrial plant species recommended for the
seed germination/root elongation, early seedling growth, and
plant uptake test guidelines are as follows:
Lycopersicon esculentum (tomato)
Cucumis sativus (cucumber)
Lactuca sativa (lettuce)
Glycine max (soybean)
Brassica oleracea (cabbage)
Avena sativa (oat)
Lolium perenne (perennial ryegrass)
Allium cepa (common onion)
Daucus carota (carrot)
Zea mays (corn)
In addition, other species of economic or ecological
importance to the region of impact may also be tested in
lieu of these species. Two species are to be used in the
plant uptake test.
These species have been selected for the following
reasons:
o As food, forage, or ornamentals they are
economically important and constitute major cash
crops .
o Their distribution, abundance and taxonomic
representation suggest broad coverage of the plant
kingdom.
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o They are also specified for phytotoxicity testing
of pesticides (Subpart J, Pesticide Registration
Guidelines). Additional justification for
selection of these test species is provided in
these guidelines (see FR 45(214): 72948-72978).
o They are sensitive to many toxic compounds and have
been used to some degree in previous bioassays.
Their use in herbicide bioassays, heavy metal
screening, salinity and mineral stress tests and
allelopathic studies indicates a sensitivity to a
wide variety of stressors (Guenzi and McCalla 1966,
Geronimo et al. 1973, Puerner and Seigel 1972,
Wiedman and Appleby 1972, Reynolds 1978, Chang and
Foy 1971) .
o They are compatible with the environmental growth
conditions and time constraints of the test
method. Seed from the selected species germinate
quickly and easily. Root growth is rapid and
uniform. The seed contain no natural inhibitors and
require no special pretreatment to germinate (such
as soaking, chilling, prewashing, light, or
scarification).
Other species may be substituted for any or all of these
ten species when appropriate. For example, forest or desert
species may constitute the population at greatest risk. If
so, those of most value to man or of ecological dominance to
the affected ecosystem should be selected. The rationale
for selection of alternative species should be discussed
with the Agency and/or supported in the report of findings.
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2. Facilities
a. Apparatus
Greenhouse space, growth rooms, or environmental
chambers are equally acceptable as long as the specified
environ-mental conditions are maintained. The environment
affects growth, metabolism, evapotranspiration, and
photosynthesis and thereby directly and indirectly affects
the plants' abilities to take up, metabolize, or accumulate
chemicals. Laboratory facilities typically needed include
standard laboratory glassware, a source of distilled water,
work areas to clean and prepare equipment and to measure
chemical concentrations and plant responses, drying ovens,
appropriate equipment for chemical analysis, and proper
disposal facilities. Without these facilities, the testing
cannot be adequately conducted.
b. Containers and Support Media
Containers for hydroponic solution or root support media
should be composed of inert materials, such as polyethylene,
in order to prevent reaction of test chemicals and/or
carriers with the container. The root support material
should be composed of inert materials for these same
reasons. Quartz sand and glass beads are suitably inert
materials and do not readily adsorb substances. This lack
of adsorption is important because the chemical should be
available for root uptake; it would not be as readily
available if it were adsorbed to the root support material
or container wall.
Sand or glass beads are used, rather than soil, to avoid
complications associated with variable physical and chemical
properties and microbial populations indigenous to native
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soils. Native soils are undesirable because of the varying
clay, sand, and humus components, the types and proportions
of which vary within the same soil type. Microbial
populations also vary between soil types. These variables
alter moisture holding capacity, chemical binding capacity,
aeration, and nutrient and trace element content (Audus
1964, Beetsman et al. 1969, Beall and Nash 1969). In
addition to the variations in these physical factors, there
will also be variation in such chemical properties as pH and
redox potential. Because of the impossibility of
controlling physical and chemical properties of native
soils, inert material is required to support the plants with
the only variables being the presence and concentration of
test chemical. The purpose of using glass beads or sand
instead of native soils is not to make test results more
directly applicable to natural systems, rather it is to
eliminate sources of variation in the test.
c. Cleaning and Sterilization
Standard good laboratory practices are recommended to
remove dust, dirt, other debris, and organic and inorganic
residues from the test containers and support media which
might confound test results. Residues could enter the
hydroponic or nutrient solution and be taken up by plants,
affecting their growth and/or other metabolic activity,
resulting in misleading data. A dichromate solution should
not be used for washing containers or beads because
dichromate may enter the nutrient solution, be taken up by
plants, and affect their growth and metabolic activity.
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d. Nutrient Media
The nutrient medium specified in the protocol is
modified half-strength Hoagland solution (Downs and Helmers
1975). This nutrient solution is recommended because all
the constituent compounds are relatively easily obtained and
because it works well for the culture of terrestrial plants.
Hydroponic or nutrient culture techniques eliminate
spatial gradients within the growth medium and maintain the
root system at uniform levels of aeration, nutrients, and
water status (Rawlins 1979). The simplest and most
practical method for routine plant growth in controlled
environments is a sand or gravel culture in which the plants
are grown in containers of sand which are periodically
filled to provide water and nutrients, and drained to
provide aeration. The frequency of irrigation depends on
the storage capacity of the sand and the rate of water use
by the plant. Use of aerated hydroponic solution without a
root support media may be preferred by some laboratories.
Reliability is the primary consideration in the design
of the system which automatically fills and drains the
containers. One of the simplest systems includes a
standpipe connected to an outlet at the bottom of each
container through a manifold. The tops of the containers
are at a uniform elevation. Periodically, the solution
level in the standpipe is raised to the corresponding level
and then lowered below the bottom of the containers. This
system requires only one conduit to each container (for both
filling and draining) and permits variation in the number of
containers. The standpipe is filled by a pump from a lower
storage reservoir. Electrical power to start the pump and
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close the drain valve is controlled by a clock timer and,
when the nutrient solution eaches the desired level in the
standpipe, a float switch turns off the power to the pump
and opens the drain allowing the solution to drain back to
the reservoir.
If the rate of uptake for test chemicals applied in the
nutrient solution differs from the rate of nutrient and
water uptake, the exposure concentration will change with
time and differ from that originally specified. In
addition, chemicals applied to foliage as sprays or dust
will enter the rooting zone and accumulate in the nutrient
solution. While foliar application is designed to allow for
chemical movement through, and uptake from, the rooting
zone, excessive concentrations of test chemical should not
be allowed to accumulate in the nutrient solution. The test
solution should be replaced when or if the test chemical
concentration differs from that specified by ± 20 percent.
When the chemical is applied to the foliage, the nutrient
solution should be replaced weekly if detectable quantities
move through and accumulate in the solution following
spraying or dusting of foliage.
3. Environmental Conditions
Controlled environmental conditions are necessary to
maintain uniform growth and ensure reliable data.
Maintenance of specified environmental conditions before,
during, and after plant exposure to the chemical is
essential to successful execution of this test. Variability
of plant response to chemical exposure as a result of
fluctuations in environmental conditions, has been noted
(Darwent and Behrens 1972, Dunning and Heck 1973, Juhren et
al. 1957, Leone and Brennan 1970).
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Environmental conditions affect growth, metabolism,
evapotranspiration, and photosynthesis of plants. In
addition to mineral nutrition the conditions that should be
standardized and maintained include: (a) carbon dioxide
concentration; (b) relative humidity; (c) irradiation; (d)
photoperiod; (e) day and night temperatures.
Standardization of environmental conditions is
essential. Several investigators have demonstrated that
differences in environmental conditions influence the
response of plants exposed to chemicals. These include pre-
conditioning by light and humidity (Dunning and Heck 1973),
effects of temperature, photoperiod, and light intensity
during the growth (Juhren et al. 1957), air movement during
exposure (Brenann and Leone 1968), and mineral nutrition
(Leone and Brenann 1970). Environmental conditions between
growth chambers should be closely maintained, as specified
in this protocol, to ensure common test conditions. If
large growth rooms or greenhouse facilities are used,
comparability of the environment between small groups of
plants is not as critical, and environmental specifications
may be relaxed.
a. Carbon Dioxide Concentration
The carbon dioxide concentration should be high enough
for photosynthesis to occur at a level which will allow
normal plant growth and biomass accumulation.
Photosynthetic rates vary directly with carbon dioxide
partial pressures. In addition, abnormally high levels of
carbon dioxide can affect carbohydrate translocation from
the leaves. Therefore, the concentration of carbon dioxide
in the air surrounding test plants should be kept within
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limits conducive to normal plant growth (350 ± 50 ppm). The
carbon dioxide concentration is not expected to be of
concern in greenhouse facilities.
In growth chambers, however, carbon dioxide depletion by
rapidly photosynthes izing plants may be a real concern. The
extent of depletion is a function of chamber volume, rate of
air exchange between the chambers and the outside, the
number, size, and type of plants, and the growth conditions
maintained in the chambers (Hellmers and Giles 1979, Pallas
1979). Attempts to reduce carbon dioxide depletion by
increasing the air exchange rate between the outside air and
the chamber would probably eliminate temperature and
humidity control within the chambers (Downs and Hellmers
1975). An appropriate system for carbon dioxide control in
up to 5 chambers is described by Hellmers and Giles
(1979). Alternatively, the use of fewer plants per chamber
may be an appropriate means of avoiding carbon dioxide
depletion problems.
b. Relative Humidity
A literature review by Hoffman (1979) indicates that
most plants grow well when the atmospheric saturation
deficit is maintained between 5 and 10 mb. At 25°C, the
optimum relative humidity range is 65 to 85 percent. Most
environmental control facilities with provision for humidity
control can maintain humidity in this range with an accuracy
at 25°C of ± i.o to 1.6 mb of vapor pressure (± 3 to 5
percent relative humidity). Humidity levels have been
observed to affect plant growth and development, rates of
carbon dioxide exchange, flowering, nutrient transport, and
susceptibility to air pollution (Thurtell 1979). However,
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these are generally the responses of plants to their total
environments and are not unique to the humidity of the
air. During the daylight period the relative humidity in
the growth chamber should be maintained within the optimal
range of 65 to 85 percent. When several chambers should be
maintained as comparable environments, a relative humidity
of 70 ± 5 percent is recommended. During the dark period,
relative humidity should approach saturation, as it normally
does in natural environments.
c. Irradiation
The growth chamber irradiation spectrum should be as
similar to natural sunlight as possible in order for test
plants to respond normally. Light drives the photo-
synthetic process and specific wavelengths cause particular
responses (e.g., the phytochrome reactions) in plants.
Therefore, it is important that light quality and intensity
remain constant throughout the test. Irradiation in
chambers should be 350 ± 50 uE/m2 sec at 400-700 nm measured
at the top of the plant canopy. This corresponds roughly to
full, direct sun plus diffuse radiation. Artificial
lighting is used in growth chambers and as supplemental
lighting in greenhouses. However, lamps may weaken, cease
to function, or function abnormally and fail in the course
of long-term tests. Lamp failures should be corrected as
soon as observed and the changes in light quality and
intensity resulting from the lamp failures should be
recorded.
For light quality and intensity to approach that of
natural sunlight, the light source commonly consists of
either a fluorescent-incandescent system, a high intensity
21
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discharge system (HID) composed of metal halide lamps, or a
combination of metal halide and high-pressure sodium
lamps. Combinations of lamps are necessary for artificial
lighting because there is no single lamp which is capable of
emanating light of the same quality and intensity as natural
sunlight, although the metal halide lamp closely approaches
ideal lighting conditions.
HID systems produce intense light that may not be
obtainable in other ways (McFarlane 1978). Because these
lamps emit radiation in discrete line spectra, researchers
frequently combine two different types of HID lamps in
growth chambers to obtain a more balance spectrum. Metal
halide lamps appear to be the most useful type of HID lamp
because their emission spectra are almost continuous over
the 400 to 700nm wavebands. However, a combination of metal
halide and sodium HID lamps will provide superior plant
growth to either one alone.
A fluorescent-incandescent system should be composed of
70-80 percent input wattage of cool-white fluorescent lamps
and 20-30 percent wattage of incandescent lamps (Downs
1975). Although incandescent lamps are capable of emanating
light which qualitatively approaches natural sunlight, the
intensity will not approach the 350_+50 uE/m2 sec specified
for plant growth. Increasing the number of incandescent
lamps would result in overheating the growth chamber.
Fluorescent lighting alone can approach the intensity of
natural sunlight without overheating the growth chamber,
since very little of the output is in the infrared range
(McFarlane 1978).
22
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August, 1982
Incandescent lights should be included to provide
radiation in the red and infrared regions. Uniformity and
intensity of lighting within chambers is routinely improved
by covering the walls with highly reflective materials.
d. Photoperiod
Plants exhibit three basic photoperiodic responses.
Long-day plants will flower only when the light period is
longer than a certain minimum number of hours in a 24 hour
day/night cycle. Short-day plants will flower only when the
light period is shorter than a certain maximum number of
hours in a 24 hour day/night cycle. Day-neutral plants will
flower regardless of the length of the light period. In this
test, long day conditions (16 hours light/8 hours darkness)
are recommended for all long-day and day-neutral plants in
order to maximize biomass. Continuous light, however, is
not recommended as a dark period is required for the
phytochrome interconvers ions necessary in photo-synthetic
vascular plants (Salisbury and Ross 1969).
e. Day/Night Temperature
For any particular plant species, there is an optimal
temperature regime for maximum growth and development. This
regime may differ between the phenophases of germination,
vegetative growth, flowering, and fruit development. The
concept of thermoperiodicity specifies that the temperature
during the light period should be different from that of the
dark period for optimal plant growth and development (Downs
1975). It has been demon-strated that plants grow and
develop better with a day/night temperature differential
(Kramer 1957, Went 1957). While specific temperature optima
can be identified for each species, a regime of 25°C/20°C
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August, 1982
± 3°C is recommended as it will promote suitable growth and
be cost effective. If growth chambers or rooms are used,
tolerances of ± 1°C ace recommended to ensure environmental
comparibility between growth facilities.
D. Reporting
The sponsor should submit to the Agency all data
developed during the test. If testing specifications are
followed, the sponsor should report that specified
procedures were followed and present the results. If
alternative procedures were used instead of those
recommended in the test guideline, then the protocol used
should be fully described and justified.
Environmental test (growth) conditions, chemical
concentrations, quantity of chemical applied, number of
applications, test data, and statistical analyses should all
be reported. The justification for this body of information
is contained in this support document. If plant species
other than those recommended were used, the rationale for
the selection of the other species should also be provided.
The data obtained from the plant uptake and
translocation test should demonstrate whether or not there
is uptake and translocation of the test chemical.
Concentrations of substances in plants are commonly
expressed in terms of weight per unit plant weight (e.g.,
ug/g dry plant). A subsample of plant material is selected
after harvest and dried at 70°C for moisture content
determination and expression of plant material on a dry
weight basis. Concerns for chemical loss by volatilization
in the drying ovens mandate that the entire mass of plant
material not be dried. Concentrations of the test chemical
24
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August, 1982
may be identical in two plants of different mass with the
smaller plant containing less total chemical than the larger
one. Test chemical concentration should be normalized with
respect to mass in order to prevent the misleading reporting
of uptake data. Both total quantity and concentration of
chemical are required data.
If the chemical enters the root in bulk flow,
transpiration and evaporation from the leaves will
proportionately increase the uptake of the chemical. Plants
that transpire more rapidly may have greater uptake rates.
Consequently, a record of evapotranspiration rates is
required. Chemical concentrations should be normalized with
respect to evapotranspiration, as well as with biomass, in
order to prevent misleading reporting of uptake data.
Furthermore, any effects of the chemical on transpiration
may also be identified.
The free parent chemical is that which is applied to the
plant either in the nutrient media or as spray or dust and
is extractable from the plant tissue in organic or aqueous
solution. It is very important to quantify the free parent
chemical as it is potentially toxic to living systems and
appears in portions of plants consumed by man and domestic
animals. Because of its potential toxicity, it is
undesirable that such chemicals should be passed from one
food web component to another. Data on the concentration of
free parent chemical should be included in the report of
testing because it probably is a measure of the maximum
amount of chemical that can be expected to be found in
plants exposed to the chemical in the natural environment.
The quantity of chemical taken up by the plant is probably
25
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August, 1982
more than would occur in the natural environment because the
test is designed to provide data on "worst case" conditions.
The chemical may not remain intact in its original form
in plants. Instead may be metabolized, combined with
smaller or larger molecules, or otherwise modified. If the
chemical or its degradation products enter normal plant
metabolic pathways, metabolites may be formed. Since the
chemical is not a normal substrate for metabolism, the
metabolites are not likely to be those normally expected
from plant metabolic activities. These abnormal metabolites
may be chemicals that are as toxic or more toxic to living
systems than the parent chemical. Although metabolite
identification, because of analytical problems, is not the
primary objective of the test, it may be possible to
determine their presence in order to give some indication of
chemical fate in plants and the mode of dissemination in
food chains. This is especially important when the
metabolites are potentially toxic. Bound residues, those
forms of the test chemical which are not extractable in any
of the solvents used, are portions of the chemical, or the
intact molecule itself, which combine with structural
components of plant cells. Bound residues may also be toxic
to living systems because they may not remain bound when
they are consumed and disseminated in food chains. Because
of analytical problems, identification of bound residues is
not an objective of the test; however, they should be
quantified in order to give some indication of the fate of
the chemical in plants and is mode of dissemination in food
webs .
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August, 1982
The total amount of chemical recovered in plants and
nutrient media, after known addition of the chemical to the
system, would indicate, by difference, the quantity lost by
volatilization and/or degradation. Since the test from
which these data are obtained is primarily an uptake test,
mass balance data should be included to account for all the
chemical added to the system as opposed to that which was
actually taken up.
III. Economic Aspects
The Agency awarded a contract to Enviro Control, Inc. to
provide us with an estimate of the cost for performing a
plant uptake and translocation test according to this
Guideline. Enviro Control supplied us with two estimates; a
protocol estimate and a laboratory survey estimate.
The protocol estimate was $20,814. This estimate was
prepared by identifying the major tasks needed to do a test
and estimating the hours to accomplish each task.
Appropriate hourly rates were then applied to yield a total
direct labor charge. An estimated average overhead rate of
115%, other direct costs of $4,463, a general and
administrative rate of 10%, and a fee of 20% were then added
to the direct labor charge to yield the final estimate.
Enviro Control estimated that differences in salaries,
equipment, overhead costs and other factors between
laboratories could result in as much as 50% variation from
this estimate. Consequently, they estimated that test costs
could range from $10,407 to $31,221.
The laboratory survey estimate was $22,100, the mean of
the estimates received from two laboratories. The estimates
ranged from $19,200 to $25,000 and were based on the costs
to perform the test according to this Guideline.
27
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IV. References
Audus L J. 1964. Herbicide behavior in the soil. II.
Interactions with soil microorganisms. In: The
physiology and biochemistry of herbicides. New York:
Academic Press, pp. 168-206.
Beall ML Jr. and Nash RG. 1969. Crop seedling uptake of
DDT, dieldrin, endrin, and heptachlor from soil. Agron.
J. 61:571-575.
Beetsman GD, Kenney DR, Chesters G. 1969. Dieldrin
uptake by corn as affected by soil properties. Agron.
J. 61:247-250.
Bennett JH, Hill C, Gates DM. 1973. A model for
gaseous pollutant sorption by leaves. J. Air. Poll.
Control Assoc. 23:957-962.
Black VJ and Unsworth MH. 1979. Resistance analysis of
sulphur dioxide fluxes to Vicia faba. Nature 282:68-69.
Brenann E and Leone IA. 1968. The response of plants
to sulfur dioxide or ozone-polluted air supplied at
varying flow rates. Phytopathology 58:1661-1669.
Chang I and Foy CL. 1971. Effect of picloram on
germination and seedling developmnt of four species.
Weed Sci. 19:58-64.
Chou SF, Jacobs LW, Penner D, Tiedje JM. 1978. Absence
of plant uptake and translocation of polybrominated
biphenyls (PBBs). Environ. Health Perspective 23:9-12.
Cole LK, Sanborn JR, Metcalf RL. 1976. Inhibition of
corn growth by aldrin and the insecticide's fate in the
soil, air, crop, and wildlife of a terrestrial model
ecosystem. Environ. Entomology 5:583-589.
Darwent AL and Behrens R. 1972. Effect of pretreatment
environment on 2,4-D phytotoxicity. Weed Sci. 20:540-
544.
Downs RJ. 1975. Controlled Environments for Plant
Research. New York: Columbia University Press.
28
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August, 1982
Downs RJ and Helmers H. 1975. Environment and the
environmental control of plant growth. New York:
Academic Press.
Dunning JA and Heck WW. 1973. Reponse of pinto bean
and tobacco to ozone as conditioned by light intensity
and/or humidity. Environ. Sci. and Tech. 7:824-826.
Fowler D and Unsworth MH. 1974.
sulphur dioxide on wheat. Nature
Dry deposition
249:389-390.
of
on
Fuhr F and Mittelstaedt W. 1980. Plant experiments
the bioavailability of unextracted [carbonyl-14C]
methabenzthiazuron residue from soil. J. Agric. Food
Chem. 28:122-125.
Geronimo J, Smith LL, Jr., Stockdale GD, Goring CAI.
1973. Comparative toxicity of nitrapyrin and its
principal mtabolite, 6-chloropicolinic acid. Agron. J.
65:689-692.
Guenzi WD and McCalla TM. 1966. Phenolic acids in
oats, wheat, sorghum and corn residues and their
phytotoxicity. Agron. J. 58:303-304.
Hammer PA and Uquhart NS. 1979. Precision and
replication: Critique II. In: Controlled Environment
guidelines for plant research [Tibbitts TW and Kozlowski
II, eds.] New York: Academic Press, pp. 364-368.
Hellmers H and Giles LJ. 1979. Carbon dioxide:
Critique I. In: Controlled environment guidelines for
plant research [Tibbitts TW and Kozlowski TT, eds.] New
York: Academic Press, pp. 229-234.
Hoffman GL. 1979. Humidity. In: Controlled environment
Guidelines for plant research [Tibbitts TW and Kozlowski
TT, eds.] New York: Academic Press, pp. 141-172.
Juhren M. Nobel W, Went FW. 1957. The standardization
of Poa annua as an indicator of smog concentrations. I.
Effects of temperature, photoperiod, and light intensity
during growth of the test plants. Plant Physiol.
32:576-586.
29
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August, 1982
Keller T. 1980. The simultaneous effect of soil-borne
NaF and air pollutant S02 On C02 - uptake and pollutant
accumulation. Oecologia (Berl.) 44:283-285.
Kelly JM, Parker GR, McPee WW. 1980. Heavy metal
accumulation and growth of seedlings of five forest
species as influenced by soil cadmium level. J. Environ.
Qual. 8:361-364.
Kramer PJ. 1957. Some effects of various combinations
of day and night temperatures and photoperiod on height
growth of lobololly pine seedlings. Forest Sci. 3:45-
55.
Leone IA and Brenann. 1970. Ozone toxicity in tomatoes
as modified by phosphorous nutrition. Phytopathology
60:1521-1524.
McFarlane JC. 1979. Radiation: Guidelines. In
Controlled environment guidelines for plant research
[Tibbits TW and Kozlowski TT, eds. ] Neww York: Academic
Press, pp. 55-74.
O'Dell RA, Takeri M. Kabel RL. 1977. A model for uptake
of pollutants by vegetation. J. Air. Poll. Control
Assoc. 27:1104-1109.
Pallas JE. 1979. Carbon dioxide. In: Controlled
environment guidelines for plant research [Tibbits TW
and Koslowski TT, eds.] New York: Academic Press, pp.
207-228.
Puerner N J, Siegel SM. 1972. The effects of mercury
compounds on the growth and orientation of cucumber
seedlings. Physiol. Plant. 26:310-312.
Rawlins SL. 1979. Watering. In: Controlled environment
guidelines for plant research [Tibbitts TW and Kozlowski
TT, eds.] New York: Academic Press, pp. 271-289.
Reynolds T. 1978. Comparative effects of aromatic
compound on inhibition of lettuce fruit germination.
Ann. Bot. 42:419-427.
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Rogers HH, Jeffries HE, Stahel EP, Heck WW, Ripperton
LA, Witherspoon AM. 1977. Measuring air pollutant
uptake; by plants: A direct kinetic technique. J. Air
Pollut. Control Assoc. 27:1192-1197.
Rogers HH, Campbell JC, Volk RJ.
dioxide uptake and incorporation
(L.) Science 206:333-335.
1979. Nitrogen-15
by Phaseelus vulgaris
Salisbury FB and Ross C,
California: Wadsworth.
1969. Plant Physiology.
Thompson CR, Kats G, Lennox RW. 1979.
air pollutants formed by high explosive
Environ. Sci. Technol. 13:1263-1268.
Phytotoxici ty
production.
of
Thurtell GW. 1979. Humidity: Critique I. In:
Controlled environment guidelines plant research
[Tibbits TW and Kozlowski TT, eds.] New York: Academic
press, pp. 173-175.
USEPA. 1979. U.S. Environmental Protection Agency.
Toxic substances control. Discussion of premanufacture
testing policy and technical issues; request for
comment. Fed. Regist. March 16, 1979. 44:16240-16292.
Went FW. 1957. Environmental control of plant growth.
Chronica. Botanica, Vol. 17. New York: Ronald Press.
Wheeler RM and Salibury FB. 1979. Water spray as a
convenient means of imparting mechanical stimulation to
plants. Hortscience 14:270-271.
Wickliff C, Evans HJ,
Cadmium effects on
Carter KR, Russell SA. 1980.
the nitrogen fixation system of
alder. J. Environ. Qual. 9:180-184.
red
Wiedman SJ and Appleby
simulation by sublthal
Weed Res. 12:65-74.
AP. 1972. Plant
concentrations of
growth
herbic ides
Winner WE and
resistance: I
deciduous and
Mooney HA. 1980. Ecology of SG>2
, Effects of fumigations on gas exchange
evergreen shrubs. Oecologia 44:290-295.
ot
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AVIAN DIETARY TEST
OFFICE OF TOXIC SUBSTANCES
OFFICE OF PESTICIDES AND TOXIC SUBSTANCES
U.S. ENVIRONMENTAL PROTECTION AGENCY
WASHINGTON, D.C. 20460
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Office of Toxic Substances EE-15
Guideline for Testing Chemicals August, 1982
AVIAN DIETARY TOXICITY TEST
(a) Purpose. The guideline in this subpart is designed to
develop data on the dietary toxicity to bobwhite and mallard of
chemical substances and mixtures subject to acute environmental
effects test regulations under the Toxic Substances Control Act
(TSCA) (Pub.L. 94-469, 90 Stat. 2003 15 U.S.C. 2601 et seq.) .
The Agency will use these and other data to assess the acute
hazard to birds and to provide an indication of potential chronic
hazard thac these chemicals may present to the environment.
(b) Def initions . The definitions in section 3 of tne Toxic
Substances Control Act (TSCA) and Part 792—Good Laboratory
Practice Standards apply here. In addition, the following
definitions apply to this guideline:
(1) "Acclimation" Physiological or behavioral adaptation of
test animals to environmental conditions and basal diet
associated with the test procedure.
(2) "LC50" The empirically derived concentration of the
test substance in the diet that is expected to result in
mortality of 50 percent of a population of birds which is exposed
exclusively to the treated diet under the conditions of the test.
(3) "Test substance" The specific form of a chemical or
mixture of chemicals that is used to develop the data.
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August, 1982
(4) "Exposure period" The five day period during which test
birds are offered a diet containing the test substance.
(5) "Post-exposure period" The portion of the test that
begins with the test birds being returned from a treated diet to
the basal diet. This period is typically three days in duration,
but may be extended if birds continue to die or demonstrate other
toxic effects.
(6) "Test period" The combination of the exposure period and
the post-exposure period; or, the entire duration of the test.
(7) "Hatch" Eggs or young birds that are the same age and
that are derived from the same adult breeding population, where
the adults are of the same strain and stock.
(8) "Basal diet" The food or diet as it is prepared or
received from the supplier, without the addition of any carrier,
diluent, or test substance.
(c) Test procedures — (1) Summary of test. (i) After birds
have been obtained, they should be acclimated for at least seven
days .
(ii) Test birds should be randomly assigned to the various
treatment levels and controls.
(iii) Definitive test concentrations should be established,
possibly requiring a range-finding test to be conducted first.
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August, 1982
(iv) The test substance should be mixed thoroughly and
evenly into the diet. Three treatment levels should be analyzed
for test substance concentrations.
(v) Birds should be weighed at the beginning of the exposure
period.
(vi) Birds should be observed regularly for mortality or
abnormal behavior; any findings should be reported.
(vii) Food treated with the test substance should be
replaced by untreated food (basal diet) after five days of
exposure. Food consumption during the exposure period should be
carefully estimated on a pen by pen basis.
(viii) Food consumption should be estimated for the post-
exposure period and birds should be weighed at the end of eight
days. Additional weights and food consumption estimates should
be determined if the test period is longer than the typical eight
days .
(ix) The mortality pattern should be examined, and a
statistical analysis should be conducted. The LC50 slope, and
confidence limits should be reported. A test for heterogeneity
of data should be conducted.
(x) Treated or positive control birds should be sacrificed
and disposed of properly. Negative control birds may be kept as
breeding stock, but should not be used in any other tests.
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August, 1982
(xi) The material to be tested should be analytically pure
and the degree of purity should be reported along with the
percentage of each impurity at levels specified in the test
rule. If specifically required by a test rule for a particular
substance or mixture, the technical grade should be tested. The
test rule will specify the degree of purity or a range of
compositions of the test substance.
(xii) A test is unacceptable if more than 10 percent of the
control birds die during the test.
(2) [Reserved]
(3) Range-finding test. Unless the approximate toxicity of
the test substance is k-nown already, a range-finding test should
be conducted to determine the test substance concentrations to be
used in the definitive test, under paragraph (d)(4)(iii) of this
section for details on concentrations for definitive tests.
Procedures for range-finding tests may vary, but generally,
groups of a few birds are fed three to five widely-spaced
concentrations for five days. A concentration series of 5, 50,
500, and 5,000 ppm is suggested. The results of the range-
finding test then may be used to establish the definitive test
concentrations .
(4) Definitive test—(i) Controls . (A) A concurrent
control is required during every test. The control birds should
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August, 1982
be from the same hatch as the test groups. Control and test
birds should be kept under the same experimental conditions. The
test procedures should be the same for control and treated birds,
except that no test substance should be added to the diets of
control birds. If a carrier is used in preparation of the test
diets, the same carrier should be added to the diets of control
birds in the highest concentration used for test diets. The use
of shared controls is acceptable for concurrent tests as long as
the same carrier is used for all the tests.
(B) A test is not acceptable if more than 10 percent of the
control birds die during the test period.
(C) A positive control (e.g., dieldrin standard) may be run,
but is not required for each test. However, a quarterly or semi-
annual laboratory standard (positive control) is recommended as a
means of detecting possible interlaboratory or temporal
variation. A laboratory standard is also recommended when there
is any significant change in food, housing, or source of birds.
(ii) Number of animals tested. In the definitive test, a
minimum of ten birds should be used for each dietary
concentration of the test substance. A minimum of twenty birds
should be used for the negative or carrier control. Thirty or
more control birds are preferable. If a positive control or
laboratory standard is used, ten or more birds should also be
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August, 1982
used for each concentration of the positive control. When a test
substance is known or expected to result in high experimental
variation, it may be appropriate or required by the test rule to
use additional birds.
(iii) Concentrations and dosage-mortality data. A minimum
of five concentrations of the test substance should be used in
the definitive test. These concentrations should be spaced
geometrically. The recommended spacing is for each concentration
to be at least 60 percent of the next higher dose (less than 1.67
times the next lower dose). If concentrations are spaced more
widely than is recommended, then at least three concentrations
should result in mortality between, but not including, 0 percent
and 100 percent. For any concentration spacing, at least one
concentration should kill more than 50 percent
(including 100 percent) and at least one concentration should
kill less than 50 percent (including 0 percent) of the birds in a
pen. For some test substances, it may be necessary to use more
than five concentrations to achieve these results.
(iv) Duration of test. The definitive test should include
five days of exposure to the test substance in the diet (exposure
period) followed by at least three days of additional observation
(post-exposure period) while the test birds are receiving an
untreated diet. If any test birds die during the second or third
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August, 1982
day of the post-exposure period or if toxic signs are evident on
the third day of the post-exposure period, the test period should
be extended until two successive mortality-free days and one day
free of toxic signs occur, or until 21 days after beginning the
test, whichever comes first.
(v) Observations of record. (A) Throughout the test
period, all signs of intoxication, other abnormal behavior, and
mortality s;hould be recorded and reported by dose level and by
day. Signs of intoxication are those behaviors apparently due to
the test chemical and may include a wide array of behaviors, such
as labored respiration, leg weakness, hemorrhage, convulsions,
ruffled feathers, etc. All signs of intoxication and any other
abnormal behavior, such as excessive aggression, toe-picking etc.
that may or may not be attributed to the test substance should be
reported. Among survivors, remission of signs of intoxication
and cessation of abnormal behavior should be recorded by dose
level and by day. When differential signs of intoxication are
observed within a dose level, an estimate of the number of birds
exhibiting such signs should be recorded. Observation of test
birds should be made, at a minimum, three times on the first day
of the exposure period. Observations also should be made at
least daily throughout the remainder of the test period; twice
daily observations are recommended, where feasible.
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August, 1982
(B) Average body weights of birds should be recorded and
reported for each pen within each treatment and control group at
the beginning of the exposure period and the end of the normal 3-
day post-exposure period of each test. Body weights 72 hours
before the exposure period are not required, but would provide
valuable base-line data. Average food consumption should be
measured in control pens and pens with the second lowest and
second highest concentration levels either daily or every other
day. Any significant amount of food spilled onto litter pans
should be estimated and reported. For all other pens, average
food consumption should be measured for both the exposure period
and the normal 3-day post-exposure period. If the study is
continued beyond eight days, body weight and food consumption
data should be recorded weekly.
(C) Gross pathology examinations are not required, but they
may provide valuable information on target site, mode of action,
etc .
(5) [Reserved]
( 6 ) Analytical measurements — ( i ) Statistical analysis.
(A) A statistical analysis should be conducted by transforming
the dietary concentrations to logarithmic values and the
mortality pattern to probits. Other acceptable methods that will
result in a theoretically straight line through _+_ 2 standard
8
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August, 1982
deviations from the LC50 valae mav be used' The LC5° V
slope of the transformed concentration-resonse curve should be
determined for mortality at the end of test period. Probit
analysis by calculation or graphical probit methods are
preferred. Any standard method that is used should provide the
slope of the transformed concentration-response curve as well as
the LC50 va-'-ae* A statistical test for goodness-of-f i t (e.g.,
chi-square test) also should be performed.
(B) All methods used for statistical analysis should be
described completely.
(ii) Analysis for test substance concentrations. (A)
Samples of treated diets should be analyzed to confirm proper
dietary concentration of the test substance. Analyses should be
conducted at the beginning of the exposure period with samples
from high, middle, and low concentrations. If not already
available, data should be generated to indicate whether or not
the test substance degrades or volatilizes. If the test
substance is known or found to be volatile or labile to the
extent that. 25 percent or more loss occurs over a five day
period, then a second series of analyses of the same
concentrations previously analyzed should be conducted at the end
of the exposure period.
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August, 1982
(B) The assay method used to determine actual concentrations
should be reported.
(iii) Analysis of basal diet. A nutrient analysis of the
basal diet should be included in the test report. For
commercially prepared basal diets, the list of ingredients
supplied by the company is normally sufficient if it is
detailed. The composition of any vitamin or other supplements
should also be reported.
(d) Test conditions — (1) Test species — (i) Selection.
(A) Bobwhite, Colinus virginianus (L.), and mallard, Anas
platyrhynchos L. , are the test species. Birds may be reared in
the laboratory or purchased from a breeder. If bobwhite are
purchased, it is preferable that they be obtained as eggs which
then are hatched and reared in the testing facility. During
incubation, a temperature of 39°C and relative humidity of 70
percent are recommended for bobwhite. It is feasible to purchase
live young bobwhite chicks if they can be obtained locally;
however, young bobwhite may suffer adverse efects if shipped by
air or other commercial means. Young mallard ducklings normally
can be shipped without undue adverse effects.
(B) All control and treatment birds used in a test should be
from the same source and hatch. Birds should be obtained only
from sources whose colonies have known breeding histories. Birds
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should be phenotypically indistinguishable (except for size) from
wild stock. It is recommended that birds be obtained from flocks
that have been outbred periodically with genetically wild stock
in order to maintain a genetic composition that approximates the
natural heterogeneity of the species.
(C) Birds used in the test should be in apparent good
health. Deformed, abnormal, sick, or injured birds should not be
used. Birds should not be used for a test if more than 5 percent
of the total test population die during the 72 hours immediately
preceding the exposure period. Purchased birds should be
certified as disease free or as bred from disease free stocks.
Birds should not have been selected in any way for genetic
resistance to toxic substances. Birds should not have been used
in a previous test, either in a treatment or control group.
(D) Test birds should be 10 to 17 days old at the beginning
of the exposure period. All treatment and control birds in a
test should be the same age _+ one day. The exact age should be
recorded and reported .
(E) Test birds should be acclimated to test facilities and
basal diet for a minimum of seven days. Acclimation to test pens
may be either in the actual pens used in the test or in identical
pens. Birds used in the test should be assigned randomly to
treatment and control pens without respect to sex. Randomization
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may be done either at the initiation of the acclimation period or
at the time when the birds are weighed at the beginning of the
exposure period.
(F) During holding, acclimation, and testing, birds should
be shielded from excessive noise, activity, or other
disturbance. Birds should be handled only as much as is
necessary to conform to test procedures.
(ii) Diet. (A) A standard commercial game bird (for
bobwhite) or duck (for mallard) starter mash, or the nutritional
equivalent, should be used for diet preparation. Antibiotics or
other medication should not be used in the diet before or during
the test. For bobwhite only, an antibiotic demonstrated to fully
depurate in 72 hours may be added to the drinking water, if
necessary, for birds up through 10 days of age; however, only
clean unmedicated water should be offered during the 96 hours
preceding the exposure period and-during the test period. It may
not be possible to obtain food that is completely free of
pesticides, heavy metals, and other contaminants; however, diets
should be analyzed periodically, and should be selected to be as
free from contaminants as possible. A nutrient analysis and list
of the ingredients in the diet should be included with the test
report.
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(B) The test substance should be mixed into the diet in a
manner that will result in even distribution of the test
substance throughout the diet. If possible, the test substance
should be added to the diet without the use of a diluent. If a
diluent is needed, the preferred diluent is distilled water; but
water should not be used as a diluent for test substances known
to hydrolyze readily. When a test substance is not water
soluble, it may be dissolved in a reagent grade evaporative
diluent (e.g., acetone, methylene chloride) and then mixed with
the test diet. The diluent should be completely evaporated prior
to feeding. Other acceptable diluents may be used, if necessary,
and include table grade corn oil, propylene glycol, and gum
arabic (acacia). If a diluent is used, it should not comprise
more than 2 percent by weight of the treated diet, and an
equivalent amount of diluent should be added to control diets for
untreated birds.
(C) Diets can be mixed by commercial, mechanical food
mixers. For many test substances, it is recommended that treated
diets be mixed under a hood. Mashes and test substances should
be mixed freshly just prior to the beginning of the test. For
cetain volatile or other test substances, the Test Rule may
require preparation of fresh diets at frequent intervals.
Analysis of the diet for test substance concentrations is
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required under paragraph (c)(6)(ii) of this section.
(D) Clean water should be available ad libitum. Water
bottles or automatic watering devices are recommended. If water
pans or bowls are used, water should be changed at least once a
day.
(2) Facilities. (i) Tests should be conducted with birds
being maintained in commercial brooder pens or pens of similar
construction. Pens should be constructed of galvanized metal,
stainless steel, or perfluorocarbon plastics. Materials that are
toxic, may affect toxicity, or may sorb test substances should
not be used. Wire mesh should be used for floors and external
walls; solid sheeting should be used for common walls and
ceilings. Wire mesh for floors should be fine enough so as to
not interfere with the normal movement of young birds. Pens for
housing ten young birds should have a floor area of at least 3000
square centimeters (approximately 500 square inches) for bobwhite
and 6000 square centimeters (approximately 1000 square inches)
for mallards and should be at least 24 centimeters (approximately
9.5 inches) high. Pens should be disassembled (if feasible) and
should be cleaned thoroughly between tests. Steam cleaning of
cages is recommended. Cages may be brushed thoroughly, as an
alternative method. The use of detergents or bleach is
acceptable, but other chemical disinfectants such as quaternary
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ammonium compounds should not be used. When necessary to control
disease vectors, hot or cold sterilization techniques are
recommended, as long as such techniques will not leave chemical
residues on the cages. For cold sterilization, ethylene oxide is
recommended. Pens should not be cleaned during a test.
(ii) Pens should be kept indoors to control lighting,
temperature, and other environmental variables. Pens should be
heated, preferably by thermos tatic control. A temperature
gradient in the pen of approximately 35°C to approximately 22°C
will allow young birds to seek a proper temperature. Temperature
requirements for young birds typically decline over this range
from birth through the first several weeks of life. Relative
humidity is not as critical, but the test room should be
maintained at a relative humidity of 45-70 percent. A
photoperiod of 14 hours light and 10 hours dark is recommended.
Other light/dark cycles should not be used, but continuous
lighting is acceptable. Lighting may be either incandescent or
fluorescent. Pens and lights should be positioned so that all
pens will receive similar illumination. The facilities should be
well ventilated.
(iii) Where feasible, it is recommended that pens not be
stacked upon each other. If pens are stacked, only one test
substance is allowed in any single stack. If a test substance
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volatilizes or otherwise forms aerosols or vapors in the air, no
more than one test substance should be tested in a room in order
to avoid cross-contamination. Pens should be randomly arranged,
whether or not in a stack, with respect to dose levels and
controls. Pens, such as stacked, unmodified, commercial pens
with external feeders, that allow food to be spilled from one pen
to a lower pen, should be avoided. Any modifications that
prevent cross contamination of concentration levels are
acceptable. For example, commercially available, 30 cm (one
foot) long chick feeders may be placed inside the pens and be
covered with 1.27 cm (0.5 inch) mesh hardware cloth over the
food, for bobwhite. The same feeders covered with approximately
2.5 cm (one inch) mesh wire are appropriate for mallards. For
either species, external feeders can be covered with the
appropriate size wire mesh and a solid piece of metal extended
from the bottom of the cage to a point exterior to the feeder.
Spillage may occur, but the added metal will prevent food from
spilling into another feeder.
(3) [Reserved]
(e) Reporting. (1) The test report should include the
following information:
(i) Name of test, sponsor, test laboratory and location,
principal investigator(s), and actual dates of beginning and end
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of test.
(ii) Name of species tested (including scientific name), age
of birds (in days) at the beginning of the test, average body
weights for birds in each pen at the beginning of the test, the
end of the exposure period, and end of the test, and individual
weights of all birds that die during the test.
(iii) Description of housing conditions, including type,
size, and material of pen, pen temperatures, approximate test
room humidity, photoperiod and lighting intensity.
( iv) Detailed description of the basal diet, including
source, diluents (if used), and supplements (if used). A
nutrient analysis of the diet should be included in the test
report.
(v) Detailed description of the test substance including its
chemical name(s), source, lot number, composition (identity of
major ingredients and impurities), and known physical and
chemical properties that are pertinent to the test (e.g.,
physical state, solubility, etc.).
(vi) The number of concentrations used, nominal and (where
required) measured dietary concentration of test substance in
each level, assay method used to determine actual concentrations,
number of birds per concentration and for controls, and names of
toxicants used for positive controls (if applicable).
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(vii) Acclimation procedures and methods of assigning birds
to tes t pens .
(viii) Frequency, duration, and methods of observation.
( ix) Description of signs of intoxication and other abnormal
behavior, including time of onset, duration, severity (including
death), and numbers affected in the different dietary
concentrations and controls each day of the test period.
(x) Estimated food consumption per pen daily or every other
day in the second highest and second lowest concentration and
control pens. For other pens, food consumption should be
estimated for the exposure period and for the post-exposure
period.
(xi) Location of raw data storage.
(xii) Results of range finding tests (if conducted).
(xiii) The calculated LC50 value, 95 prcent confidence
limits, slope of the concentration-reponse curve, the results of
the goodness-of-f it test (e.g., chi-square test), and a
description of statistical methods used. The same statistics for
positive controls (when used). The methods used for statistical
analysis should be described completely.
(xiv) Anything unusual about the test, any deviation from
these procedures, and any other relevant information.
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(2) In addition to the above information required in every
report, the following information should be available upon
reques t:
(i) A general description of the support facilities.
(ii) A description of the Quality Control/Quality Assurance
program, including the Average Quality Level for the program
element performing the test, procedures used, and documentations
that these levels have been achieved.
(iii) The names, qualifications, and experience of personnel
working in the program element performing the test, including
the study director, principle investigator, quality assurance
officer, as well as other personnel involved in the study.
(iv) Standard operating procedures for all phases of the
test and equipment involved in the test.
(v) Sources of all supplies and equipment involved in the
test.
(vi) Originals or exact copies of all raw data generated in
performing the test.
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TECHNICAL SUPPORT DOCUMENT
FOR
AVIAN DIETARY TEST
OFFICE OF TOXIC SUBSTANCES
OFFICE OF PESTICIDES AND TOXIC SUBSTANCES
U.S. ENVIRONMENTAL PROTECTION AGENCY
WASHINGTON, D.C. 20460
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Table of Contents
Subject Page
I. Purpose 1
II. Scientific Aspects 1
General 1
Issues 7
Test Procedures 3
Range Findings and Definitive Dose Levels 8
Controls 10
Number of Animals Tested 12
Duration of Test 14
Observations and Measurements 15
Required Analysis 16
Statistical 16
Test Substance Concentration 19
Basal Diet 20
Acceptability Criteria 20
Test Conditions 21
Test Species 21
Selection 21
Maintenance of Test Species 26
Acclimation 26
Diet 26
Feeding 27
Facilities 29
Environmental Conditions 33
Temperature (See Section II.C.2) 33
Humidity (See Section II.C.2) 33
Light (See Section II.C.2) 33
Reporting 33
III. Economic Aspects 34
IV. References 35
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TEST SUPPORT DOCUMENT FOR AVIAN DIETARY TEST
I. Purpose
The purpose of this document is to provide the
scientific background and rationale used in the development
of Test Guideline EG-15 which uses Bobwhite and Mallard to
evaluate the toxicity of chemical substances in the diets of
these two species. The Document provides an account of the
scientific evidence and an explanation of the logic used in
the selection of the test methodology, procedures, and
conditions prescribed in the Test Guideline. Technical
issues and practical considerations are discussed, In
addition, estimates of the cost of conducting these tests
are provided.
II. Scientific Aspects
A. General
Investigations of the dietary toxicity of chemicals to
native birds began when the Fish and Wildlife Service was
directed by Public Law 85-582 to evaluate and report upon
the effects of pesticides on wildlife. As a result of this
law, DeWitt et al. (1962) reported dietary toxicities of
various pesticides to mallards and bobwhite exposed to
treated diets at the Patuxent Wildlife Research Center
(PWRC). Tests were conducted for less than 10 or less than
100 days with one objective being the determination of the
quantity of pesticide producing at least 50% mortality
within 10 or between 10 and 100 days (DeWitt et al. 1963).
The shorter period was designed to simulate acute toxicity,
whereas the longer period was to simulate chronic
toxicity. Tests were conducted with adult and/or young
bobwhite, mallard, and ring-necked pheasant.
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Similar studies were conducted at PWRC in 1963 by
Stickel (1964), who reported at this time that young birds
used in dietary tests were 1-2 days old. Japanese quail
were also used as a test species beginning in 1963. The
1963 tests were from 3 to 185 days in length.
Heath and Stickel (1965) presented a protocol for avian
dietay tests. This protocol recommended a test period of 5
days exposure to a treated diet followed by 3 days of
observation while the birds received untreated diets. The
standardized length of the test period was designed to
permit quantitative comparisons of the relative toxicities
of different test substances. This protocol also
recommended that the birds be 5-7 days in age when first
exposed to treated diets.
This protocol was generally followed by PWRC for 10
years, although some deviations were occasionally necessary
because of shortages of facilities or birds (Hill et al.
1975). Hill et al. (1975) reported that the ages of the
test birds were standardized in 1970. Quail (both bobwhite
and Japanese quail) were tested when they were 14 days old;
mallards and pheasants were 10 days old. In 1973 tests,
mallards were five days old. Heath et al. (1972) did not
specify the ages of test birds precisely, but they did state
that dosage was never initiated before birds were nine days
old.
EPA adopted the basic protocol described by Heath and
Stickel (1965) and Heath et al. (1972) and published it as a
test method to be used in conjunction with pesticide
registration (US EPA 1975). The EPA protocol was intended
primarily for bobwhite and mallard and specified the use of
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10-15 day old birds. In other respects, the EPA protocol
generally followed that developed by PWRC. The Agency (US
EPA 1978a) published a later protocol that was slightly
modified from the first EPA protocol. This later protocol
specified, as a guideline, that birds be 10-17 days old at
the beginning of the test period and also recommended a
continuous lighting regime. Other modifications were minor.
Most avian dietary tests conducted since 1965 have more
or less followed either the Heath and Stickel (1965)
protocol or one of the EPA protocols. The largest number of
avian dietary tests have been conducted for the purpose of
pesticide registration and have been classified as
confidential business information by the sponsors. PWRC has
conducted the vast majority of avian dietary tests that have
been published in the open literature (Heath et al. 1972,
Hill et al., 1975). Very recently, ASTM (1979a) has
developed a draft protocol for avian dietary tests that also
is based on the PWRC method.
Because all known avian dietary test protocols are based
upon the method of Heath and Stickel (1965), there has been
no need to compromise between widely divergent methods. At
the same time, there have been numerous deviations from the
basic method that preclude precise comparisons of all of the
data. There continue to be conflicts of ideas among
investigators conducting avian dietary tests. Yet, there
are few published data to support either side of many
conflicts. Also, there is little ongoing research that
might resolve these conflicts. PWRC does conduct methods
research to some extent, but reports from most other avian
dietary test facilities usually are classified as
conf identicil.
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To promote uniformity and comparability of tests, some
recommendations and requirements are standardized in this
test guideline. Where such recommendations and requirements
are controversial and are not addressed sufficiently by
published data, it is hoped that research will be stimulated
to resolve questions. If feasible, conditions and
procedures that approximate natural conditions have been
specified in preference to other options. Controversial
points are addressed in section 1.2 of this support
document.
The use of avian dietary tests in the assessment of
chemical impacts on the environment is based on several
factors. First, birds are an obvious and important
component of the environment. Congress has indicated
repeatedly that birds are worthy of protection by passing
such laws as the Lacey Act of 1900, Migratory Bird Treaty
Act of 1918, Migratory Bird Conservation Act of 1929,
Pittman-Robertson Act of 1937, Fish and Wildlife Act of
1956, Endangered Species Act of 1973, and others. The
United States also has entered into treaties with Great
Britain and Canada (1916), Mexico (1937), Japan (1974), and
Russia (1976) for the protection of migratory birds. The
people of the United States have also indicated a desire to
protect birds through their support of Audubon Society,
Nature Conservancy, and other environmental groups.
Sportsmen's organizations support protection of birds,
although their interests often focus heavily on game birds.
Second, birds have a definite economic importance.
Federal and State Agencies spend large sums for the
preservation and propagation of birds. Hunters and
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birdwatchers also spend substantial sums in pursuit of their
pastimes. Less measurable, but of definite importance, is
the substantial role of birds in insect control.
Birds consume huge quantities of insects and other
invertebrates, many of which are considered pests. Small
mammals and other vertebrates or plants are consumed by
various birds, sometimes to the extent that birds have an
important effect on populations. In turn, birds are
consumed by birds of prey, mammals and other vertebrates.
Excretory products of birds provide nutrients for plankton
and other microorganisms that in turn are food for larger
organisms. Birds are important in pollination of some
plants and in dispersal of others. Because of their
mobility, the effects of or on birds are not restricted to
specific locations.
Finally, birds are among the more sensitive terrestrial
vertebrates. Because of their high metabolic rate, high
body temperature, and the demands of flight, they require
more energy relative to their size than most other
animals. The energy requirements lead to greater food
intake and thus to greater toxicant intake when a toxicant
is in or on their food. Data presented by Kenaga (1979)
indicate that for a majority of insecticides, mallards are
more sensitive than rats in acute oral tests. No data were
presented for acute oral toxicity to bobwhite, but in
dietary tests, bobwhite are typically more sensitive than
mallards (Heath et al. 1972). These data and confidential
reports submitted to the Agency for pesticide registration
suggest that, in general, birds are the most sensitive class
of terrestrial vertebrates.
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Avian dietary tests are useful in assessing the hazard
and risk of toxic substances to avian species. They are
relatively inexpensive and of short duration. Since
ingestion is the most likely route of avian exposure,
dietary tests can be used to estimate the effects of short-
term exposures. When a concentration-response curve is
generated, as is required in this guideline, the curve can
be used to estimate a probable no-effect level and to obtain
indications of chronic effects. The results can be used to
compare toxic responses between or among species or of one
species to various test substances. The results of
laboratory dietary studies can be extrapolated to field
conditions and/or to other species. All such comparisons
and extrapolations should be made with extreme caution,
taking into account a number of physiological, ecological,
and behavioral parameters.
Avian dietary tests are different from avian acute oral
tests. Acute oral tests involve a single dose, usually
administered by capsule or gavage directly into the crop.
Amounts are varied according to the weight of the bird, so
that each bird receives the same dose on a mg/kg basis.
Dietary tests, on the other hand, involve ingestion of a
test substance that is incorporated into the diet. Because
it is not practical to house birds in individual cages, food
consumption, and thus test substance intake, cannot be
measured individually but rather are determined on an
average basis for each pen. Some birds may eat more or less
than the average. In addition some food is spilled, and
some birds may reduce consumption because of aversion to the
food and test substance or because of toxic effects of the
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test substance that they have ingested. Therefore, the
precise amount of test substance taken in cannot be
determined and may vary among the test birds. As a result,
the LCc-Q values determined in dietary tests are not as
precise as LD50 values obtained in acute oral tests.
However, dietary tests do simulate exposure under natural
conditions. They take into account not only the route of
exposure, but also individual variation in intake, aversion,
reactions to test substance and, to a limited extent,
detoxification of the test substance. Acute oral tests have
definite value, particularly in comparative toxicology or in
assessing hazards of such concentrated toxicants as granular
insecticides. But it is the dietary test that is most
useful in assessing the hazard and risk of most test
substances to birds.
1. Is s ue s
The avian dietary test guideline and support document
contain some controversial points. Data are insufficient or
absent to support either side of most points. For other
points, there are data to support each side, or the
controversy may relate to the use of the test. Issues are
merely identified below and are discussed in the appropriate
sections of this document.
o Is continuous lighting more appropriate than
dark/light cycles, or should either be considered
acceptable?
o What is the most appropriate age for test birds?
How narrowly should the age be defined?
o Is a positive control group(s) necessary for each
test?
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o What carriers, if any, should be used or allowed
for incorporation of the test substance into the
diet?
o Are commercial foods adequate, or is there too much
variation and/or contamination of commercial foods?
o Should treated diets be mixed daily with the same
or decreasing concentrations, or should diets be
mixed fresh only at the beginning of the test?
B. Test Procedures
1. Range Finding and Definitive Dose Levels
The range-finding test is a highly recommended procedure
(US EPA 1978a, ASTM 1979a) which helps to determine the
appropriate dietary concentrations in the definitive test.
In some situations there may be enough toxicity information
available so that appropriate concentrations can be selected
without a range-finding test. The range-finding test (or
other available information) needs to be accurate enough to
ensure that dose levels in the definitive test are spaced so
as to result in at least one level each above and below the
LC50 value.
Unless otherwise specified in the Test Rule, it is
necessary to establish dose levels in the definitive test
that will result in an accurate determination of the slope
of the log dose-probit response curve. The slope and shape
of this regression curve are of great interpretative value
in analyzing the results of the test (Tucker and Leitzke
1979). Among other things (refer to discussion on
statistical interpretation), the slope and mortality pattern
may yield an indication of chronic effects. A shallow slope
suggests chronic effects whereas a steep slope suggests
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acute effects (Tucker and Leitzke 1979). Because this avian
dietary test directly investigates short term effects, it is
more important that a shallow slope be well defined than it
is for a steep slope. A slope can be defined well by
spacing concentrations to yield several partial mortality
levels, although individual variation in response, for any
reason, will tend to reduce precision of results. Ideally,
three or more partial mortality levels will give a
reasonable indication of the precision and accuracy of the
slope. Therefore, close spacing of concentrations is
recommended to yield these partial mortality levels. When a
spacing factor of 1.67 is used, as is recommended in the
guideline, and does not result in three partial mortality
levels, then the slope is steep, usually greater than 6
probits per log cycle. As stated above, a steep slope does
not need to be completely defined because the test itself
predicts acute effects. Thus, it is not necessary to use
closer spacing than a 1.67 ratio. However, if spacing is
wider than recommended, then it is necessary to achieve
three partial mortality levels in order to yield a reliable
slope.
The requirement for at least five concentration levels
is also based on the need for at least three partial
mortality levels to provide for reliable probit or other
statistical analysis. In the event that five concentration
levels provides five partial mortality levels, this
situation would yield an even more reliable slope and LC5Q
value. Fewer than five levels may frequently yield
sufficient information for determination of an LC5Q value by
non-parametric statistical methods, but even more
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frequently, fewer than five levels will not yield the three
partial mortality levels necessary for an accurate slope
determination.
Some test substances are relatively non-toxic and are
unlikely to pose an acute environmental problem. In terms
of acute effects, the Patuxent Wildlife Research Center
(Heath et al. 1972, Hill et al. 1975) and the Agency's
Office of Pesticide Programs (US EPA, 1978a) have both
stated that it is unnecessary to determine an accurate
dietary LC^Q when it is found to be in excess of 5000 ppm.
Some test substances may have an LC50 greater than 5000 ppm,
but may still produce mortality at the 5000 ppm level.
Since the slope of the log dose-probit regression line is as
important as the LC^Q value in interpreting the results of a
test, when mortality does occur at 5000 ppm, it is important
to determine at what level there is no observed effect. If
the no effect level is comparatively low, such as around 500
ppm, this gives an indication of the potential for chronic
toxicity and the need for additional testing. A
comparatively high no effect level, such as 4500 ppm,
suggests a steep slope and considerably less possibility of
chronic effects. In essence, the mortality at 5000 ppm and
the no effect level give a rough indication of the slope and
therefore, range-finding dose levels of 5, 50, 500, and 5000
ppm are suggested in order to incorporate the 5000 ppm level
(relatively non-toxic) and a lower no effect level in case
there is some mortality at 5000 ppm.
2. Controls
Concurrent controls are required for every test to
assure that any observed effects are a result of ingestion
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of the test substance and not of other factors. Other
factors may include such environmental factors as
temperature, lighting, vapors, sensitive or stressed test
birds, etc,, If a diluent is used in mixing the diet, this
diluent is used also in the untreated diets at the same
concentration as it occurs in treated diets. In effect,
this results in a diluent, but no completely negative,
control. Diluent selection is based upon an assumed lack of
toxicity (e.g., water, completely evaporated acetone) and it
is not cons, idered necessary, therefore, to have an
additional negative control when a diluent control is used.
A positive control, such as one with dieldrin, is
recommended as a means of detecting temporal or
interlaboratory variation. Some researchers have advocated
using a positive control for every test and then adjusting
the test substance LC^g value by dividing that value by the
positive control LC5Q value to yield a ratio that can then
be used to compare toxicities of various test substances
(ASTM 1979a). This procedure may be of academic interest
when comparing different test substances, but it is
inappropriate for the purpose of this guideline. In
addition, the mathematical manipulation of two values that
both have limits on precision as indicated by confidence
intervals will result in a further decrease in precision
when the results are combined. Another argument has been
advanced (Heinz, personal communication) that a positive
control should still be run with every test, even if no
adjustments are made. It is assumed that the procedures
outlined in this guideline are rigorous enough so that
variation within a given laboratory will be minimal as long
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as the food, cages, and source of birds remain the same. A
quarterly or semi-annual positive control, or one performed
when the food, housing, or source of birds changes, should
provide enough information on temporal or interlaboratory
variation to preclude the need to have a positive control
with every test.
3. Number of Animals Tested
Ten birds per test concentration is the minimum number
required in this test. Other test methodologies have
recommended ten birds per level but have allowed as few as
six birds per level (ASTM 1979a, US EPA 1978a). For some,
perhaps many, test substances, six birds per level will
yield statistically valid results. However, the statistical
error of variable responses will be magnified when six birds
are used, and a significant number of tests may result in
values that are not statistically valid.
It also should be noted that the protocol recommended by
the Agency requires no less than ten birds per dose level in
avian acute oral tests required for pesticide registration
(US EPA 1978b). Since the acute oral tests inherently
control more variables than dietary tests (see section 1),
it would be inconsistent to allow fewer birds in the dietary
tests than in the acute oral tests. The vast majority of
avian dietary tests have been conducted either by Patuxent
Wildlife Reseach Center (Heath et al. 1972, Hill et al.
1975) or by contract and company laboratories for the
purpose of pesticide registration. Most of these tests have
utilized ten birds per level and only a small fraction have
used less than ten. These facilities apparently favor the
use of ten birds per level because it markedly reduces the
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number of tests that have to be rerun to achieve statistical
validity and because it is nearly as easy to maintain ten
birds in the appropriate size pen as it is to maintain six
birds in the same size pen.
Ten birds per concentration usually are adequate to
produce a valid test, although some test substances may
require more birds to accommodate excessive variation in
response. Although 20 birds per level may be considered
statistically superior to ten birds per level, there is a
substantial data base for using the latter, and most tests
using 10 birds per level have achieved statistical
significance. For the small number of tests that resulted
in heterogeneous data, it is more cost-effective to repeat a
test with more birds per level than it is to require all
tests to be conducted with more birds. The use of 20 birds
per dose level would require twice the pens, space, food,
care, handling, etc. and would significantly increase the
cost of a test. Not only would the cost be increased, but
due to limited avian testing facilities, the time between
requiring a test and beginning the test would be lengthened
(on the average) if 20 birds per concentration were
required. Some types of tests, such as aquatic tests, may
require 20 organisms per level to offset the greater number
of variables involved or the variable responses of the test
animals. It is not necessary that avian tests also require
20 birds per dose level.
Twenty birds are the minimum required, however, for
negative control groups. Many investigators use the same
number of control birds as are used in all experimental
levels tested, and this number (50-60) is required in one
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method (ASTM 1979a). Frequently, such a large control group
is used for several concurrent tests. In such a situation a
failure to achieve a control survival of 90% would mean that
all of the concurrent tests would be unacceptable. In
addition, 20 control birds, or preferably 30 or more, will
provide better data on food consumption and growth values
with which to compare performance of treated birds.
Although natural mortality of young birds usually occurs
prior to reaching the test age, natural mortality during the
test may occur, more often with bobwhite than with
mallards. Since control mortality in excess of 10%
invalidates a test, the use of 20 birds in a control will
nearly always yield sufficient survival for validity. The
use of a single extra pen of 10 control birds is a cost-
effective preventive measure that in addition yields a
statistically more powerful test.
4. Duration of Test
Acute toxicity tests with birds and mammals are normally
done on an oral single dose basis and historically the
single dose has been followed by 14 or 21 days of
observation. This was originally true of laboratory mammals
and also was adopted by those investigators conducting acute
single dose oral tests on birds (Tucker and Crabtree
1970). Avian dietary tests, however, have developed along
other lines. The duration of the test was first
standardized by the Patuxent Wildlife Research Center (Heath
and Stickel 1965). The test period was set at 5 days
dietary exposure to the test substance followed by 3 days of
observation while birds were receiving untreated diet.
Using the PWRC format as a basis, the Agency's Office of
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Pesticide Programs (US EPA 1978a) and subsequently ASTM
(1979a) both presented test methods with the same
duration. Because of this historical basis and the data
base developed from these methods, the duration of the
currently proposed dietary test is the same.
The three day post-exposure observation period was
originally included to detect chemical mortality induced
beyond the exposure period (Heath et al. 1972). It was
noted that one chemical caused a delayed mortality pattern,
and for that chemical, the observation period was extended
an additional six days. Subsequent adaptations of this
protocol (US EPA 1978a, ASTM 1979a) included statements that
the observation period "shall be extended. . .as long as test
birds exhibit toxic symptons and continue to die." In tests
submitted to support pesticide registration, however, it has
been noted that tests were rarely extended beyond eight
days, even when mortality occurred on the last day. It was,
therefore, determined that the currently proposed avian
dietary guideline should be moje definitive about the length
of the post-exposure period under certain conditions. It is
not expected that the post-exposure period will need to be
extended for most test substances.
5. Observations and Measurements
When birds are caged together, the amount of food
ingested cannot be determined for individual birds. In
addition, feed can be spilled out of the pen or scattered
among litter and droppings. Therefore, only estimates of
average consumption per pen can be determined. This
admittedly does not yield precise intake data per bird, but
the dietary test is considered an "applied" measurement that
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includes factors of ingestion and digestion in addition to
toxicity (Heath et al. 1972).
It would be desirable to have the body weights of test
birds recorded at the beginning of the test, at the end of
the exposure period, and at the end of the test. However,
weighing birds at the end of the exposure period requires
handling, and the stress on the birds, especially in
combination with test substances that produce stimulant
effects, could easily lead to anomalous results. Therefore,
body weights are required only at the beginning and at the
end of the test.
Observations of signs of intoxication are important for
several reasons. First, they obviously yield information on
the mode of action of the test substances. If there is
additional information from other sources on the mode of
action, signs of intoxication other than those known
indicate either a difference in action between the other
test animals and birds or that factors other than the test
substance are involved. The duration of signs of
intoxication may yield information on the potential for
cumulative effects or the possiblity that effects are
irrevers ible.
6. Required Analyses
a. Statistical Analysis
A coherent theory of the dose-response relationship was
introduced by Bliss (1935), and is accepted widely today.
This theory is based on four assumptions:
o Response is a positive function of dosage, i.e., it
is expected that increasing treatment rates should
produce increasing responses.
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o Randomly selected animals are distributed normally
in Gaussian fashion as to their sensitivity to a
toxicant.
o Due to homeostasis, response magnitudes are
proportional to the logarithm of the dosage, i.e.,
it takes geometrically increasing dosages
(stresses) to produce arithmetically increasing
responses (strains) in test animal populations.
o In the case of direct dosage of animals, their
resistance to effects is proportional to body mass.
Stated another way, the treatment needed to produce
a given response in a population of test animals is
proportional to the size of the animals treated.
If percent mortality is plotted as a function of the
logarithm of test solution concentration, the form of the
resulting curve will generally be sigmoidal (Casarett and
Doull 1975). The sigmoidal shape of the concentration-
response curve makes it less than ideal for predictive
purposes. It is often more useful to transform the data so
that the concentration- response curve may be represented by
a linear equation of the form:
y=ax+b
While a number of data transformations are possible, the
probit or probability unit transformation (Bliss 1934) is
the most widely accepted means of linearizing the sigmoidal
concentration-response curves (Finney 1971).
Probits are standard deviations to which the number five
has been added arbitrarily to avoid negative numbers. For
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example, the 50 percent response point is equivalent to the
median and is called probit five. An 84 percent response is
the median plus one standard deviation and is also known as
probit 6 (0+1+5=6). Likewise, a 16 percent response is the
median minus one standard deviation or probit 4 (0-1+5=4).
Once the mortality data have been transformed onto a
probit scale, a straight line can be fitted graphically
according to the methods of Litchfield and Wilcoxon (1949)
or Miller and Tainter (1944) or by calculated methods such
as that developed by Finney (1971). The LC^g value can be
determined from the regression line, and the 95% confidence
limits can be calculated. Other values (e.g., LC16, LCQ4)
also can be determined from the line along with the
corresponding confidence limits. However, the confidence
limits are narrowest at the L^Q value and wider as the LCX
value is further from the LCcQ point.
The LC^Q value itself has utility in that it provides an
indication of the toxicity of a test substance. This value
can be used to help predict the likelihood of short-term
adverse effects to animals exposed to the test substance in
the environment. It should be noted, however, that the
endpoint of this test is mortality and the exposure period
is 5 days. Thus, the LC50 value is most pertinent to
relatively short term effects.
The mortality pattern and slope (probits divided by log
cycles) of the concentration-response line have more
implications. A low (<2 probits/log cycle) slope or delayed
mortality is usually indicative of slow and often cumulative
effects, whereas a steep slope (> 6 probits/log cycle)
suggests rapidly acting effects (Tucker and Leitzke 1979).
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When the effects of a test substance are slow and
cumulative, there is a strong possibility that some effects
may occur at concentration levels well below the LC^Q value
and also that sublethal effects have a higher likelihood of
occurring in many animals. When a toxic substance
accumulates in an animal at sublethal levels, there is a
likelihood that chronic effects, such as reproductive
impairment or secondary toxicity, may occur. Significant
amounts of some toxic accumulative chemicals, such as DDE
and heptachlor epoxide, are excreted via the eggs of birds
(Stickel, 1973) and may adversely affect reproductive
success. Thus, a low slope or delayed mortality that
suggests a potential for accumulation and chronic effects
may have long lasting consequences for the populations of
exposed birds. Therefore, not only the placement, but also
the mortality pattern and slope of the concentration-
response line are important pieces of information derived
from this test.
b. Test Substance Concentrations
Samples of treated diets will be analyzed to determine
the actual levels to be used in the test. Analysis will
help to detect mathematical errors in calculating
concentrations, technicians' errors in mixing diets, and
manufacturers ' errors in determining the amount of active
ingredient in a test substance (Heinz, personal
communication). Intentional manipulations of dietary
concentrations will not normally be detected simply through
analysis, unless it can be ensured that those diets analyzed
are those diets actually used. Therefore, it is sufficient
to analyze dietary concentrations at several, rather than
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all concentration levels, with the assumption that
variations between measured and nominal concentrations will
be consistent for all levels. When volatile or labile test
substances are used, it is important to analyze
concentrations at the end of the exposure period also.
c. Basal Diet
Most testing facilities use diets prepared by commercial
feed companies. Some facilities may have a commercial
company prepare a diet to order. Normally, such diets are
supplied with a quantitative list of ingredients, and such a
list should be supplied with the test report. If there are
supplements added to the diet, a list of all supplemental
ingredients also should be submitted. Analysis of
ingredients in the basal diet is important because there are
a number of potential test substances, such as certain
metals, that may interact with components of the diet and
possibly affect the results of a test. A nutrient analysis
will allow for a better evaluation of such results. In
addition, it is possible that dietary deficiencies or
imbalance of ratios of nutrients also could affect the
results. Even though commercial companies normally supply a
nutritionally adequate diet, it is important to know the
components because no rigid requirements exist far the type
and constitution of the diet used.
7. Acceptability Criteria
A typical avian dietary test will have no control
mortality. Occasionally, a bobwhite control death may
occur, but only rarely will a mallard control death
happen. In well conducted tests, control mortality will
never (p >0.99) be greater than 10%. If control mortality
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exceeds 10%, it is very likely that it is due to improper
test conditions or procedures, such as failure of
temperature control, excessive handling, etc. It can be
assumed that factors causing control mortality will also
contribute to mortality of treated birds and result in
inaccurate results. Therefore, a test is considered
unacceptable if more than 10% of the control birds die.
C. Test Conditions
1. Test Species
a. Selection
The mallard, Anas platyrhynchos, and/or bobwhite,
Colinus virginianus, are the species to be tested. The
choice of these species is based on a number of factors.
Although no single species would satisfy all criteria for
species selection, mallard and bobwhite each have a number
of favorable attributes.
The mallard has a widespread distribution, not only in
the United States, but also in Eurasia. Such distribution
means that mallards may be exposed to toxic substances in
the environment regardless of the location of the toxic
substance. As a waterfowl, mallards may also be exposed to
toxic substances in the water, in sediments, and on land.
Because the mallard is the most abundant and widely
distributed duck in the northern hemisphere (Bellrose 1976),
it is also suitable as a native test species for many
countries belonging to the Organization for Economic
Cooperation and Development (OECD).
The bobwhite has a widespread distribution throughout
much of the United States, and it is an important part of
the avifauna, of the southern U.S. It occurs in a variety of
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errestrial habitats and is likely to be exposed to toxic
ubs tances that occur in such habitats. Because it is
idely distributed, the bobwhite represents itself for many
nvironments as well as serving as a surrogate for other
errestrial avian species.
Bobwhite and mallards also are amenable to testing in
he laboratory. They can be bred in captivity and are
eadily available from commercial sources so that testing of
hese species will not deplete wild stocks. There is
ufficient information on the nutritional/ habitat, and
ehavioral characteristics of natural populations of
allards and bobwhite in order to meet the basic nutritional
nd physical requirements of the species in the laboratory.
Mallards and bobwhite have been demonstrated to be
ensitive to challenges with toxic chemicals (Tucker and
rabtree 1970, Hill et al. 1975). In a comparison of four
pecies (mallard, bobwhite, Japanese quail, ring-necked
heasant) that all were tested with the same 39 chemicals in
ietary studies, bobwhite was the most sensitive to 47.4% of
he chemicals (Heath et al. 1972). Even though this
omparison showed bobwhite to be the species most sensitive
o the largest number of toxic chemicals, relative species
ensitivity is variable depending upon the particular test
ubstance or type of test substance. Therefore, dietary
ensitivity, an important criterion, should not be the
verriding factor in selection of species for these tests.
or example, waterfowl have demonstrated a particular
usceptibil i ty to eggshell thinning in reproductive studies
nd bobwhite have been shown to be sensitive to the
eproductive effects of a few chemicals such as toxaphene
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(DeWitt et al. 1962). Both species have been widely used in
actual and simulated field tests. It is advantageous to
select for short term tests a species which also can be used
in reproductive and field tests. This allows for more
direct comparisons between the results of two test types
than can be obtained when one species is tested in short-
term tests and another species is tested in reproductive or
field studies.
In addition, mallards and bobwhite are generally
considered to have a positive economic value. Although the
Agency is charged with the protection of all species in the
environment, the choice of an economically valuable species
for testing is appropriate to the cost-benefit or risk-
benefit analyses upon which Agency decisions are frequently
based .
Finally there is as good or better a data base for
toxicity tests with mallards and bobwhite as for any other
native avian species. This data base permits comparisons
with results of tests with other toxic substances.
If a test is to simulate toxicity to naturally occurring
populations of mallard or bobwhite, then it is important to
use birds that are phenotypically indistinguishable from
wild birds. Since many chemicals act upon specific enzymes
and enzymes are based on a genetic code, the use of birds
genotypically similar to wild birds would be desirable.
However, the determination of phenotype is a simple
observational process, whereas genotypic determination is
impractical, if possible at all. In addition, wild birds
have a degree of heterogeneity that would not be typical of
any given genotype.
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The necessity for using healthy birds is obvious since
the test is designed to determine toxicity to healthy,
typical populations. The health of birds also is important
in reducing the number of variables that limit comparisons
between tests. There are several checks in this test
guideline that help to ensure that healthy birds are used.
The use of previously untested birds not selected for
resistance and being from disease-free flocks provides a
basically healthy stock. Visual observations select out
apparently abnormal or unhealthy birds from that stock. A
final check on health is based on the birds' ability to
survive the three-day period preceding the test. The 5%
maximum mortality during this period, as established by the
Committee on Methods (1975) for effluent fish tests and as
proposed by ASTM (1979a) for avian dietary tests, allows for
the natural mortality that may occur in young birds, but
precludes the use of populations that do not demonstrate a
reasonable survival rate.
Historically, birds used in dietary tests usually have
been immature birds. Hill et al. (1975) reported dietary
toxicities of pesticides to four avian species. All birds
were between 5 and 24 days old, with 85% being 10 to 17 days
old. The only significantly repeated deviation from the 10
to 17 day age group was that 15% of the mallards were aged 5
days. Guidelines for the Agency's Office of Pesticide
Programs specify using 10 to 17 day old birds (US EPA
1978a). A large number of avian dietary tests have been
conducted with this age class, although these data are
confidential and generally not available to the public.
Because of the historical precedents and substantial data
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base for mallards and bobwhite of this age, 10 to 17 day old
birds are specified.
Certain investigators (Heinz et al. 1979) have argued
that mallards should be tested when they are five days
old. This has been proposed because five day old mallards
could not survive the test period if they refused to eat and
because it is thought that the younger birds are more
sensitive. However, it is felt that the death of birds in a
test from starvation does not approximate natural conditions
because wild birds are rarely restricted enough to have only
one food source. Further, death by starvation confounds the
interpretation of LC^Q values. With many chemicals, younger
birds do appear to be more sensitive, but age sensitivity is
not constant, even for a given class of chemicals (Hudson et
al. 1972). Because of these factors, since adult birds are
more likely to be exposed to toxic chemicals than are young
birds, and since use of younger birds is likely to result in
increased control mortality, there appears to be no
compelling argument to change the historical precedence and
use 5 day old mallards.
Randomization of test birds into the test cages prevents
biases from being introduced. Most test birds are
artificially incubated and subtle differences in incubators
can result in hatchlings from one part of the incubator
having different characteristics, such as body weight, from
hatchlings in another part of the incubator. Randomization
is done without regard to sex because young birds cannot be
sexed reliably without surgery. Since all test pens are
supposed to be treated equally in a manner to reduce
variables, it is not necessary to acclimate the birds in the
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actual pens used in the test. Birds are normally handled
after hatching or upon receipt and again when they are
weighed at the beginning of the test period. Randomization
at either of these times is preferable to handling the birds
an additional time simply for the purpose of random
ass ignment.
2. Maintenance of Test Species
a. Acclimation
Acclimation of birds to test cages and basal diet is
important so that the effects of a new environment on the
toxicity of test substances are limited. Ideally, birds
should be maintained in test cages and on basal diet from
the time of hatching until the test period, However, many
test facilities purchase young birds rather than rear them
from eggs. A seven day acclimation period allows enough
time for adaptation and permits these facilities to purchase
birds if such facilities cannot rear their own.
b. Diet
There are few data on the nutritional requirements of
mallards or bobwhite. This subject is being investigated
under a current contract and is a proposed research need.
At the present time, commercial starter mash is recommended,
based upon historical precedents (Hill et at. 1975, US EPA
1978a, ASTM I979a) and a lack of data that support
alternatives. Changes may be made in the future when
additional data become available.
There are several different ways of considering the test
concentrations with respect to degradation and actual amount
of test substance present. One view is that the test
substance should be mixed freshly with the diet every day so
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that a constant level is achieved. A second is that the
diets should be mixed only at the beginning of the test in
order to simulate natural degradation in the environment. A
third view holds that diets should be freshly mixed each
day, but in decreasing concentrations that simulate natural
degradation. The second alternative above has been used in
most avian dietary tests in the past. Although degradation
rates in food may differ from rates in the environment,
there are differences in environmental degradation rates
among the various compartments (soil, on plants, in plants,
etc.). If there were a single degradation rate, the third
alternative would offer advantages, possibly enough
advantage to counterbalance the additional cost of mixing
diets daily. The first alternative is more costly and would
not simulate natural conditions unless release of the test
substance was continuous. Since the history and previous
data are based on the second alternative and since this
alternative does simulate degradation to an extent, the test
gu.ideline specifies that diets should be mixed once at the
beginning of the test (unless otherwise specified in the
Test Rule). If degradation is expected to be substantial
(greater than 25%), then concentrations should be analyzed
at the end of the exposure period in order to ascertain the
extent of degradation.
c. Feeding
When unmodified commercial brooder pens with external
feeders are used, food spillage is likely to occur and some
of the spilled food may fall into the feeders of lower pens
in a stack. This could result in changes of test substance
concentrations, and in some cases could significantly affect
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the outcome of the test. A stacking arrangement of control
pens on top with doses in lower pens being progressively
higher toward the bottom of the stack has been used in the
past in order to minimize the effects of spillage. Such an
arrangement would provide the least amount of dosage change,
but the resultant errors would tend to make a test substance
appear somewhat safer than would occur without any
spillage. In the majority of tests, the effects of spillage
would probably be negligible. However, there are means to
eliminate the effects of spillage.
The easiest method to avoid effects of food spillage is
to not stack pens, however, this is usually impractical for
reasons of space conservation. An effective method of
minimizing spillage is to cover the feeding trays with
hardware cloth or wire mesh of a size appropriate to the
species being tested. Although this will reduce spillage, it
will not eliminate it. Patuxent Wildlife Research Center has
had good success by using 30 cm long commercial chick
feeders and covering these with wire mesh (Hill, personal
communication). These feeders are placed inside the pens
and thus protected from spillage into other feeders.
Another method for external feeders is to extend a solid
piece of metal out from the pen (perhaps attached to the
litter pan) to a point outside the feeder. Spillage may
occur, but it will not fall into a lower tray. In addition
to avoiding cross-contamination of concentration levels, the
wire mesh over the feeder will help to provide accurate food
consumption data.
Cross-contamination of two or more different test
substances is avoided by requiring that only one substance
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be used in any one stack of pen.
3. Facilities
Bobwhite dietary tests generally have been conducted in
galvanized metal pens; mallard tests have been conducted
either in galvanized metal pens or in wooden pens on straw-
covered concrete slabs (Heath et al. 1972, Hill et al.,
1975). The wooden pens were outdoors but were
weatherproofed. The Agency's pesticide guidelines (US EPA
1978a) recommended commercial brooder units of wire mesh and
galvanized sheeting, but no mention was made of outdoor
pens. ASTM (1979a) stated that pens are best kept indoors
where lighting, temperature, and other factors may be
controlled. They recommended commercial brooder units of
wire mesh and galvanized sheeting, but suggested that other
materials would be adequate if they can be kept clean. The
suggested construction materials were stainless steel,
galvanized steel, and perfluorocarbon plastics. Materials
that are toxic, capable of adsorbing test substances, or
tnat alter or otherwise affect toxicity should not be used
(ASTM 1979a).
For this test guideline, indoor pens were selected
because they provide for control of more variables and
contribute to the reproducibility of results. Outdoor pens,
even when "weatherproofed", would result in test birds being
exposed to different seasons, light regimes, temperature,
disturbances, etc. In effect, they provide a substantially
different environment. Therefore, the results of tests run
in outdoor pens are not legitimately comparable with the
results of tests run indoors. In addition, mallard tests
conducted in 'outdoor pens limit the utility of comparisons
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with bobwhite and other species that are normally tested
indoors (Heath et al. 1972, Hill et al. 1975).
The effect of using various construction materials has
not been documented. The original use of galvanized metal
pens was based on what was available when the tests were
developed. Galvanized metal is inexpensive and readily
cleaned. Stainless steel is expensive, but is easier to
clean than galvanized metal. There is some concern that
studies with certain metals, such as zinc or nickel, could
be affected by using galvanized metal or stainless steel,
respectively (ASTM 1979b). Therefore, ASTM protocols
(1979a, 1979b) include the possibility of using
perfluorocarbon plastics (e.g., Teflon). Unfortunately, the
perfluorocarbon plastics are not a sound structural material
and have a tendency to warp and bend. Thus, cages
constructed primarily of such materials are not particularly
desirable. However, perfluorocarbon plastics may be used to
coat other structural materials. Such pens should be
examined regularly to ensure that the coating has maintained
its integrity and has not exposed the underlying structural
material. At the present time, it seems appropriate to
allow the use of all three materials, although
perfluorocarbon plastics are not recommended for structural
parts.
The space requirements for birds are based on commercial
pens that are commonly in use. Although no native birds are
considered, ILAR (1978) specified 1451.7 square centimeters
for individual pigeons and 232.3 square centimeters for
Coturnix quail and for chickens under 500 grams weight. The
sizes specified in the test rule are somewhat larger. The
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larger size is expected to be less stressful because it
allows some extra space appropriate to the interaction of
test birds that are housed in groups. ILAR (1978) is also
oriented towards laboratory animals that may be inbred to
the point of tolerating more crowded conditions. Since the
avian dietary test is designed to kill birds, extra space
over that used for general laboratory animals will reduce
the potential for stress from overcrowding to affect the
results of the test.
Pens should be cleaned and sanitized between tests.
Brushing and/or steam cleaning appear to be the most
appropriate since they do not involve the use of chemicals
that could affect subsequent tests. Detergents and bleach
have been used by Denver Wildlife Research Center (Tucker,
personal communication) and Patuxent Wildlife Research
Center (Heinz, personal communication). The use of chemical
disinfectants, such as quaternary ammonium compounds, should
be avoided because of possibility that these compounds can
leave toxic residues. However, the widely used cold
sterilization method with ethylene oxide is acceptable, if
needed for disease control. Pens should not be cleaned
during a test in order to minimize disturbance to the test
birds.
The choice of temperatures, humidity levels, and
lighting regimes has been based upon historical precedents
(Heath et al. 1972, Hill et al. 1975, US EPA 1978a, ASTM
1979a) and a lack of any proposed alternatives. A
completely rcindomized cage arrangement is required. Pens
are usually stacked on top of each other to enhance the
utilization of space. A temperature gradient is strongly
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recommended because the thermoregulatory ability of young
birds varies among individuals and changes with age. The
use of a gradient gives the young birds temperature options
that are appropriate at varying times in their development
of thermoregulatory ability.
The protocols developed by Patuxent Wildlife Research
Center (Heath and Stickel 1965, Heath et al. 1972, Hill et
al. 1975) did not specify any particular photoperiod. The
Agency's Office of Pesticide Programs (US EPA 1975) stated
that photoperiod may follow diurnal variation or be
continuous for 24 hours per day. The next revision of the
Pesticide guidelines (US EPA 1978a) stated a preference for
continuous lighting but accepted a diurnal variation. ASTM
(1979a) considered continuous lighting to be "probably
optimum" but stated that a schedule of 14 hours light and 10
hours dark (14 L/10D) is acceptable. For this avian test
guideline, a 14L/10D lighting schedule is recommended
because it approximates natural lighting. A photoperiod
that approximates the natural light regime is expected to be
less stressful than exposure to constant lighting.
Consideration was given to requiring, rather than
recommending, a light-dark cycle. However, a lack of
published data on the effects of continuous lighting versus
a diurnal variation in young birds, along with a data base
using continuous lighting (Hill et al. 1975) suggests that
constant lighting should not be prohibited at this time.
Research on this subject has been recommended in order to
provide the Agency with additional information.
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4. Environmental Conditions
a. Temperature (See Section II C.3)
b. Humidity (See Section II C.3)
c. Light (See Section II C.3)
D. Reporting Requirements
The information that is required to be reported in
section II.B 6a, Statistical Analysis is essential to a
proper evaluation of the test results. These required items
are needed (1) to establish that the test was conducted
according to specifications, (2) to evaluate those
conditions and procedures that could affect the results of
the test, and (3) to supply the Agency with sufficient
information to conduct an independent analysis of statistics
and conclusions. The location of the raw data storage will
allow the Agency to find additional information that may
have been left out of the report or that may be needed for
enforcement purposes. The location is necessary because
some chemical companies request the testing facility to keep
these data, while other companies keep their own. The
information is needed in a detailed manner because the avian
dietary guideline contains few rigid requirements. Even
when minimuns or maximums are specified, it is important to
know how much the test may have exceeded specifications,
such as; if test birds were observed more frequently than
required, if the number of test concentrations exceeded the
five levels required, etc.
The information required in section II.B.6.b to be
available, but not included in the test report, may be
needed if there are serious concerns about the results or
validity of the test. This information will not normally be
33
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August, 1982
needed and therefore is not required routinely.
III. Economic Aspects
Three laboratories were surveyed to estimate costs for
performing the test outlined in this guideline. The
individual laboratories gave prices of $1850-2600, $850-
1200, and $1500. The "best estimate" based upon the survey
was $1510 for bobwhite and $1640 for mallards. A cost
estimate also was made by separating the protocol into
components and estimating the cost of each component,
including direct labor cost, overhead cost, other direct
cost, general and administrative cost, and fee. The best
estimated final cost, based upon this calculation method,
was $1973 for bobwhite and $2785 for mallards, with an
estimated range of +_ 50% of the best estimate. The
calculated estimate is higher than the best estimate based
on the survey. Differences in estimated prices or prices
obtained from the different laboratories may have resulted
from a number of factors, such as nature of the chemical,
overhead rates, outside consultants, automation, marketing
strategies, and other factors as outlined in a cost analysis
report by Enviro Control (1980). The cost estimates were
made assuming that the requirements of the Good Laboratory
Practice Standards, as specified in Section (d) of the avian
dietary toxicity guideline, are being satisfied.
In a cost analysis of subpart E pesticides guidelines,
laboratories were surveyed in 1978 and in 1980 to determine
the cost of testing (US EPA 1980a). The cited costs did not
differentiate between species, however, the unit cost for an
avian dietary LC5Q test was $2000 in 1978 and $3600 in 1980.
34
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August, 1982
IV. References
ASTM. 1979a. American Society for Testing and
Materials. Standard practice for conducting LC50 tests
with birds. Draft No. 1. 13 pp.
ASTM. 1979b. American Society for Testing and
Materials. Standard practice for conducting
reproductive studies with birds. Draft No. 1. 20 pp.
Bellrose FC. 1976. Ducks, Geese & Swans of North
America. Stackpole Books, Harrisburg, Pa. 543 pp.
Bliss CI. 1934. The method of probits. Science 79:
38-39.
Bliss CI. 1935. The calculation of the dosage-mortality
curve. Ann. Appl. Biol. 22: 134-307.
Casarett LJ, Doull J. 1975. Toxicology, the Basic
Science of Poisons. Macmillan Publishing Co., New
York. 768 pp.
Committee on Methods for Toxicity Tests with Aquatic
Organisms. 1975. Methods for acute toxicity tests with
fish, macroinverteb rates, and amphibians. Ecological
Research Series No. EPA-660/3-75-009. U.S.
Environmental Protection Agency, Duluth, Minn. 61 pp.
DeWitt JB, Crabtree DG, Finley RB, George JL. 1962.
Effects on wildlife. Pp. 4-10 (+Tables) in USDI,
Effects of Pesticides on Fish and Wildlife: A Review of
Investigations during 1960. Bureau Sport Fish. Wildl.
Circ. No. 143. 52 pp.
Wildlife
pp. 74-96
DeWitt JB, Stickel WH, Springer PF. 1963.
studies, Patuxent Wildlife Research Center.
In USDI, Pesticide-Wildlife studies: a review of Fish
and Wildlife Service investigations during 1961 and
1962, Fish and Wildl. Serv. Circ. No. 167. 109 pp.
Enviro Control, Inc. 1980. Cost analysis methodology
and protocol estimates: ecotoxicity standards.
Rockville, MD: Enviro Control, Inc., Borriston
Laboratories, Inc.
35
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ES-12
August, 1982
Finney DJ. 1971 Probit Analysis/ 3rd ed. Cambridge
University Press, London. 333 pp.
Heath RG, Stickel LF. 1965. Protocol for testing the
acute and relative toxicity of pesticides to penned
birds, pp. 18-21 in USDI. Effects of Pesticides on
Fish and Wildlife. Fish and Wildl. Serv. Circular
226. 77 p.
Heath RG, Spann JW, Hill EF, Kreitzer JF. 1972.
Comparative dietary toxicities of pesticides to birds.
U.S. Fish and Wildlife Service, Spec. Kept. Wildl. No.
152. 57 pp.
Heinz GH, Hill EF, Stickel WH, Stickel LF. 1979.
Environmental contaminant studies by the Patuxent
Wildlife Research Center, pp. 9-35. in Kenago, EE
(ed). Avian and Mammalian Wildlife To"xTcology, STP 693,
ASTM, Philadelphia. 97 pp.
Hill EF, Heath RG, Spann JW, Williams JD. 1975. Lethal
dietary toxicities of environmental pollutants to
birds. U.S. Fish and Wildl. Serv., Spec. Sci. Rept.
Wildl. No. 191. 61 pp.
Hudson RH, Tucker RK, Haegele MA. 1972. Effect of age
on sensitivity: acute oral toxicity of 14 pesticides to
mallard ducks of several ages. Toxicol. Appl.
Pharmacol. 22: 556-561.
ILAR. 1978. Institute of Laboratory Animal Resources,
National Research Council. Guide for the care and use
of laboratory animals. U.S. Department of Health,
Education, and Welfare Publication 78-23. 70 pp.
Kenaga EE. 1979. Acute and chronic toxicity of 75
pesticides to various animal species. Down to Earth 35
(2): 2531.
Litchfield JT, Jr, Wilcoxon F. 1949. A simplified
method of evaluating dose-effect experiments. J.
Pharmacol. Exp. Therap., 96(2): 99-133.
Miller LC, Tainter ML. 1944. Estimation of the ED50
and its error by means of logarithmic-probit graph
paper. Proc. Soc. Exp. Biol., 57: 261-264.
36
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ES-12
August, 1982
Stickel L. 1964. Wildlife Studies, Patuxent Wildlife
Research Center, pp. 77-115 in USDI, Pesticide-wildlife
studies, 1963: a review of FisTT and Wildlife Service
investigations during the calendar year. Fish and
Wildlife Service Circ. No. 199. 130 pp.
Stickel LF. 1973. Pesticide residues in birds and
mammals, pp 254-312 in Edward CA. (ed .). Environmental
Pollution by Pesticides. Plenum Press, London. 542 pp.
Tucker RK, Crabtree DG. 1970. Handbook of toxicity of
pesticides to wildlife. U.S. Fish and Wildl. Serv.,
Resource Publ. No. 84. 131 pp.
Tucker RK, Leitzke JS. 1979. Comparative toxicology of
insecticides for vertebrate wildlife and fish.
Pharmac. Ther. 6:167-220.
U.S. Environmental Protection Agency. 1975. Protocol
for determing lethal dietary concentration of chemicals
to birds. Federal register, 40 CFR 162.82 (appendix):
26915. June 25, 1975.
U.S. Environmental Protection Agency. 1978a. Avian
dietary LC50. Federal Register, 40 CFR 163.71-2:29727-
29728. July 10, 1978.
U.S. Environmental Protection Agency. 1978b Avian
single-dose oral LD50. Federal Register, 40 CFR 163.71-
1: 29726-29727. July 10,1978.
U.S. Environmental Protection Agency. 1979. Toxic
substances control. Discussion of premanufacture
testing policy and technical issues; request for
comment. Federal Register 44: 16240-16292. March 16,
1979.
U.S. Environmental Protection Agency. 1980a. Cost
analysis: Guidelines for registering pesticides in the
United States, Subpart E. Draft. May 1980.
U.S. Environmental Protection Agency. 1980b.
Guidelines for registering pesticides in the United
States. Subpart E. Hazard Evaluation: Wildlife and
Aquatic Organisms. Draft. November 3, 1980.
37
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August, 1982
Personal Communications:
Gary Heinz (11/16/79)
Patuxent Wildlife Research Center
Laurel, Md.
Elwood F. Hill (5/8/80)
Patuxent Wildlife Research Center
Laurel, Md.
Richard K. Tucker (11/8/79)
EPA, Office of Toxic Substances
Washington, D.C.
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EG-16
August, 1982
BOBWHITE REPRODUCTION TEST
OFFICE OF TOXIC SUBSTANCES
OFFICE OF PESTICIDES AND TOXIC SUBSTANCES
U.S. ENVIRONMENTAL PROTECTION &3ENCY
WASHINGTON, D.C. 20460
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Office of Toxic Subs tnces SG-16
Guideline for Testing Chemicals August, 1982
BOBWHITE REPRODUCTION TEST
(a) Purpose. This guideline is designed to develop data
on the reproductive effects on the bobwhite of chemical
substances and mixtures subject to chronic environmental effects
test regulations under the Toxic Substances Control Act (TSCA)
(Pub.L. 94-469, 90 Stat. 2003 15 U.S.C. 2601 _et_. s eg.) . The
Agency will use these and other data to assess the reproductive
effects on birds that these chemicals may present to the
environment.
(b) Def initions . (1) The definitions in section 3 of the
Toxic Substances Control Act (TSCA) and Part 792—Good Laboratory
Practice Standards apply here. In addition, the following
definitions apply generally to this guideline:
(i) "Acclimation" Physiological and behavioral adaptation
to environmental conditions (e.g., housing and diet) associated
with the test procedure.
(ii) "Test substance" The specific form of a chemical or
mixture of chemicals that is used to develop the data.
(iii) "Photoperiod" The light and dark periods in a 24
hour day. This is usually expressed in a form such as 17 hours
light/ 7 hours dark or 17L/7D.
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August, 1982
( iv) "Basal diet" The untreated form of the diet, such as
the diet obtained from a commercial source.
(2) The definitions in this section refer specifically to
the production and quality of eggs and the subsequent development
of these eggs up to the point where young are 14 days old.
(i) "Eggs laid" This term refers to the total egg
production during the test, which normally includes ten weeks of
laying. Values are expressed as numbers of eggs per pen per
season (or test) .
(ii) "Eggs cracked" Eggs determined to have cracked
shells when inspected with a candling lamp. Fine cracks cannot
be detected without using a candling lamp and if undetected will
bias data by adversely affecting embryo development. Values are
expressed as a percentage of eggs laid by all hens during the
test.
(iii) "Eggs set" All eggs placed under incubation, i.e.,
total eggs minus cracked eggs and those selected for analysis of
eggshell thickness. The number of eggs set, itself, is an
artificial number, but it is essential for the statistical
analysis of other development parameters.
(iv) "Viable embryos (fertility)" Eggs in which
fertilization has occurred and embryonic development has begun.
This is determined by candling the eggs 11 days after incubation
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August, 1982
has begun. It is difficult to distinguish between the absence of
fertilization and early embryonic death. The distinction can be
made by breaking out eggs that appear infertile and examining
further. This distinction is especially important when a test
compound induces early embryo mortality. Values are expressed as
a percentage of eggs set.
(v) "Live 18-day embryos" Embryos that are developing
normally after 18 days of incubation. This is determined by
candling the eggs. Values are expressed as a percentage of
viable embryos (fertile eggs).
(vi) "Hatchability" Embryos that mature, pip the shell, and
liberate themselves from the eggs on day 23 or 24 of
incubation. Values are expressed as percentage of viable embryos
(fertile eggs).
(vii) "14-day old survivors" Birds that survive for two
weeks following hatch. Values are expressed both as a percentage
of hatched eggs and as the number per pen per season (test).
(viii) "Eggshell thickness" The thickness of the shell and
the membrane of the egg at several points around the girth after
the egg has been opened, washed out, and the shell and membrane
dried for at. least 48 hours at room temperature. Values are
expressed as the average thickness of the several measured points
in millimeters.
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August, 1982
(c) Test procedures--(1) Summary of the test; (i)
After birds have been obtained they should be observed for health
and acclimated for at least two weeks.
(ii) Test birds should be randomly assigned to control and
various treatment groups.
(iii) The test substance should be thoroughly and evenly
mixed into the diet at concentrations specified in the test
rule. All treatment levels should be analyzed for test substance
concentrations at the beginning and midway through the test.
(iv) Birds should be weighed at the beginning of the test,
at 14-day intervals until the onset of laying, and at termination
of the test.
(v) Photoperiod should be carefully controlled on a short-
day basis during the initial exposure phase, then increased to
16-17 hours to induce egg laying.
(vi) Birds should be observed regularly for abnormal
behavior or mortality throughout the test.
(vii) Eggs should be removed daily and stored until there is
a sufficient quantity for incubation. All eggs should be candled
for cracks and cracked eggs removed. Once every two weeks, all
eggs produced that day should be analyzed for eggshell
thickness. Incubated eggs should be candled on day 11 and day
18. Hatching should be completed by day 24.
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EG-lb
August, 1982
(viii) Hatchlings should be maintained until they are 14
days old. Abnormal behavior or death should be reported. Chicks
should be weighed on day 14.
(ix) A statistical analysis should be performed, preferably
by analysis of variance or regression analysis.
(x) The report should include all conditions, procedures,
and results. Data should be sufficiently detailed for an
independent statistical analysis.
(xi) All treated birds should be sacrificed and disposed of
properly. Control birds may be kept as breeding stock, but
should not be used in any other tests. Control offspring may be
reared and used in another test as adults.
(2) [Reserved]
(3) [Reserved]
(4) Definitive test—(i) Test substance. (A) The
concentrations of test substance in the diet will be specified in
the test rule. Generally, three treatment groups and a control
group will be used. The higher two treatment concentrations will
be multiples (often 5x, lOx, or 20x) of the lowest treatment
level. The highest treatment levels usually will be below lethal
levels, unless predicted environmental exposure levels are high
enough to approximate lethal concentrations.
(B) The material to be tested should be analytically pure
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August, 1982
and the degree of purity should be reported along with the
percentage of each impurity at levels specified in the test
rule. If specifically required by a test rule for a particular
substance or mixture, the technical grade should be tested. The
test rule will specify the degree of purity or a range of
compositions of the technical grade material.
(ii) Controls. A concurrent control is required during
every test. The control birds should be from the same hatch as
the test groups. Control and test birds should be kept under the
same experimental conditions. The test procedures should be the
same for control and treated birds, except that no test substance
should be added to the diets of control birds. If a carrier or
diluent is used in preparation of the test diets, the same
carrier should be added to the diets of control birds in the
highest concentration used for test diets. The use of shared
controls is acceptable for concurrent tests as long as the same
carrier is used for all the tests.
(iii) Test groups and numbers of birds. (A) Each of the
three treatment groups and the control group should consist of a
minimum of 12 replicate pens. Each pen should contain one male
and one female, or alternatively one male and two females. The
use of 20 replicate pens in the control group may yield a test
with greater statistical power. Either arrangement is acceptable
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August, 1982
productivity reaches the definitive values given in paragraph
(c)(4)(xii) of this section. Because the behavioral interactions
of birds in the two arrangements is likely to be different,
testing facilities using an arrangement with which they are not
familiar are advised to experiment first without test substances
in order to determine the feasibility of obtaining acceptable
productivity levels.
(B) All control and treatment birds should be randomly
distributed to pens from the same population.
(iv) Duration of test. (A) The test consists of three
phases following acclimation to test facilities. The initial
phase begins with exposure of treatment groups to diets
containing the test substance and is typically six to eight weeks
long. After the initial phase, the photoperiod is manipulated
according to paragraph (c)(4)(v) of this section to bring the
hens into laying condition. This second phase ends with the
onset of egg-laying and is typically two to four weeks long. The
final phase begins with the onset of laying and lasts for at
least eight weeks, preferably ten weeks. A withdrawal study
period may be added to the test phase if reduced reproduction is
observed. The withdrawal period, if used, need not exceed three
we eks .
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EJG-16
August, 1982
(B) Exposure of adult birds to the test substance should be
continuous throughout the test. Unless otherwise specified in
the test rule, test birds should be exposed for at least ten
weeks prior to the onset of egg laying.
(v) Preparation for reproduction (photoperiod). (A)
Lighting regimes (photoperiod) are critical to successful
reproduction. Various photoperiod regimes have been demonstrated
to give acceptable results. Any photoperiod regime that results
in productivity that meets the definitive values given in
paragraph (c)(4)(xii) of this section is acceptable as long as
birds are exposed to treated diets a minimum of ten weeks prior
to the onset of laying. Regardless of the methods selected,
lighting should be controlled carefully. It is important during
the initial phase to not interrupt the dark period unless
absolutely necessary.
(B) A suggested photoperiod regime would consist of
maintaining birds under a photoperiod for seven or eight hours of
light during the initial phase. At the end of the initial phase,
the photoperiod may be increased to 16-17 hours of light per
day. The photoperiod may be maintained at this level for the
remainder of the study, although an increase each week of 15
minutes per day is acceptable.
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August. 1982
(vi) Observations of record on adult birds. (A) Body
weights should be recorded for each adult bird at the beginning
of the treatment period, at 14-day intervals until the onset of
egg laying,, and at termination of treatment. Recording of body
weights during egg laying is discouraged because of possible
adverse effects on egg production. Food consumption should be
measured and recorded by pen as often as body weights are
measured prior to the onset of laying and at least bi-weekly
throughout the rest of the study.
(B) Observations on adult birds should be made at least once
a day. Any mortality or other signs of toxicity should be
described and recorded by date or day of test. Gross
pathological examinations should be conducted on all birds that
die during the test period, and for all survivors at the end of
the test. Analysis of two or more tissues (e.g., muscle, fat)
for test substance residues is encouraged, but not required
(unless specified in the Test Rule).
( v i i) Egg collection, storage, and incubation. Al 1 eg gs
should be collected daily, marked according to the pen from which
collected, and should be stored at 16°C and 55-80 percent
relative humidity. Storage in plastic bags may improve
uniformity of hatching. Stored eggs should be turned daily. At
weekly or bi-weekly intervals, eggs should removed from storage
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EG-16
August, 1982
and be candled to detect eggshell cracks. Except for eggs with
cracked shells and those eggs removed for eggshell thickness
measurements, all eggs should be set after candling for
incubation in a commercial incubator. If incubators are not
equipped to automatically turn eggs, they should be turned daily
by hand. During the incubation period, eggs should be maintained
at 37.5°C and approximately 70 percent relative humidity. Eggs
should be candled again on day 11 of incubation to determine
fertility and early death of embryos. A final candling should be
done on day 18 to measure embryo survival. On day 21, eggs
should be removed to a separate incubator or hatcher. Hatching
will normally be complete by the end of day 24.
(viii) Chick maintenance. By day 24 of incubation, the
hatched bobwhite chicks should be removed from the hatcher or
incubator. Chicks should be either housed according to the
appropriate parental pen group or individually marked (such as by
leg bands) as to parental group and housed together. Chicks
should be maintained in commercial brooder pens or pens of
similar construction. Pens should be constructed of galvanized
metal or stainless steel. Temperature in the pens should be
controlled, preferably by a thermostatically control device. A
temperature gradient in the pen from approximately 35°C to
approximately 22°C will allow young birds to seek a proper
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EG-lb
August, 1982
temperature. Temperature requirements for young birds typically
decline over this range from birth through the first several
weeks of life. Chicks should be provided a standard commercial
game bird starter ration, or its nutritional equivalent. No test
substance nay be added to the diet of chicks. Chicks should be
maintained until they are 14 days old.
(ix) Observations of record on chicks. The hatchability,
percentage of normal hatchlings, percentage of 14-day old
survivors, and number of 14-day old survivors per hen should be
recorded arid reported. Chicks should be observed daily from
hatching until they are 14 days old. Mortality, signs of
toxicity, and other clinical abnormalities should be recorded at
least cumulatively through day 5 and recorded by age from days 5
through 14. Average body weights should be recorded for chicks
at day 14.
(x) Eggshell thickness. Once every two weeks all eggs newly
laid that day should be removed and measured for eggshell
thickness. Eggs should be opened at the girth (the widest
portion), the contents washed out (or used or saved for egg
residue analysis), and the shells air dried for at least 48
hours. The thickness of the shell plus the dried membrane should
be measured at a minimum of 3 points around the girth using a
micrometer calibrated at least to 0.01 mm units.
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August, 1982
(xi) Typical observed values The values reported here
represent those observed from a few testing facilities under
their conditions. These values are not necessarily
representative of those from all facilities, however, if a
reproduction test does not meet or at least approach these values
for control birds, then there is likely to be a problem with test
procedures or conditions that should be investigated and
corrected.
(A) Eggs laid. Normal values for bobwhite - 28 to 38 eggs
per hen per season.
(B) Eggs cracked. Normal values for bobwhite - 0.6 percent
to 2.0 percent of eggs laid.
(C) Viable embryos (fertility). Normal fertility values for
bobwhite - 75 percent to 90 percent of eggs set.
(D) Live 18-day embryos. Normal values for bobwhite - 97 to
99 percent of viable embryos.
(E) Hatchabili ty. Normal values for bobwhite - 50 percent
to 90 percent of viable embryos (fertile eggs).
(F) 14-day-old survivors. Normal values for bobwhite - 75
percent to 90 percent of eggs hatched.
(G) Eggshell thickness. Normal average values for bobwhite
- 0.19 mm to 0.24 mm.
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EG-lb
August, 1982
(xii) Definitive test criteria. (A) A test is unacceptable
if bobwhite chick productivity in control groups does not average
twelve 14-day old survivors per pen over a ten week period.
(B) A test is unacceptable if the average eggshell thickness
in control groups is less than 0.19 mm.
(C) A test is unacceptable if more than 10% of the adult
control birds die during the test.
(5) [Reserved]
(6) Analytical measurements — (i) Statistical analysis.
Experimental groups should be individually compared to the
control group by analysis of variance. Other accepted
statistical methods may be used as long as they are documented.
In particular, regression analysis is highly desirable if the
data and number of dose levels allow the use of this technique.
Sample units are the individual pens within each treatment level
or control. Analysis should include:
(A) Body weight of adults.
(B) Food consumption of adults.
(C) Percentage of hens laying eggs. This should always be
determined when pens contain a single pair; if feasible, it
should be determined when pens contain groups.
(D) Number of eggs laid per pen.
(E) Percentage of cracked eggs.
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August, 1982
(F) Percent viable embryos of eggs set.
(G) Percent live 18-day embryos of viable embryos.
(H) Percent hatching of viable embryos.
(I) Percentage of hatchlings that are normal.
(J) Percent 14-day-old survivors of normal hatchlings.
(K) Number of 14-day-old survivors per hen.
(L) Body weights of 14-day-old survivors.
(M) Eggshell thickness.
(ii) Test substance concentrations. (A) Samples of treated
diets should be analyzed to confirm proper dietary concentrations
of the test substance. If samples cannot be analyzed
immediately, they should be stored appropriately (e.g., frozen at
a temperature of -15°C or lower) until analysis can be
performed. Analyses should be conducted on all test substance
concentrations at the beginning of the test period and again 10
to 12 weeks later. If not otherwise available, data should be
generated to indicate whether or not the test substance degrades
or volatilizes. If the test substance is known or found to be
volatile or labile to the extent that 25 percent or more loss
occurs within one week, then test substance diets should be
prepared (freshly or from frozen concentrate) at a frequency that
will prevent more than 25 percent% loss of test substance.
14
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August, 1982
(B) The assay method used to determine actual concentrations
should be reported according to paragraph (e)(l)(vi) of this
section.
(iii) Analysis of Basal Diet. A nutrient analysis of the
basal diet should be included with the test report. For
commercially prepared basal diets, the list of ingredients
supplied by the manufacturer is normally sufficient if it is
detailed. The composition of any vitamin or other supplements
should also be reported.
(d) Test conditions — (1) Test species — (i) Selection.
(A) Bobwhite, Colinus virginianus (L.), is the test species.
Test birds should be pen-reared. They may be reared in the
laboratory or purchased from commercial breeders. Rearing stock
and/or test birds should be obtained only from sources that have
met the requirements for "U.S. Pullorum-Typhoid Clean"
classification. Birds should be obtained only from sources whose
colonies have known breeding histories. If possible, a history
of rearing practices for test birds should be obtained and made
available upon request. This history should include lighting
practices daring rearing, disease record, drug and any other
medication administered, and exact age. Test birds should be
phenotypically indistinguishable (except for size) from wild
stock. Conscientious breeders of such birds will periodically
15
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EG-16
August, 1982
outbreed their flocks with genetically wild stock in order to
maintain a genetic composition that approximates the
heterogeneity of naturally occurring birds.
(B) All control and experimental birds used in a test should
be from the same source and strain. If shipped, all birds should
be examined following shipment for possible physical injury that
may have occurred in transit. All birds should have a health
observation period of at least two weeks prior to selection for
treatment. Birds should be in apparent good health. Deformed,
abnormal, sick, or injured birds should not be used. A
population of birds should not be used if more than 3 percent of
either sex die during the health observation period. Birds
should not have been selected in any way for resistance to toxic
substances. Birds should not have been used in a previous test,
either in a control or treatment group. Offspring of birds used
in a treatment group in a previous test should not be used, but
offspring of birds used as a control in a previous test are
acceptable.
(C) Test birds should be approaching their first breeding
season and should be at least seven months old. All test birds
should be the same age within one month. The age of test birds
should be reported.
16
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BG-16
August, 1982
(D) Bobwhite should be acclimated to test facilities and
untreated basal diet for at least two weeks. Acclimation may be
in the actual pens used in the test or in identical pens. The
acclimation period may coincide with the health observation
period. Birds should be randomly assigned to treatment and
control pens. However, when birds in a pen are incompatible,
they may be rearranged within a control or treatment group at any
time prior to initiating treatment.
(E) During holding, acclimation, and testing, birds should
be shielded from excessive noise, activity, or other disturb-
ance. Birds should be handled only as much as is necessary to
conform to test procedures.
(ii) Eiiet—(A) Adult birds . (^L_) A standard commercial
game bird breeder ration, or its nutritional equivalent, should
be used for diet preparation. This ration or basal diet should
be used for both control and treatment birds and should be
constant throughout the duration of the study. Antibiotics or
other medication should not be used in the diet or water of
breeding birds. It may not be possible to obtain food that is
completely free of pesticides, heavy metals, and other con-
taminants. However, diets should be analyzed periodically for
these substances and should be selected to be as free from
contaminants as possible. A nutrient analysis (quantitative list
17
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EG-16
August, 1982
of ingredients) of the diet should be included with the test
report.
(2) The test substance should be mixed into the diet in a
manner that will ensure even distribution of the test substance
throughout the diet. If possible, the test substance should be
added to the diet without the use of a carrier or diluent. If a
diluent is needed, the preferred diluent is distilled water; but
water should not be used for test substances known to hydrolyze
readily. When a test substance is not water soluble, it may be
dissolved in a reagent grade evaporative diluent (e.g., acetone,
methylene chloride) and then mixed with the test diet. The
solvent should be completely evaporated prior to feeding. Other
acceptable diluents may be used, if necessary, and include table
grade corn oil, propylene glycol, and gum arabic (acacia). If a
diluent is used, it should comprise no more than 2 percent by
weight of the treated diet, and an equivalent amount of diluent
should be added to control diets.
(3) Diets may be mixed by commercial or mechanical food
mixers . Other means are acceptable as long as they result in
even distribution of the test substance throughout the diet.
Screening of the basal diet before mixing is suggested to remove
large particles. For many test substances, it is recommended
that diets be mixed under a hood. Frequently, the test substance
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August, 1982
is added to an aliquot of the basal diet to form a premix
concentrate. The premix concentrate should be stored so as to
maintain the chemical concentration. For final preparation of
test diets, the premix is mixed with additional basal diet to
form the proper concentrations. The frequency with which final
treated diets are prepared will depend upon the stability and
other characteristics of the test substance. Unless otherwise
specified in the test rule or determined by degradation or
volatility studies, it is recommended that final diets be
prepared weekly, either fresh or from a concentrate. For
volatile or labile test substances, test diets should be mixed
frequently enough so that the concentrations are not reduced from
initial concentrations by more than 25 percent. Analysis of
diets for test substance concentration is required as specified
in paragraph (c)(6)(ii).
(_£) Clean water should be available ad libitum. Water
bottles or automatic watering devices are recommended. If water
pans or bowls are used, water should be changed daily or more
often.
(B) Young birds . Young birds produced during the test
should be fed a commercial game bird starter ration, or its
nutritional equivalent. No test substance should be added to the
diets of young birds. No antibiotics or medication may be used
in the diet. Bacitracin, or one of its forms, may be added to
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August, 1982
the drinking water of young birds, if necessary.
(2) Facilities . (i) Bobwhite should be housed in breeding
pens or cages of adequate size conforming to good husbandry
practices. Space requirements for bobwhite have not been well
defined, but it is recommended that there be at least 5000 square
centimeters (approximately 2.7 square feet) of floor space per
bird. Documentation that reproductive parameters and health of
birds are not adversely affected should be provided for cages
much smaller than this area. The preferred construction
materials are stainless steel, galvanized sheeting, and wire
mesh. For enclosed cages, floors and external walls may be wire
mesh; ceilings and common walls should be solid sheeting. Wire
mesh for floors should be fine enough so as to not interfere with
normal movement of bobwhite. Open-topped pens may be constructed
of the same materials for the side walls with open tops and wire
mesh or concrete floors. Concrete floors should be covered witn
litter such as straw, wood shavings, or sawdust. Other
construction materials, except wood, are acceptable if they can
be kept clean. Wood may be used as vertical framing posts for
the support of wire mesh or for horizontal framing along the top
of the pen. Wood should not be used for floors or lower sides of
pens unless it has been coated with a non-adsorbent material such
as perfluorocarbon plastic (e.g., Teflon), or unless the wood is
20
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August, 1982
replaced between tests.
(ii) Pens should be disassembled (if feasible) and should be
cleaned thoroughly between tests. Steam cleaning of enclosed
cages is recommended. Enclosed cages may be brushed thoroughly,
as an alternative method. For open-topped pens, the sides and
vertical supports should be thoroughly brushed. Any used floor
litter should be discarded. The floor composition will dictate
methods used to clean the floor. If litter is used on the floor,
it should be fresh and clean when birds are placed in the pen.
The use of detergents or bleach is acceptable, but other chemical
disinfectants (such as quaternary ammonium compounds) should not
be used. Vfhen necessary to control disease vectors, hot or cold
sterilization techniques are recommended, as long as such
techniques will not leave chemical residues on the cages. For
cold sterilization, ethylene oxide is recommended.
(iii) Pens should be kept indoors in order to better control
lighting, temperature, humidity, and other factors. Outdoor pens
may be used only during the normal breeding season. The
photoperiod should be carefully controlled, preferably by
automatic timers. A 15-30 minute transition period is
desirable. The photoperiod regime is described under test
procedures under paragraph (c)(4)(v). Lights should emit a
spectrum simulating that of daylight. The use of shorter wave-
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August, 1982
length "cool-white" fluorescent lights that do not emit the
daylight spectrum should be avoided. Illumination intensity
should be about 6 foot-candles at the level of the birds.
(iv) Temperature and humidity should be controlled during
the study. Recommended levels are 21°C and 55 percent relative
humidity. Temperature should be recorded at least weekly at the
same time of day and should be reported. For tests conducted
without temperature control, temperature minimums and maximums
should be recorded daily. Continuous temperature monitoring is
desirable. Temperature recordings should be made at a level of
2.5-4 cm above the floor of the cage. Recording of approximate
humidity levels is also desirable. Good ventilation should be
maintained. Suggested ventilation rates are 4 changes per hour
in winter and 15 changes per hour in summer.
(v) If facilities are being used for the first time, it may
be desirable to allow birds to breed in the facility prior to
testing in order to ensure that controls will have acceptable
productivity according to the requirements given in paragraphs
(c)(4)(xi) and (xii).
(3) [Reserved]
(e) Reporting. (1) The test report should include the
following information:
(i) Name of test, sponsor, test laboratory and location,
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August, 1982
principal investigator(s), and actual dates of beginning and end
of test.
(ii) Name of species tested (including scientific name), age
of birds (in months) at the beginning of the test, source of
birds, and body weights for adult birds throughout tne test.
(iii) Description of housing conditions, including type,
size, and material of pen, temperature, humidity, photoperiod and
lighting intensity, and any changes during the test.
(iv) Detailed description of the basal diet, including
source, composition, diluents (if used), and supplements (if
used). A nutrient analysis of the basal diet should be included.
(v) Detailed description of the test substance including its
chemical name(s), source, lot number, composition (identity of
major ingredients and impurities), and known physical and
chemical properties pertinent to the test (e.g., solubility,
volatility,, degradation rate, etc.).
(vi) The number of concentrations used, nominal and measured
concentrations of test substance in each level, assay method used
to determine actual concentrations, storage conditions and
stability of treated diets, number of birds per pen and number of
replicate pens per concentration and for controls.
(vii) Acclimation procedures and methods of assigning birds
to test peris, including method of randomization, and any
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August, 1982
rearrangement due to incompatibility.
(viii) Frequency, duration, and methods of observation.
( ix) Description of any signs of intoxication, including
time of onset, duration, severity (including death), and numbers
affected, including accidental deaths or injuries.
(x) Food consumption per pen and any observations of
repellancy or food palatability.
(xi) Method of marking all birds and eggs.
(xii) Details of autopsies.
(xiii) Egg and hatching data in summary and by pen per week
in sufficient detail to allow an independent statistical
analysis. Data should be presented for all of the parameters
listed in paragraph (c)(6)(i). The number of eggs set should
also be reported.
(xiv) Egg storage, incubation, and hatching temperatures,
relative humidities, and turning frequencies.
(xv) Observations of health and weights of young at 14 days
of age.
(xvi) Location of all raw data storage.
(xvii) Methods of statistical analysis and interpretation of
results .
(xviii) Anything unusual about the test, any deviation from
these procedures, and any other relevant information.
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August, 1982
(2) In addition, the following information should be
available apon request:
(i) A general description of the support facilities.
(ii) A description of the Quality Control/Quality Assurance
program, including the Average Quality Level for the program
element performing the test, procedures used, and documentations
that these levels have been achieved.
(iii) The names, qualifications, and experience of personnel
working in the program element performing the test, including the
study director, principal investigator, quality assurance
officer, as well as other personnel involved in the study.
(iv) Standard operating procedures for all phases of the
test and equipment involved in the test.
(v) Sources of all supplies and equipment involved in the
test.
(vi) Diagram of the test layout.
(vii) Originals or exact copies of all raw data generated in
performing the test.
(viii) A detailed description, with references, of all
statistical methods.
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TECHNICAL SUPPORT DOCUMENT
FOR
BOBWHITE REPRODUCTION TEST
OFFICE OF TOXIC SUBSTANCES
OFFICE OF PESTICIDES AND TOXIC SUBSTANCES
U.S. ENVIRONMENTAL PROTECTION AGENCY
WASHINGTON, D.C. 20460
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TABLE OF CONTENTS
Subject Page
I. Purpose 1
II. Scientific Aspects 1
General 1
Issues 5
Test Procedures 7
Test Substance Concentrations 7
Controls 7
Test Groups and Number of Animals 7
Duration of Test 3
Preparation for Reproduction 9
Observations and Measurements 11
Adult Birds 11
Chicks 11
Eggshell Thickness 12
Typical Observed Values 13
Egg Collection, Storage and Incubation 14
Required Analysis 15
Statistical 15
Test Substance Concentration 16
Basal Diet 17
Acceptability Criteria 17
Test Conditions 19
Test Species 19
Selection 19
Maintenance of Test Species 22
Acclimation 22
Diet 22
Facilities 23
Environmental Conditions 25
Temperature (See Section II.C.3) 25
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Subject Page
Humidity (See Section II.C.3) 25
Reporting 25
III. Economic Aspects 26
IV. References 27
11
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Office of Toxic Substances ES-13
August, 1982
TEST SUPPORT DOCUMENT FOR BOBWHITE REPRODUCTION TEST
I. Purpose
The purpose of this document is to provide the
scientific background and rationale used in the development
of Test Guideline EG-16 which uses Bobwhite to evaluate the
effect of chemical substances on reproduction. The Document
provides an account of the scientific evidence and an
explanation of the logic used in the selection of the test
methodology, procedures and conditions prescribed in the
Test Guideline. Technical issues and practical
considerations are discussed. In addition, estimates of the
cost of conducting the test are provided.
II. Scientific Aspects
A. General
The earliest investigations of the effects of chemicals
on reproduction of native birds were in the 1950s (DeWitt
1956, Genelly and Rudd 1956). Chemicals were administered
in the diet, but procedures varied. Laboratory
investigation of reproductive effects of pesticides
continued at Patuxent Wildlife Research Center, but methods
were not reported well (DeWitt et al. 1962, DeWitt et al.
1963). In 1964, a very brief protocol for reproduction
studies was developed at that center (Stickel and Heath
1965). This protocol outlined the egg parameters to be
studied. Ratcliffe (1967), in a classic paper, correlated
the decline of certain avian populations with thin eggshells
that apparently had resulted primarily from exposure to DDT
and DDE. Heath et al. (1969) presented the first clearcut
experimental data showing that DDE caused thin eggshells in
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mallards. The protocol used by Heath et al. (1969),
although not fully detailed, formed the basis for most
future avian toxicological reproduction studies. In 1968,
USDA developed a guideline for evaluating reproductive
effects of toxic chemicals to birds (US EPA 1975). This
guideline apparently was developed in conjunction with
Patuxent personnel as it bears a great similarity to methods
used at Patuxent, but was available prior to the publication
of Heath et al. (1969). The Agency (US EPA 1975) developed
composite protocols for reproduction tests from the limited
published information and unpublished information,
especially the USDA protocol which was presented as an
exhibit. No complete, suitable protocol for bobwhite was
available from the published literature at that time (US EPA
1975), although Heath et al. (1972b) and Heath and Spann
(1973) had published the results of bobwhite reproduction
studies. The Agency's pesticide guidelines were revised (US
EPA 1978a) but the basic method of the earlier guidelines
for reproduction tests was retained. There have been very
few bobwhite reproduction tests published even in recent
years, although a number of tests have been conducted,
classified as confidential, and submitted to the Agency to
support pesticide registration. ASTM (1979a) has prepared
a draft avian reproduction method. Although this ASTM
protocol is similar to the Office of Pesticides Programs
protocol (US EPA 1978a), it is designed for a variety of
species. Thus, the history of avian reproduction test
methodology is basically a history of Patuxent and EPA
methods. This guideline continues the trend because these
methods appear to be the most appropriate for developing
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data needed to make regulatory decisions and because no
other methods have become accepted widely.
Even though the basic method is similar for most
investigators, there have been a few points of difference or
controversy. Yet there are very few data to address these
differences and little ongoing research that might resolve
conflicts. To promote uniformity and comparability of
tests, some recommendations and requirements are
standardized in this test guideline. Where such
recommendations and requirements are controversial and are
not sufficiently addressed by published data, it is hoped
that research will be stimulated to resolve questions. If
feasible, conditions and procedures that approximate natural
conditions have been selected in preference to other
options.
The use of avian reproduction tests in the assessment of
chemical impacts on the environment is based on several
factors. First, birds are an obvious and important
component of the environment. Congress has indicated
repeatedly that birds are worthy of protection by passing
such laws as the Lacey Act of 1900, Migratory Bird Treaty
Act of 1913, Migratory Bird Conservation Act of 1929,
Pittman-Robertson Act of 1937, Fish and Wildlife Act of
1956, Endangered Species Act of 1973, and others. The
United States also has entered into treaties with Great
Britain and Canada (1916), Mexico (1937), Japan (1974), and
Russia (1976) for the protection of migratory birds. The
people of the United States also have indicated a desire to
protect birds through their support of the Audubon Society,
Nature Conservancy, and other environmental groups.
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August, 1982
Sportsmen's organizations support protection of birds,
although their interests often focus heavily on game birds.
Second, birds have a definite economic importance.
Federal and State Agencies spend large sums for the
preservation and propagation of birds. Hunters and
birdwatchers also spend substantial sums in pursuit of their
pastimes. Less measurable, but of definite importance, is
the substantial role of birds in insect control.
Birds have an important ecological role. Insectivorous
birds consume huge quantities of insects and other
invertebrates, many of which are considered pests. Small
mammals and other vertebrates or plants are consumed by
various birds, sometimes to the extent that birds have an
important effect on populations. In turn, birds are
consumed by birds of prey, mammals, and other vertebrates.
Excretory products of birds provide nutrients for plankton
and other microorganisms that in turn are food for larger
organisms. Birds are important in pollination of some plants
and in dispersal of others. Because of their mobility, the
effects of birds are not restricted to specific locations.
Finally, birds are among the more sensitive terrestrial
vertebrates. Because of their high metabolic rate, high
body temperature, and the demands of flight, they require
more energy relative to their size than most other
animals. The energy requirements lead to greater food
intake and thus to greater toxicant intake when a toxicant
is in or on their food. There are abundant data showing
that some birds, particularly raptors, pelicans, and
waterfowl, are very sensitive in their reproductive
responses to toxic chemicals (e.g., Ratcliffe 1967, Anderson
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August, 1982
and Hickey 1972) and that some species have suffered from
drastic population reductions apparently as a result. Avian
reproduction is unique, and no surrogate is adequate to
predict effects on eggshells, the primary mode of impairment
for many chemicals such as DDE (Heath et al. 1969).
Avian reproduction tests are extremely valuable in
assessing the potential population effects of exposure of
birds to toxic chemicals. The route of intake simulates
natural exposure to chemicals on or in the food. Most
physiological effects can be assessed under laboratory
conditions, although many behavioral effects such as nest
desertion are difficult to study in the laboratory. A
positive finding of impairment in the laboratory is highly
predictive (qualitatively) in the field when exposure is the
same. However, negative findings in the laboratory may not
preclude adverse effects under field conditions. Thus, some
extrapolation may be made from laboratory to field, but
quantitative extrapolation is risky.
1. Issues
The avian reproduction test guideline and support
document contain some controversial points. Data are
insufficient or absent to support either side of most
points. For other points, there may be data supporting each
side. A number of controversial points have been selected
as potential research projects. Issues are merely
identified below and are discussed in the appropriate
sections of this document.
o Is productivity from tests run out of normal
season sufficient to evaluate the potential for
reproductive impairment?
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August, 1982
o Should methods involving natural incubation of
eggs be incorporated into tests?
o Should the current use of first year birds only be
continued or should proven breeders be used?
Would either age produce results suitable for
comparison?
o What carriers, if any, should be used or allowed
for incorporation of the test substance into the
diet?
o Are commercial foods adequate, or is there too
much variation and/or contamination of commercial
foods?
o How often should treated diets be mixed? Is there
an advantage in mixing diets with decreasing
concentrations to simulate natural degradation of
test substances?
o What is the optimum number of birds to be tested
in order to attain statistically valid results and
still be cost-effective? Can tests using pairs
only of birds in pens be successfully conducted by
a variety of testing facilities?
o Can more useful results be obtained by testing
enough dose levels to use linear regression
analys is?
o Should outdoor tests be allowed?
o Should medication be allowed as is needed to treat
individual sick birds?
o Are the typical values for productivity and other
egg parameters realistic for a wide diversity of
testing facilities, or are they really only valid
for a few?
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B. Test Procedures
1. Test Substance Concentrations
Test substance concentrations in the diet will be
specified in the test rule. These concentrations will be
based upon the properties of the test substance, the lethal
and no-effect levels, if known, or the amount of test
substance known or likely to be found in the environment.
Three concentration levels are specified because, for many
test substances, three levels will allow for a dose-response
regression analysis from which a no- effect level can be
calculated. (See section 2.1.8 on statistics for further
discuss ion).
2. Controls
Concurrent controls are required for every test to
assure that any observed effects are a result of ingestion
of the test substance and not of other factors. Such other
factors may include environmental factors such as
temperature or lighting, vapors, sensitive or stressed test
birds, etc,. If a diluent is used in mixing the diet, this
diluent also is used in the untreated diets in the same
concentration as it occurs in treated diets. In effect,
this results in a diluent, but no completely negative,
control. Diluent choices are based upon their lack of
toxicity (e.g., water, completely evaporated acetone) and it
is not considered necessary, therefore, to have an
additional negative control when a diluent control is used.
3., Test groups and numbers of animals
A minimum of 12 replicate pens with one cock and two
hens or 20 replicate pens with one cock and one hen is
required for each test concentration. The number of
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August, 1982
replicates is needed to achieve a test with statistical
significance. There is enough variation in productivity of
bobwhite hens that fewer replicates would be insensitive to
all but the most severe effects.
The most recommended arrangement is for use of group
pens containing two females each. This is based upon the
rather agressive behavior of courting males that could
result in severe stress to the female, if only one hen were
present. In reproduction tests different from this
guideline and other published protocols, Kendall et al.
(1973) used pairs of bobwhite. Currently, K. Stromberg
(personal communication) has been working on reproduction
tests with pairs of bobwhite at Patuxent Wildlife Center
(USDI). Data from Stromberg's work are still being
analyzed, but it is expected that methods for keeping
bobwhite pairs successfully will result from that
research. The alternative arrangement of 20 bobwhite pairs
per level is suggested to accommodate those test facilities
that believe they will have success or that wish to
contribute to the development of methods. A warning is
included in the guideline so that relatively inexperienced
investigators should not expect unqualified success. The
additional pens in pair testing will strengthen notably the
statistical analysis of the testo This alternative is
included because it is statistically stronger than the more
familiar method of group testing.
4. Duration of Test
The avian reproduction test lasts approximately 22
weeks. The initial part of the test is an exposure phase
where birds are receiving treated diet. Exposure to treated
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August, 1982
diets begins with this phase and allows for the test
substance to act upon the reproductive mechanisms of the
body, and also for birds to accumulate residues of
lipophilic and other test substances. The development of
reproductive capacity actually begins months before egg
laying starts (Kirkpatrick 1959). Therefore, exposure
to the test chemical should be well in advance of egg
laying, if the test is to investigate reproductive effects
in general.
The second phase, following lengthening of the
photoperiod, directly brings the birds into readiness for
egg laying. The duration of this phase is dependent upon
the response of the test birds.
The third phase is the egg-laying portion of the test.
This is to be a minimum of eight weeks. The duration of
this phase is based upon two main factors. First, it is
important to determine if egg laying is within normal levels
or if it declines, or otherwise varies, over a period of
time. This yields information on speed of action and has a
role in using the test for assessment purposes. Second, the
eight week period is needed to provide sufficient data for a
strong statistical analysis.
A withdrawal period is optional when impairment has been
detected. This period may provide data on recovery of
reproductive capacity that could be useful for assessment
purposes.
5. Preparation for reproduction (photoperiod)
Because photoperiod is critical to reproduction, it
should be controlled in indoor tests. Under natural
conditions, photoperiod is lengthening gradually just prior
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August, 1982
to the reproductive season. The earlier bobwhite
reproduction tests were conducted outdoors (Stickel and
Heath 1965, Heath et al. 1972b, Heath and Spann 1973), and
therefore, used natural photoperiod. More recently,
bobwhite tests involving reproduction have been conducted
indoors (Fergin and Schafer 1977, Kendall et al. 1978) and
egg laying has been induced by increasing photoperiod to 16
or 17 hours of light per day. The ASTM (1979a) and US EPA
(1978a) protocols both recommended seven hours of light in
the first phase and 16-17 hours light in the second phase.
Without giving supporting data, both of these protocols
stated that the regime is for maximum egg production, and
both specify at least an option of gradually increasing the
length after egg laying has started. The photoperiod in
this guideline is based on the above data and a lack of any
suggested alternatives.
The dark period in the photoperiod should not be
interrupted, even briefly, except as absolutely necessary.
Kirkpatrick (1955) found that even as little as 15 minutes
interruption on a short day/long night regime caused an
increase in gonadal development, and all birds exposed to a
60 minute interruption became fully active sexually. It is
highly probable, on the basis of Kirkpa trick's (1955, 1959)
data, that it is the length of the dark period rather than
the light period that controls reproductive preparation.
Kirkpatrick (1955) also tested light intensity as a
factor in bobwhite reproduction. Responses were very
similar at intensities of 0.1 to 100 foot candles on a 17
hour light photo- period. With shorter days, the 0.1 foot
candle intensity did not achieve the same results as 1,10,
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August, 1982
and 100 foot candles. These data support the approximate 6
foot candle intensity suggested by ASTM (1979a) and US EPA
(1978a).
6. Observations and Measurements
a. Adult Birds
Observations of food consumption, body weights, and
signs of toxicity are required. Body weights are required
three times during the test. More frequent body weights
might be informative, but the stress of handling may offset
the collection of data, particularly for laying hens. Food
consumption is to be estimated at frequent intervals. This
will provide data both on test substance ingestion and on
energy intake for test birds. The latter data will be
particularly helpful in the absence of frequent weighings.
Clinical signs of toxicity contribute substantially to
the analysis of the data, in addition to providing
information on the mode and speed of action. If dose levels
are finely tuned, there should be minimal observed acute
toxicity. However, without adequate preliminary data, one
or more of the test levels may cause lethal or notable
sublethal effects. Such effects may affect the results of
the test without acting on the specific reproductive
parameters being investigated. For example, a severely
stressed bird may not be able to mobilize internal resources
to produce eggs. Signs of toxicity will give valuable
information in evaluating the results of the test.
b. Ch i cks
The incubation period for bobwhite eggs has been
reported to be 559± 7 hours (23.3 days) (Wetherbee and
Wetherbee, as cited in ILAR 1977). Some variation may occur
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August, 1982
as a result of temperature or humidity conditions (ILAR
1977). Bobwhite eggs have been incubated for 24 days
(Fergin and Schafer 1977) or 25 days (Heath and Spann
1973). US EPA (1978a) stated that eggs should be removed
from hatchers on day 24. The meager data available suggest
that 24 days is sufficient and this incubation length has
been selected for this guideline.
Because the pen is the basic unit for statistical
analysis, it is necessary that the eggs and hatchlings be
identified as to pen of origin. This can be done either by
housing all chicks from one pen together, or by individually
marking each egg and bird.
Environmental conditions for young chicks are based upon
historical precedents (Heath et al. 1972, Hill et al. 1975)
and other current methods for maintaining young chicks (ASTM
1979b, US EPA 1978b). Parameters given in these references
include temperature of 35°C with a lower temperature outside
the cage to provide a gradient, galvanized brooder cages
maintained indoors, and a commercial game bird starter
ration. These conditions have been included in this
guideline.
Observations on chicks are necessary to determine if and
when toxicity might be expressed in the offspring. Although
most test substances exert their action prior to hatching,
some may affect growth, development, or survival of chicks.
c . Eggshell Thickness
The classic example of reproductive impairment in birds
is the eggshell thinning effect of DDE, a metabolite or
degradate of DDT (Tucker and Leitzke 1979). It is important
to measure eggshell thickness because as little as 11%
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thinning may have an effect on avian populations (Tucker and
Leitzke 1979). Techniques given in this guideline are
widely used (Heath and Spann 1973, ASTM 1979a, US EPA
1978a), although many papers do not give a full description
of the process. It is particularly important for several
measurements to be made around the girth of bobwhite eggs,
because these eggs may have small calcium deposits that
could affect results if only one or two measurements are
taken.
d . Typical observed values
The typical observed values presented in this guideline
have been taken from the Agency's pesticide guidelines (US
EPA 1975, 1978a). A number of avian reproduction tests have
been submitted to the Agency for the purpose of pesticide
registration and have been classified as confidential. Most
of the tests that have been conducted during the normal
reproductive season have achieved for control birds the
typical values as presented in the test guideline. Heath
and Spann (1973) and Fergin and Schafer (1977) achieved
these values in control birds insofar as could be determined
from their data. Heath et al. (1972b) did not quite reach
the egg production values in an eight week season for a test
conducted outdoors with three hens and 2 cocks per pen.
They did achieve all other normal values except that for
cracked eggs. Ten week indoor tests according to this
guideline would likely have egg production values that
correlated with these normal values.
The values presented in the guideline have been
identified as an issue. It is known that some testing
facilities routinely
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August, 1982
meet these norms. However, it is not known if the values
are realistic when many testing facilities are considered.
7. Egg Collection, Storage, and Incubation
Egg storage prior to setting for incubation has been
reported for bobwhite in only a few papers and has ranged
from 16°C and 55% humidity (Heath et al. 1972b) to 19°C and
70° humidity (Kendall et al. 1978). Stromberg (personal
communication) stated that he stored eggs at 16° and as high
a humidity as could be obtained from evaporating water; this
was up to 80%, but at least 50% relative humidity. He also
stated that turning the eggs daily was important to keep
part of the shell and egg from excessive drying. US EPA
(1978a) recommends 16°C and 65% relative humidity. This
guideline specifies 16°C, but only a range of relative
humidity between 55% and 80%, since many testing facilities
do not have the means to control storage humidity and since
this range has produced good viability.
No papers were found that reported incubation
temperatures and humidities for bobwhite. Neither ASTM
(1979a) nor US EPA (1978a) specify incubation temperatures
although the latter does recommend 39°C and 70% relative
humidity during the hatching phase from day 21 to day 24.
Stromberg (personal communication) stated that 99.75°F
(37.6°C) and 50-70% relative humidity are standard
commercial poultry incubation temperatures and that all
testing facilities with which he is acquainted use
commercial poultry incubators. Mallards have been incubated
at temperatures of 37.4-37.5°C and relative humidities of
62-80% (Heinz 1976a, Heinz 1976b, Greenwood 1975, Holmes et
al. 1978). These same parameters have been specified in
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this bobwhite test guideline.
Candling procedures are a standard practice for
determining eggshell cracks, fertility, and embryo
survival. These procedures have been used in most bobwhite
tests, but timing has been inadequately reported. US EPA
(1978a) specifies, for bobwhite, candling on day 0 for
cracks, on day 11 for fertility and on day 18 for embryo
survival. This timing has been adopted in this guideline
because it will permit comparisons of test data with
existing studies in Agency files. At least one major avian
testing laboratory protocol uses the same timing.
US EPA (1978a) recommended moving eggs to a separate
hatcher or incubator for hatching, but gave no reasons.
None of the available published papers mentioned this
procedure. However, Stromberg (personal communication)
stated that this procedure is highly recommended as a means
to minimize disease vectors. This procedure has been
recommended in the guideline.
This test guideline does not consider the effects on
incubation behavior, nest desertion, and care of young.
Techniques for investigating the effects of chemicals on
these behaviors are in their infancy and, at present, are
prohibitively expensive. Agency research has been proposed
to investigate methods of incorporating natural incubation
into future reproduction guidelines at a reasonable cost.
8,. Required Analyses
a. Statistical
The statistical analysis of avian reproduction studies
typically has been analysis of variance (Heath and Spann
1973, Heinz 1976a, US EPA 1978a, ASTM 1979a) using the
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August, 1982
parameters set forth in this guideline. Angular
transformations have been used to a lesser extent (Heath et
al. 1972b). Regression analysis is a powerful tool that may
be used if there are data at sufficient dose levels to
obtain a dose-response curve. Regression analysis has been
recommended, as an option by ASTM (1979a), and methods
directed toward regression analysis are being developed
(Stromberg, personal communication). It is the opinion of
the author of this test guideline that regression analysis
is a more useful tool than currently typical methods because
it yields a dose response curve and this curve can be used
for extrapolation. Analysis of variance provides only
significance at a particular level and does not lend itself
to extrapolation on a reliable basis. At the present time,
methods and background work have not been developed for a
test oriented primarily towards a dose-response curve. More
dose levels would be needed, possibly with fewer animals per
level. This subject may be included in the Agency's
research projects. In the meantime, this guideline uses
methods appropriate to analysis of variance.
b. Test Substance Concentrations
Samples of treated diets will be analyzed to determine
the actual levels to be used in the test. Analysis will
help to detect mathematical errors in calculating
concentrations, technicians' errors in mixing diets, and
manufacturers' errors in determining the amount of active
ingredient in a test substance (Heinz, personal
communication). All test substance concentrations will be
analyzed so that, even with only three test concentrations,
a dose-response curve (if obtained) would be based on
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measured concentrations, rather than on normal levels. A
second analysis about midway through the test will
corroborate initial levels.
c. Basal Diet
Most testing facilities use diets prepared by commercial
feed companies. Some facilities may have a commercial
company prepare a diet to order. Normally, such diets are
supplied with a quantitative list of ingredients, and such a
list should be supplied with the test report. If there are
supplements added to the diet, a list of all supplemental
ingredients also should be submitted. Analysis of
ingredients in the basal diet is important because there are
a number of potential test substances, such as certain
metals, that may interact with components of the diet and
possibly affect the results of a test. A nutrient analysis
will allow for a better evaluation of such results. In
addition, it is possible that dietary deficiencies or
imbalance of ratios of nutrients also could affect the
results. Even though commercial companies normally supply a
nutritionally adequate diet, it is important to know the
components because no rigid requirements exist for the type
and constitution of the diet used.
9. Acceptability Criteria
Test acceptability is dependent upon following mandatory
requirements and having acceptable control productivity and
survival. When control values do not reach the typical
observed values, as given in section (h)(ll) of the test
guideline, it is very likely that there are problems with
some aspect of the test. Since reproductive parameters of
treated birds are compared with controls, it is essential
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that values for control birds are normal as determined from
similar reproduction studies.
Rather than requiring every parameter to achieve normal
values, it was decided that acceptability of a test should
be based on the final parameter of 14-day old survivors per
control hen as one criterion. For bobwhite, the requirement
of 12 survivors per hen over a ten week period was selected
because data in the Agency's pesticide files show that
acceptable tests have reached and usually surpassed this
level. Heath and Spann (1973) passed these levels in test
birds over eight weeks, and had comparable control
productivity of 7.5 survivors per hen in four weeks. Heath
et al. (1972b) reported 11.5 control survivors per hen in
eight weeks.
Eggshell thickness of control birds is also a criterion
of test acceptability because thin eggshells among control
birds are usually a sign of inadequate diet, which in turn
could affect other aspects of the test. Heath et al.
(1972b) reported bobwhite shell thickness of control groups
as .207 mm. Heath and Spann (1973) reported control values
as ranging from .22 to .25 mm. McGinn is et al. (1976)
reported average bobwhite values of .226 and .236 mm
( ranges-. 196-. 267 and .185-.274, respectively) in a non-test
investigation. It might be unrealistic to expect all eggs
to meet the designated thickness of .19 mm, but if average
thickness is less, then a problem probably exists.
A well conducted test with adult birds should not result
in any but an occasional mortality in control groups, even
though the test is relatively long. This is especially true
since this guideline provides for rearrangement of
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incompatible birds during the acclimation period.
Therefore, control mortality in excess of 10% indicates
problems with some aspect of the test.
C. Test Conditions
1. Test Species
a. Selection
The bobwhite, Colinus virginianus (L), is the species to
be tested. The choice of bobwhite is based on several
factors. Although no single species would satisfy all
criteria for species selection, the bobwhite has a number of
favorable attributes. The bobwhite has a widespread
distribution throughout much of the United States, and it is
an important part of the avifauna of the southern U.S. It
occurs in a variety of terrestrial habitats and is likely to
be exposed to toxic substances that occur in such
habitats. As an ecologically relevant species, the bobwhite
represents its own species as well as serving as an
indicator for other species.
The bobwhite has not been used extensively in
reproduction tests that have been published. In general,
waterfowl are considered to be more susceptible to
reproductive impairment than gallinaceous birds. However,
DeWitt (1962) found bobwhite to be 50 times more sensitive
than mallards to reproductive impairment from toxaphene.
Similarly, gallinaceous birds have exhibited reproductive
impairment from PCBs, but mallards have not (Stickel
1975). Other confidential data in the Agency's pesticide
files have confirmed that bobwhite are sensitive to some
chemicals although not as frequently as are waterfowl. When
reproductive tests are conducted on both gallinaceous birds
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and waterfowl, this combination of species will often
indicate impairment that might not be found if only one
species were tested.
The bobwhite is also amenable to testing in the
laboratory. Bobwhite can be bred in captivity and are
readily available from commercial sources so that testing of
this species will not deplete wild stocks. There is
sufficient information on the nutritional, habitat, and
behavioral characteristics of natural populations of
bobwhite in order to meet the basic nutritional and physical
requirements of the species in the laboratory.
A major advantage of reproduction testing with bobwhite
instead of other galliforras is that bobwhite is typically
one of two species used in short-term toxicity tests with
birds. There is an advantage in using the same species in
reproductive tests that also can be used in short-term test
or field tests. Bobwhite have been used for short-term
laboratory test and actual or simulated field tests from the
beginnings of ecological effects testing CDeWitt et al.
1962) to the present (US EPA
1980b). Thus, the choice of bobwhite as a test species
facilitates comparisons of the results from different kinds
of tests.
In addition, bobwhite are generally considered to have a
positive economic value. Although the Agency is charged
with the protection of all species in the environment, the
choice of an economically valuable species for testing is
appropriate to the cost-benefit or risk-benefit analyses
upon which Agency decisions frequently are based.
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August, 1982
If a test is to simulate toxicity to naturally occurring
populations of bobwhite, then it is important to use birds
that are phenotypically indistinguishable from wild birds.
Since many chemicals act upon specific enzymes and enzymes
are based on the genetic code, the use of birds
genotypically similar to wild birds would be desirable.
However, the determination of phenotype is a simple
observational process, whereas genotypic determination is
impractical, if possible at all. In addition, wild birds
have a degree of heterogeneity that would not be typical of
any given genotype.
The necessity for using healthy birds is obvious since
the test is designed to determine toxicity to typical
populations. It is admitted that not all birds occurring in
natural populations are healthy, but the majority of
survivors in natural environments are healthy. The health
of birds is also important in reducing the number of
variables that limit comparisons between tests. There are
several checks in this test guideline that help to insure
that healthy birds are used. The use of previously untested
birds not selected for resistance and being from disease
free flocks provides a basically healthy stock. Visual
observations select out abnormal or unhealthy birds from
that stock. A final check on health is based on the birds
ability to survive two weeks immediately preceding
exposure. The 3% maximum mortality during this period
allows for an occasional death that may occur during
acclimation at the time when unfamiliar birds in a pen are
becoming adapted to each other.
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August, 1982
The choice of first year birds was made in this
guideline primarily because it facilitates comparisons with
existing data and data that may be generated according to
the US EPA (1978a) protocol for bobwhite. It also ensures
that birds will be the same age; whereas if no age were
specified, birds in the same test could be various ages.
Use of first year birds also reduces the cost of birds and
ensures that test birds will not have been used in previous
reproduction tests.
2. Maintenance of Test Species
a. Acclimation
An acclimation period is necessary for birds to become
familiar with the test environment. Ideally, birds will be
maintained in test cages for several months, but this is
impractical for testing facilities that purchase adult
birds. It is also sometimes necessary to alter the
composition of birds in a pen because of excessive
aggression or other incompatibilities. The acclimation
period allows time for rearranging incompatible birds.
b. Diet
There are few data on the detailed nutritional
requirements for bobwhite. This subject is being
investigated under a current contract and is a proposed
research need. At the present time, a commercial game bird
breeder ration is recommended. All known testing facilities
in this country use a commercial ration or a similar but
specially prepared ration made by a commercial company.
There are no known data to support an alternative diet for
bobwhite. Changes may be made in the future when additional
data become available. The recommendation of a commercial
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August, 1982
game bird starter ration is made for the same reasons.
Samples of treated diets will be analyzed to determine
the actual test concentrations used in the test. See
section 2.2.8 for details
3. Facilities
Very few bobwhite reproduction tests have been reported
in the literature, although a number have been submitted to
the Agency and classified as confidential. As a result, it
is difficult to define optimal conditions and most
conditions have been recommended, rather than required.
Heath et al. (1972b) and Heath and Spann (1973) used outdoor
pens with wire mesh floors for their bobwhite reproduction
studies. Heath and Spann (1973) specified the size of cages
as 3 X 6 feet; they housed 6 hens and 3 cocks in each pen.
Heath et al. (1972b) may or may not have used the same pens;
they housed 3 hens and 2 cocks in each pen. Kendall et al.
(1978) used pairs of bobwhite housed in quail battery
breeding pens (size unspecified) indoors, apparently at
24°C. They also used .6 x 1.5m (2x5 ft) pens, housing 5
hens and 5 cocks, in a poultry house. Illumination
intensity, temperature, and humidity were not reported by
Heath et al. (1972) and Heath and Spann (1973). Presumably,
these were not controlled since pens were outdoors.
US EPA (1978a, 1980b) and ASTM (1979a) have both
developed protocols for avian reproduction tests.
Rationales for selecting particular conditions were not
spelled out in either protocol. Reasons were apparently
based on experience with reproduction tests that have been
submitted to the Agency for pesticide registration and that
have been classified as confidential. These three protocols
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August, 1982
have specified size only as being "adequate". All three
protocols recommend 21°C, 55% relative humidity, and
adequate ventilation. The same conditions have been
recommended in this guideline, since these protocols are the
most widely used and data developed from this guideline will
be comparable with data gathered using all three protocols.
Galvanized metal is the recommended construction
material. This material has been widely used in toxicity
tests (Heath et al. 1972a, Hill et al. 1975) and has been
recommended in ASTM (1979a) and EPA (1978a) protocols for
reproduction tests. The ASTM (1979a) protocol suggests
stainless steel or perfluorocarbon plastics (e.g., Teflon)
as alternatives. As a relatively non-adsorbent material,
perf luorocarbon plastics may be used to coat wood or other
materials that might be contaminated by chemicals if used
uncoated. Because of the tendency of wood to sorb
chemicals, uncoated wood may not be used where it is likely
to become contaminated and come in contact with birds in
subsequent tests.
Pens should be cleaned and sanitized between tests.
Brushing and/or steam cleaning appear to be the most
appropriate since they do not involve the use of chemicals
that could affect subsequent tests. Detergents and bleach
have been used by Denver Wildlife Research Center (Tucker,
personal communication) and Patuxent Wildlife Research
Center (Heinz, personal communication). The use of chemical
disinfectants, such as quaternary ammonium compounds, should
be avoided because of possibility that these compounds can
leave toxic residues. However, the widely used cold
sterilization method with ethylene oxide is acceptable, if
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August, 1982
needed for disease control. Pens should not be cleaned
during a test in order to minimize disturbance to the test
birds.
4. Environmental Conditions
a. Temperature (See Section II.C.3)
b. Humidity (See Section II.C.3)
D. Reporting
The information that is required to be reported in
section II.B.S.b is essential to a proper evaluation of the
test results. These required items are needed (1) to
establish that the test was conducted according to
specifications, (2) to evaluate those conditions and
procedures that could affect the results of the test, and
(3) to supply the Agency with sufficient information to
conduct and independent analysis of statistics and
conclusions. The location of the raw data storage will
allow the Agency to find additional information that may
have been left out of the report or that may be needed for
enforcement purposes. The location is necessary because
some chemical companies request the testing facility to keep
these data, while other companies keep their own. This
information is needed in a detailed manner because this
avian reproduction guideline contains few rigid
requirements. Even when minimums or maximums are specified,
it is important to know how much the test may have exceeded
specifications, such as; if test birds were observed more
frequently than required, if the number of test
concentrations exceeded the three levels required, etc.
The information required in section II.B.S.b on Test
Substance Concentration to be available, but not included in
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August, 1982
the test report, may be needed if there are serious concerns
about the results or validity of the test. This information
will not normally be needed and therefore is not required
routinely.
III. Economic Aspects
Three laboratories were surveyed to estimate costs for
performing the test outlined in this guideline. The
individual laboratories gave prices of $17,000, $10,780, and
$38,000. The "best estimate" based upon the survey was
$21,927. A cost estimate also was made by separating the
protocol into components and estimating the cost of each
component, including direct labor cost, overhead cost, other
direct cost, general and administrative cost, and fee. The
best estimated final cost, based upon this calculation
method, was $20,562, with an estimated range of $10,281 to
$30,844 based on _+_ 50% of the best estimate. The calculated
estimate is similar to the best estimate based on the
survey. Marked differences in prices obtained from the
different laboratories may have resulted from a number of
factors, such as nature of the chemical, overhead rates,
outside consultants, automation, marketing strategies, and
other factors as outlined in a cost analysis report by
Enviro Control (1980). The cost estimates were made
assuming that the requirements of the Good Laboratory
Practice Standards, as specified in section (d) of the
Bobwhite reproduction guideline, are being satisfied.
In a cost analysis of subpart E pesticides guidelines,
laboratories were surveyed in 1978 and in 1980 to determine
the cost of testing (US EPA 1980a). The cited costs did not
differentiate between species, however, the unit cost for an
avian reproduction test was $24,000 in 1978 and $28,000 in
1980.
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IV. References
Anderson DW, Hickey JJ. 1972. Eggshell changes in
certain North American birds, pp. 514-540 in
Proceedings of the XV International Ornithological
Congress. Voous KH, ed. E.J. Brill, Leiden, 745 op.
1972.
ASTM 1979a. American Society for Testing and
Materials. Standard practice for conducting
reproduction studies with birds. Draft No. 1. 20 pp.
ASTM 1979b. American Society for Testing and
Materials. Standard practice for conducting LC
with birds. Draft No. 1. 13 pp.
50
tes ts
DeWitt JB. 1956. Chronic toxicity to quail and
pheasants of some chlorinated insecticides. J. Ag.
Food Chem. 4(10): 863-866.
DeWitt JB, Crabtree DG, Finley RB, George JL. 1962.
Effects on wildlife. pp. 4-10 (+Tables) in USDI,
Effects of Pesticides on Fish and Wildlife: A Review
of Investigations during 1960. Bureau Sport Fish and
Wildl. Circ. No. 143. 52 pp.
DeWitt JB, Stickel WH, Springer PF. 1963. Wildlife
studies, Patuxent Wildlife Research Center, pp. 74-96
in USDI, Pesticide - Wildlife studies: A Review of
Fish and Wildlife Service Investigations during 1961
and 1962. Fish and Wildlife Serv. Circ. No. 167. 109
pp.
Enviro Control, Inc. 1980. Cost analysis methodology
and protocol estimates: ecotoxicity standards.
Rockville, MD: Enviro Control, Inc., Borriston
Laboratories, Inc.
Fergin T J, Schafer EC. 1977. Toxicity of dieldrin to
bobwhite quail in relation to sex and reproductive
status. Arch. Envir. Contain. Toxic. 6:213-219.
Genelly RE, Rudd RL. 1956. Effects of DDT, Toxaphene,
and dieldrin on pheasant reproduction. Auk 73: 529-
539.
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August, 1982
Greenwood RJ. 1975. Reproduction and development of
four mallard lines. Prairie Natur. 7(1): 9-16.
Heath RG, Spann JW. 1973. Reproduction and related
residues in birds fed Mirex. pp. 421-435 in Pesticides
and the Environment: A continuing controversy.
Symposia Specialists, North Miami. 1973.
Heath RG, Spann JW, Kreitzer JF, Vance C. 1972b.
Effects of polychlorinated biphenyls on birds, pp. 475-
485 in Proceedings of the XV International
Ornithological Congress. Voous KH, ed. E.J. Brill,
Leiden, 745 pp. 1972.
Heinz GH. 1976a. Me thy liner car y: second year feeding
effects on mallard reproduction and duckling
behavior. J. Wildl. Manage. 40(1): 82-90.
Heinz GH. 1976b. Behavior of mallard ducklings from
parents fed 3 ppm DDE. Bull. Env. Contam. Toxic.
16(6) : 640-645.
Hill EF, Heath RG, Spann JW, Williams LD. 1975. Lethal
dietary toxicities of environmental pollutants to
birds. U.S. Fish Wildl. Serv., Spec. Sci. Rept. Wildl.
No. 191. 61 pp.
Holmes WN, Cavanaugh KP, Cronshaw J. 1978. The effects
of ingested petroleum on oviposition and some aspects
of reproduction in experimental colonies of mallard
ducks (Anas platyrhynchos). J. Reprod. Fertil.
54(2) :3l3^T48.
ILAR. 1977. Institute of Laboratory Animal
Resources. Laboratory Animal Management: Wild Birds.
Nat. Acad. Sci., Washington. 116 pp.
Kendall RJ, Noblet R, Senn LH, Holman JR. 1978.
Toxicological studies with Mirex in bobwhite quail.
Poult. Sci. 57: 1539-1545.
Kirkpatrick CM. 1955. Factors in photoperiodism of
bobwhite quail. Physiol. Zool. 28:255-264.
Kirkpatrick CM. 1959. Interrupted dark period: tests
for refractoriness in bobwhite quail hens. pp. 751-758
in Withrow RB (ed). Photoperiodism. Am. Assoc. Advan.
Sci. Publ. No. 55, Washington, D.C.
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McGinnis CH, Jr, Wallace LD, Burkhart DM. 1976.
Bobwhite quail eggs: some measurements and a method
for estimating the weight of egg contents. Bull.
Environ. Contain. & Toxicol. 15( 4 ): 497-503.
Ratcliffe DA. 1967. Decrease in eggshell weight in
certain birds of prey. Nature 215: 208-210.
Stickel LF, Heath RG . 1965. Wildlife s tudies-Patuxent
Wildlife Research Center, pp. 3-30 in USDI. Effects of
pesticides on fish and wildlife. Fish and Wildl. Serv.
Circ. No. 226. 77 pp.
Stickel WH. 1975. Some effects of pollutants in
terrestrial ecosystems, pp. 25-74 in Mclntyre AD,
Mills CF. ( eds ) . Ecological Toxicology Research.
Plenum Publishing Co. , New York.
Tucker RK, Leitzke JS. 1979. Comparative toxicology of
insecticides for vertebrate wildlife and fish.
Pharmac. Ther. 6: 167-220.
U.S. Environmental Protection Agency. 1975. Protocol
for determining lethal dietary concentration of
chemical to birds. Federal Register, 40 CFR 162.82
(appendix): 26915. June 25, 1975.
U.S. Environmental Protection Agency. 1978a. Avian
reproduction. Federal Register, 40 CFR 163.71-4:
29729-29730. July 10, 1978.
U.S. Environmental Protection Agency. 1978b. Avian
dietary LC50. Federal
29728. July 10, 1978.
dietary LC50. Federal Register, 40 CFR 163.71-2:29727
l
U.S. Environmental Protection Agency. 1979. Toxic
substances control. Discussion of premanuf acture
testing policy and technical issues; request for
comment. Federal Register 44:16240-16292. March 16,
1979.
U.S. Environmental Protection Agency. 1980a. Cost
analysis: Guidelines for registering pesticides in the
United States, Subpart E. Draft. May 1980.
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U.S. Environmental Protection Agency. 1980b.
Guidelines for registering pesticides in the United
States. Subpart E. Hazard Evaluation: Wildlife and
Aquatic Organisms. Draft. November 3, 1980.
Personal Communications:
Gary Heinz (11/16/79)
Patuxent Wildlife Research Center
Laurel, MD.
Richard K. Tucker (11/8/79)
EPA, Office of Toxic Substances
Washington, D.C.
K.L. Stromberg (2/4/80)
Patuxent Wildlife Research Center
Laurel, MD.
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MALLARD REPRODUCTION TEST
OFFICE OF TOXIC SUBSTANCES
OFFICE OF PESTICIDES AND TOXIC SUBSTANCES
U.S. ENVIRONMENTAL PROTECTION AGENCY
WASHINGTON, D.C. 20460
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Office of Toxic Substances EE-17
Guideline for Testing Chemicals August, 1982
MALLARD REPRODUCTION TEST
(a) Purpose. This guideline is designed to develop data on
the reproductive effects on the mallard of chemical substances
and mixtures subject to chronic environmental effects test
regulations under the Toxic Substances Control Act (TSCA) (Pub.L.
94-469, 90 Stat. 2003 15 U.S.C. 2601 et. seq.). The Agency will
use these and other data to assess the reproductive effects on
birds that these chemicals may present in the environment.
(b) Def initions. (1) The definitions in section 3 of the
Toxic Substances Control Act (TSCA) and Part 792—Good Laboratory
Practice Standards apply here. In addition, the following
definitions apply generally to this guideline:
(i) "Acclimation" Physiological and behavioral adaptation to
environmental conditions (e.g., housing and diet) associated with
the test procedure.
(ii) "Test substance" The specific form of a chemical or
mixture of chemicals that is used to develop the data.
(iii) "Photoperiod" The light and dark periods in a 24 hour
day. This is usually expressed in a form such as 17 hours light/
7 hours dark or 17L/7D.
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(iv) "Basal diet" The untreated form of the diet, such as
the diet obtained from a commercial source.
(2) The definitions in this section refer specifically to
the production and quality of eggs and the subsequent development
of these eggs up to the point where young are 14 days old.
(i) "Eggs laid" This term refers to the total egg
production during the test, which normally includes ten weeks of
laying. Values are expressed as numbers of eggs per pen per
season (or tes t).
(ii) "Eggs cracked" Eggs determined to have cracked shells
when inspected with a candling lamp. Fine cracks cannot be
detected without using a candling lamp and if undetected will
bias data by adversely affecting embryo development. Values are
expressed as a percentage of eggs laid by all hens during the
test.
(iii) "Eggs set" All eggs placed under incubation, i.e.,
total eggs minus cracked eggs and those selected for analysis of
eggshell thickness. The number of eggs set, itself, is an
artificial number, but it is essential for the statistical
analysis of other development parameters.
(iv) "Viable embryos (fertility)" Eggs in which
fertilization has occurred and embryonic development has begun.
This is determined by candling the eggs 14 days after incubation
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August, 1982
has begun. It is difficult to distinguish between the absence of
fertilization and early embryonic death. The distinction can be
made by breaking out eggs that appear infertile and examining
further. This distinction is especially important when a test
compound induces early embryo mortality. Values are expressed as
a percentage of eggs set.
(v) "Live 21-day embryos" Embryos that are developing
normally after 21 days of incubation. This is determined by
candling the eggs. Values are expressed as a percentage of
viable embryos (fertile eggs).
(vi) "Hatchabili ty" Embryos that mature, pip the shell, and
liberate themselves from their eggs on day 25, 26, or 27 of
incubation. Values are expressed as a percentage of viable
embryos (fertile eggs).
(vii) "14-day old survivors" Birds that survive for two
weeks following hatch. Values are expressed both as a percentage
of hatched eggs and as the number per pen per season (test).
(viii) "Eggshell thickness" The thickness of the shell and
the membrane of the egg at several points around the girth after
the egg has been opened, washed out, and the shell and membrane
dried for at least 48 hours at room temperature. Values are
expressed as the average thickness of the several measured points
in millimeters.
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August, 1982
(c) Test Procedures—-(1) Summary of test. (i) After birds
have been obtained they should be observed for health and
acclimated for at least two weeks.
(ii) Test birds should be randomly assigned to control and
various treatment groups.
(iii) The test substance should be thoroughly and evenly
mixed into the diet at concentrations specified in the test
rule. All treatment levels should be analyzed for test substance
concentrations at the beginning and midway through the test.
( iv) Birds should be weighed at the beginning of the test,
at 14-day intervals until the onset of laying, and at termination
of the test.
(v) Photoperiod should be carefully controlled on a short-
day basis during the initial exposure phase, then increased to
16-17 hours to induce egg laying.
(vi) Birds should be observed regularly for abnormal
behavior or mortality throughout the test.
(vii) Eggs should be removed daily and stored until there is
a sufficient quantity for incubation. All eggs should be candled
for cracks and cracked eggs removed. Once every two weeks, all
eggs produced that day should be analyzed for eggshell
thickness. Incubated eggs should be candled on day 14 and day
21. Hatching should be completed by day 27.
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(viii) Hatchlings should be maintained pens until they are
14 days old. Abnormal behavior or death should be reported.
Ducklings should be weighed on day 14.
( ix) A statistical analysis should be performed, preferably
by analysis of variance or regression analysis.
(x) The report should include all conditions, procedures,
and results. Data should be sufficiently detailed for an
independent statistical analysis.
(xi) All treated birds should be sacrificed and disposed of
properly. Control birds may be kept as breeding stock, but
should not be used in any other tests. Control offspring may be
reared and used in another test as adults.
(2) [Reserved]
(3) [Reserved]
(4) Definitive test—(i) Test subs tance. (A) The
concentrations of test substance in the diet will be specified in
the test rule. Generally, three treatment groups and a control
group will be used. The higher two treatment levels will be
multiples (often 5x, lOx, or 20x) of the lowest treatment
level. The highest treatment levels usually will be below lethal
levels, unless predicted exposure levels are high enough to
approximate lethal levels.
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August, 1982
(B) The material to be tested should be analytically pure
and the degree of purity should be reported along with the
percentage of each impurity at levels specified in the test
rule. If specifically required by a test rule for a particular
substance or mixture, the technical grade should be tested. The
test rule will specify the degree of purity or a range of
compositions of the technical grade material.
(ii) Controls . A concurrent control is required during
every test. The control birds should be from the same hatch as
the test groups. Control and test birds should be kept under the
same experimental conditions. The test procedures should be the
same for control and treated birds, except that no test substance
should be added to the diets of control birds. If a carrier is
used in preparation of the test diets, the same carrier should be
added to the diets of control birds in the highest concentration
used for test diets. The use of shared controls is acceptable
for concurrent tests as long as the same carrier is used for all
the tes ts.
(iii) Test groups and numbers of birds. (A) Either one of
two designs may be used for numbers of animals and pens. For one
design, each of the three treatment groups and the control group
should consist of a minimum of 8 replicate pens, with each pen
containing one male and three females. For the alternative
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design, each group should consist of 12 replicate pens containing
one male and one female per pen; the use of 20 replicate pens in
the control group may yield a test with greater statistical
power. Either design is acceptable as long as productivity
reaches the definitive values given in paragraph (c)(4)(xii) of
this section. Testing facilities using an experimental design
with which they are not familiar are advised to experiment first
without test substances in order to determine the feasibility of
obtaining acceptable productivity levels.
(B) All control and treatment birds should be randomly
distributed to pens from the same population.
(iv) Duration of test. (A) The test consists of three
phases following acclimation to test facilities. The initial
phase begins; with exposure of treatment groups to diet containing
the test substance and is typically six to eight week long.
After the initial phase, the photoperiod is manipulated according
to paragraph (c)(4)(v) of this section to bring the hens into
laying condition. This second phase ends with the onset of egg-
laying and is typically two to four weeks long. The final phase
begins with the onset of laying and lasts for at least eight
weeks, preferably ten weeks. A withdrawal study period may be
added to the test phase if reduced reproduction is observed. The
withdrawal period, if used, need not exceed three weeks.
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(B) Exposure of adult birds to the test substance should be
continuous throughout the test. Unless otherwise specified in
the test rule, test birds should be exposed to the test substance
for at least ten weeks prior to the onset of egg laying.
(v) Preparation for reproduction (photoperiod). (A)
Lighting regimes (photoperiod) are critical to successful
reproduction. Various photoperiod regimes have been demonstrated
to give acceptable results. Any photoperiod regime that results
in productivity that meets the definitive values given in
paragraph (c) (4)(xii) of this section is acceptable as long as
birds are exposed to treated diets a minimum of ten weeks prior
to the onset of laying. Regardless of the method selected,
lighting should be controlled carefully. It is important during
the initial phase to not interrupt the dark period unless
absolutely necessary.
(B) A suggested photoperiod regime would consist of
maintaining birds under a photoperiod of seven or eight hours of
light during the initial phase. At the end of the initial phase,
the photoperiod may be increased to 16-17 hours of light per
day. The photoperiod may be maintained at this level for the
remainder of the study, although an increase each week of 15
minutes per day is acceptable.
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(vi) Observations of record on adult birds. (A) Body
weights should be recorded for each adult bird at the beginning
of the treatment period, at 14-day intervals until the onset of
egg laying, and at termination of treatment. Birds may be
weighed during egg production phase of the study only if they are
not unduly stressed by the procedure. Food consumption should be
measured and recorded by pen at least as often as body weights
are measured prior to the onset of laying and at least bi-weekly
throughout the rest of the study.
(B) Observations on adult birds should be made at least once
a day. Any mortality or other signs of toxicity should be
described and recorded by date or day of test. Gross
pathological examinations should be conducted on all birds that
die during the test period, and for all survivors at the end of
the test. Analysis for test substance residues of two or more
tissues (e.g.., muscle, fat) is encouraged, but not required
(unless specified in the Test Rule).
(vii) Egg collection, storage, and incubation. All eggs
should be collected daily, marked according to the pen from which
collected, and should be stored at 16°C and 55-80 percent
relative humidity. Storage in plastic bags may improve
uniformity of hatching. Stored eggs should be turned daily. At
weekly or bi-weekly intervals, eggs should be removed from
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storage and be candled to detect eggshell cracks. Except for
eggs with cracked shells and those eggs removed for eggshell
thickness measurements all eggs should be set after candling for
incubation in a commercial incubator. If incubators are not
equipped to automatically turn eggs, they should be turned daily
by hand. During the incubation period, eggs should be maintained
at 37.5°C and approximately 70 percent relative humidity. Eggs
should be candled again on day 14 of incubation to determine
fertility and early death of embryos. A final candling should be
done on day 21 to measure embryo survival. On day 23, eggs
should be removed to a separate incubator or hatcher. Hatching
will normally be complete by the end of day 27.
(viii) Duckling maintenance. By day 27 of incubation, the
hatched mallard ducklings should be removed from the hatcher or
incubator. Ducklings should be either housed according to the
appropriate parental pen group or individually marked (such as by
leg bands) as to parental group and housed together. Ducklings
should be maintained in commercial brooder pens or pens of
similar construction. Pens should be constructed of galvanized
metal or stainless steel. Temperature in the pens should be
controlled, preferably by a thermostatic control device. A
temperature gradient in the pen from approximately 35°C to
approximately 22°C will allow young birds to seek a proper
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temperature. Temperature requirements for young birds typically
decline over this range from birth through the first several
weeks of life. Ducklings should be provided a standard
commercial duck starter ration, or its nutritional equivalent.
No test substance may be added to the diets of ducklings.
Ducklings should be maintained until they are 14 days old.
(ix) Observations of record on ducklings. The hatchability,
percentage of normal hatchlings, percentage of 14-day old
survivors, and number of 14-day old survivors per hen should be
recorded and reported. Ducklings should be observed daily from
hatching until they are 14 days old. Mortality, signs of
toxicity, and other clinical abnormalities should be recorded at
least cumulatively through day 5 and recorded by age from days 5
through 14. Average body weights should be recorded for
ducklings at day 14.
(x) Eggshell thickness. Once every two weeks all eggs newly
laid that day should be removed and measured for eggshell
thickness. Eggs should be opened at the girth (the widest
portion), the contents washed out (or used or saved for egg
residue analysis), and the shells air dried for at least 48
hours. The thickness of the shell plus the dried membrane should
be measured at a minimum of 3 points around the girth using a
micrometer calibrated at least to 0.01 mm units.
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(xi) Typical observed values. The values reported here
represent those observed from a few testing facilities under
their conditions. These values are not necessarily
representative of those from all facilities, however, if a
reproduction test does not meet or at least approach these values
for control birds, then there is likely to be a problem with test
procedures or conditions that should be investigated and
corrected. Typical values include:
(A) Eggs laid. Normal values for mallards - 28 to 38 eggs
per hen per season.
(B) Eggs cracked. Normal values for mallards - 0.6 percent
to 6 percent of eggs laid.
(C) Viable embryos (fertility). Normal fertility values for
mallards - 85 percent to 98 percent of eggs set.
(D) Live 21-day embryos. Normal values for mallards - 97 to
99 percent of viable embryos.
(E) Hatchability. Normal values for mallards - 50 percent
to 90 percent of viable embryos (fertile eggs).
(F) 14-day-old survivors. Normal values for mallards - 94
percent to 99 percent of eggs hatched.
(G) Eggshell thickness. Normal average values for mallards
- 0.34 mm to 0.39 mm.
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(xii) Definitive test criteria. (A) A test is unacceptable
if mallard duckling productivity in control groups does not
average fourteen 14-day old survivors per hen over a ten week
period.
(B) A test is unacceptable if the average eggshell thickness
in control groups is less than 0.34 mm.
(C) A test is unacceptable if more than 10% of the adult
control birds die during the test.
(5) [Reserved]
( 6 ) Analytical measurements — (1) Statistical analysis.
(i) Experimental groups should be individually compared to the
control group by analysis of variance. Other accepted
statistical methods may be used as long as they are documented
and described. In particular, regression analysis is highly
desirable if the data and number of dose levels allow the use of
this technique. Sample units are the individual pens within each
treatment level or control. Analysis should include:
(A) Body weights of adults.
(B) Food consumption of adults.
(C) Percentage of hens laying eggs. This should always be
determined when pens contain a single pair; if feasible, it
should be determined when pens contain groups.
(D) Number of eggs laid per pen.
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(E) Percentage of cracked eggs.
(F) Percent viable embryos of eggs set.
(G) Percent live 21-day embryos of viable embryos.
(H) Percent hatching of viable-embryos.
(I) Percentage of hatchlings that are normal.
(J) Percent 14-day-old survivors of normal hatchlings.
(K) Number of 14-day-old survivors per hen.
(L) Body weights of 14-day-old survivors.
(M) Eggshell thickness.
(ii) Analysis for Test Substance Concentrations. (A)
Samples of treated diets should be analyzed to confirm proper
dietary concentrations of the test substance. If samples cannot
be analyzed immediately, they should be stored appropriately
(e.g., frozen at a tempererature of -15°C or lower) until
analysis can be performed. Analyses should be conducted on all
test substance concentrations at the beginning of the test period
and again 10 to 12 weeks later. If not otherwise available, data
should be generated to indicate whether or not the test substance
degrades or volatilizes. If the test substance is known or found
to be volatile or labile to the extent that 25 percent or more
loss occurs within one week, then test substance diets should be
prepared (freshly or from frozen concentrate) at a frequency that
will prevent more than 25 percent loss of test substance.
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(B) The assay method used to determine actual concentrations
should be reported according to paragraph (e)(l)(vi) of this
section.
(C) Analysis of basal diet. A nutrient analysis of the
basal diet should be included in the test report. For
commercially prepared basal diets, the list of ingredients
supplied by the manufacturer is normally sufficient, if it is
detailed. The composition of any vitamin or other supplements
should also be reported.
( d ) Test conditions — (1) Test species ( i ) Selection.
(A) The mallard, Anas platyrhynchos L., is the test species.
Test birds should be pen-reared. They may be reared in the
laboratory or purchased from commercial breeders. Rearing stock
and/or test birds should be obtained only from sources that have
met the requirements for "U.S. Pullorum-Typhoid Clean"
classification. Birds should be obtained only from sources whose
colonies have known breeding histories. If possible, a history
of rearing practices for test birds should be obtained and made
available upon request. This history should include lighting
practices during rearing, disease record, drug and any other
medication administered, and exact age. Test birds should be
phenotypically indistinguishable (except for size) from wild
stock. Conscientious breeders of such birds will periodically
15
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August, 1982
outbreed their flocks with genetically wild stock in order to
maintain a genetic composition that approximates the
heterogeneity of naturally occurring birds.
(B) All control and experimental birds used in a test should
be from the same source and strain. If shipped, all birds should
be examined following shipment for possible physical injury that
may have occurred in transit. All birds should have a health
observation period of at least two weeks prior to selection for
treatment. Birds should be in apparent good health. Deformed,
abnormal, sick, or injured birds should not be used. A
population of birds should not be used if more than 3 percent of
either sex die during the health observation period. Birds
should not have been selected in any way for resistance to toxic
substances. Birds should not have been used in a previous test,
either in a control or treatment group. Offspring of birds used
in a treatment group in a previous test should not be used, but
offspring of birds used as a control in a previous test are
acceptable.
(C) Test birds should be approaching their first breeding
season and should be at least seven months old. All test birds
should be the same age within one month. The age of test birds
should be reported.
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August, 1982
(D) Mallards should be acclimated to test facilities and
untreated basal diet for at least two weeks. Acclimation may be
in the actual pens used in the test or in identical pens. The
acclimation period may coincide with the health observation
period. Birds should be randomly assigned to treatment and
control pens. However, when birds in a pen are incompatible,
they may be rearranged within a control or treatment group at any
time prior to initiating treatment.
(E) During holding, acclimation, and testing, birds should
be shielded from excessive noise, activity, or other
disturbance. Birds should be handled only as much as is
necessary to conform to test procedures.
(ii) Diet—(A) Adult birds. (JJ A standard commercial
duck breeder ration, or its nutritional equivalent, should be
used for diet preparation. This ration or basal diet should be
used for both control and treatment birds and should be constant
throughout the duration of the study. Antibiotics or other
medication should not be used in the diet or water of breeding
birds. It may not be possible to obtain food that is completely
free of pesticides, heavy metals, and other contaminants.
However, diets should be analyzed periodically for these
substances and should be selected to be as free from contaminants
as possible. A nutrient analysis (quantitative list of
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ingredients) of the diet should be included with the test report.
(_2_) The test substance should be mixed into the diet in a
manner that will ensure even distribution of the test substance
throughout the diet. If possible, the test substance should be
added to the diet without the use of a carrier or diluent. If a
diluent is needed, the preferred diluent is distilled water; but
water should not be used for test substances known to hydrolyze
readily. When a test substance is not water soluble; it may be
dissolved in a reagent grade evaporative diluent (e.g., acetone,
methylene chloride) and then mixed with the test diet. The
solvent should be completely evaporated prior to feeding. Other
acceptable diluents may be used, if necessary, and include table
grade corn oil, propylene glycol, and gum arabic (acacia). If a
diluent is used, it should comprise no more than 2% by weight of
the treated diet, and an equivalent amount of diluent should be
added to control diets.
(_3_) Diets may be mixed by commercial or mechanical food
mixers. Other means are acceptable as long as they result in
even distribution of the test substance throughout the diet.
Screening of the basal diet before mixing is suggested to remove
large particles. For many test substances, it is recommended
that diets be mixed under a hood. Frequently, the test substance
is added to an aliquot of the basal diet to form a premix
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concentrate. The premix concentrate should be stored so as to
maintain the chemical concentration. For final preparation of
test diets, the premix is mixed with additional basal diet to
form the proper concentrations. The frequency with which final
treated diets are prepared will depend upon the stability and
other characteristics of the test substance. Unless otherwise
specified in the test rule or determined by degradation or
volatility studies, it is recommended that final diets be
prepared weekly, either fresh or from a concentrate. For
volatile or labile test substances, test diets should be mixed
frequently enough so that the concentrations are not reduced from
initial concentrations by more than 25 percent. Analysis of
diets for test substance concentrations is required as specified
in paragraph (c)(6)(ii).
(_4_) Clean water should be available ad_ libitum. Water
bottles or automatic watering devices are recommended. If water
pans or bowls are used, water should be changed daily or more
of ten.
(B) Young birds . Young birds produced during the test
should be fed a commercial duck starter ration, or its
nutritional equivalent. No test substance should be added to the
diets of young birds. No antibiotics or medication should be
used in the diet.
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August, 1982
(2) Facilities. (i) Mallards should be housed in breeding
pens or cages of adequate size conforming to good husbandry
practices. Space requirements for mallards have not been well
defined, but it is recommended that there be at least 10,000
square centimeters (approximately 5.4 square feet) of floor space
per bird. Documentation that reproductive parameters and health
of birds are not adversely affected should be provided for cages
much smaller than this area. The preferred construction
materials are stainless steel, galvanized sheeting, and wire
mesh. For enclosed cages, floors and external walls may be wire
mesh; ceilings and common walls should be solid sheeting. Open-
topped pens may be constructed of the same materials for the side
walls with open tops and wire mesh or concrete floors. Concrete
floors should be covered with litter such as straw, wood
shavings, or sawdust. Other construction materials, except wood,
are acceptable if they can be kept clean. Wood may be used as
vertical framing posts for the support of wire mesh or for
horizontal framing along the top of the pen. Wood should not be
used for floors or lower sides of pens unless it has been coated
with a non-adsorbent material such as perfluorocarbon plastic
(e.g., Teflon) or unless the wood is replaced between tests.
(ii) Pens should be disassembled (if feasible) and should be
cleaned thoroughly between tests. Steam cleaning of enclosed
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August, 1982
cages is recommended. Enclosed cages may be brushed thoroughly,
as an alternative method. For open-topped pens, the sides and
vertical supports should be thoroughly brushed. Any used floor
litter should be discarded. The floor composition will dictate
methods used to clean the floor. If litter is used on the floor,
it should be fresh and clean when birds are placed in the pen.
The use of detergents or bleach is acceptable, but other chemical
disinfectants (such as quaternary ammonium compounds) should not
be used. When necessary to control disease vectors, hot or cold
sterilization, techniques are recommended, as long as such
techniques will not leave chemical residues on the cages. For
cold sterilization, ethylene oxide is recommended.
(iii) Pens should be kept indoors in order to better control
lighting, temperature, humidity, and other factors. Outdoor pens
may be used only during the normal breeding season. The
photoperiod should be carefully controlled, preferably by
automatic tiraers. A 15-30 minute transition period is
desirable. The photoperiod regime is described under paragraph
(c)(4)(v). Lights should emit a spectrum simulating that of
daylight. The use of shorter wave-length "cool-white"
fluorescent lights that do not emit the daylight spectrum should
be avoided. Illumination intensity should be about 6 foot-
candles at the level of the birds.
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EG-17
August, 1982
(iv) Temperature and humidity should be controlled during
the study. Recommended levels are 21°C and 55 percent relative
humidity. Temperature for indoor tests should be recorded at
least weekly at the same time of day and should be reported. For
tests conducted without temperature control, temperature minimums
and maximums should be recorded daily. Continuous temperature
monitoring is desirable. Temperature recording should be made at
level of 2.5-4 cm above the floor of the cage. Recording of
approximate humidity levels is also desirable. Good ventilation
should be maintained. Suggested ventilation rates are 4 changes
per hour in winter and 15 changes per hour in the summer.
(v) If facilities are being used for the first time, it may
be desirable to allow birds to breed in the facility prior to
testing in order to ensure that controls will have acceptable
productivity according to the requirements given in paragraphs
(c)(4)(xi) and (xii)].
(3) [Reserved]
(e) Reporting. (1) The test report should include the
following information:
(i) Name of test, sponsor, test laboratory and location,
principal investigator(s), and actual dates of beginning and end
of test.
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August, 1982
(ii) Name of species tested (including scientific name), age
of birds (in months) at the beginning of the test, source of
birds, and body weights for adult birds throughout the test.
(iii) Description of housing conditions, including type,
size, and material of pen, temperature, humidity, photoperiod and
lighting intensity, and any changes during the test.
(iv) Detailed description of the basal diet, including
source, composition, diluents (if used), and supplements (if
used). A nutrient analysis of the basal diet should be included.
(v) Detailed description of the test substance including its
chemical name(s), source, lot number, composition (identity of
major ingredients and impurities), and known physical and
chemical properties pertinent to the test (e.g., solubility,
volatility, degradation rate, etc.).
(vi) The number of concentrations used, nominal and measured
concentrations of test substance in each level, assay method used
to determine actual concentrations, storage conditions and
stability of treated diets, number of birds per pen and number of
replicate pens per concentration and for controls.
(vii) Acclimation procedures and methods of assigning birds
to test pens, including method of randomization, and any
rearrangements due to incompatibility.
(viii) Frequency, duration, and methods of observation.
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August, 1982
(ix) Description of any signs of intoxication, including
time of onset, duration, severity (including death), and numbers
affected, including accidental deaths or injuries.
(x) Food consumption per pen and any observations of
repellency or food palatability.
(xi) Method of marking all birds and eggs.
(xii) Details of autopsies.
(xiii) Egg and hatching data in summary and by pen per week
in sufficient detail to allow an independent statistical
analysis. Data should be presented for all of the parameters
listed in paragraph (c)(6)(i). The number of eggs set also
should be reported.
(xiv) Egg storage, incubation, and hatching temperatures,
relative humidities, and turning frequencies.
(xv) Observations of health and weights of young at 14 days
of age.
(xvi) Location of all raw data storage.
(xvii) Methods of statistical analysis and interpretation of
results .
(xviii) Anything unusual about the test, any deviation from
these procedures, and any other relevant information.
(2) In addition, the following information should be
available upon request:
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August, 1982
(i) A general description of the support facilities.
(ii) A description of the Quality Control/Quality Assurance
program, including the Average Quality Level for the program
element performing the test, procedures used, and documentations
that these levels have been achieved.
(iii) The names, qualifications, and experience of personnel
working in the program element performing the test, including the
study director, principal investigator, quality assurance
officer, as well as other personnel involved in the study.
(iv) Standard operating procedures for all phases of the
test and equipment involved in the test.
(v) Sources of all supplies and equipment involved in the
test.
(vi) Diagram of the test layout.
(vii) Originals or exact copies of all raw data generated in
performing the test.
(viii) A detailed description, with references, of all
statistical methods.
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ES-14
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TECHNICAL SUPPORT DOCUMENT
FOR
MALLARD REPRODUCTION TEST
OFFICE OF TOXIC SUBSTANCES
OFFICE OF PESTICIDES AND TOXIC SUBSTANCES
U.S. ENVIRONMENTAL PROTECTION AGENCY
WASHINGTON, D.C. 20460
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TABLE OF CONTENTS
Subject
I. Purpose
II. Scientific Aspects
General
Issues
Test Procedures
Test Substance Concentrations 7
Controls 7
Test Groups and Numbers of Animals 8
Duration of Test 9
Preparation for Reproduction (photoperiod) 10
Observations and Measurements 11
Adult Birds 11
Ducklings 12
Eggshell Thickness 13
Typical Observed Values 13
Egg Collection, Storage and Incubation 14
Required Analysis 16
Statistical 16
Test Substance Concentrations 16
Basal Diet 17
Acceptability Criteria 18
Test Conditions 19
Test Species 19
Selection 19
Maintenance of Test Species 23
Acclimation 23
Diet 23
Facilities 23
Environmental Conditions 25
Temperature (See Section II.C.3) 25
Humidity (See Section I I.e.3) 25
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Subject Page
Reporting 25
III. Economic Aspects 26
IV. References 28
11
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Office of Toxic Substances ES-14
August, 1982
TEST SUPPORT DOCUMENT FOR MALLARD REPRODUCTION TEST
I. Purpose
The purpose of this document is to provide the
scientific background and rationale used in the development
of Test Guideline EG-17 which uses Mallards to evaluate the
effect of chemical substances on reproduction. The Document
provides an account of the scientific evidence and an
explanation of the logic used in the selection of the test
methodology, procedures and conditions prescibed in the Test
Guideline. Technical issues and practical considerations
are discussed. In addition, estimates of the cost of
conducting the test are provided.
II. Scientific Aspects
A. General
The earliest investigations of the effects of chemicals
on reproduction of native birds were in the 1950s (DeWitt
1956, Genelly and Rudd 1956). Chemicals were administered
in the diet, but procedures varied. Laboratory
investigations of reproductive effects of pesticides
continued at Patuxent Wildlife Research Center, but methods
were not reported well (DeWitt et al. 1962, DeWitt et al.
1963). In 1964, a very brief protocol for reproduction
studies was developed at that center (Stickel and Heath
1965). This protocol outlined the egg parameters to be
studied. Ratcliffe (1967), in a classic paper, correlated
the decline of certain avian populations with thin eggshells
that apparently had resulted primarily from exposure to DDT
and DDE. Heath et al. (1969) presented the first clearcut
experimental data showing that DDE caused thin eggshells in
mallards. The protocol used by Heath et al. (1969),
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August, 1982
although not fully detailed, formed the basis for most
future avian toxicological reproduction studies. In 1968,
USDA developed a guideline for evaluating reproductive
effects of toxic chemicals to birds (US EPA 1975). This
guideline apparently was developed in conjunction with
Patuxent personnel as it bears a great similarity to methods
used at Patuxent, but was available prior to the publication
of Heath et al. (1969). The Agency (US EPA 1975) developed
composite protocols for reproduction tests from the limited
published information and unpublished information,
especially the USDA protocol which was presented as an
exhibit. No complete, suitable protocol for bobwhite was
available from the published literature at that time. More,
but insufficient, literature was available for mallards,
particularly as relates to eggshell thinning (US EPA, 1975)
since Heath et al. (1969, 1972b) and Heath and Spann (1973)
had published the results of mallard reproduction studies.
The Agency's pesticide guidelines were revised (US EPA
1978a), but the basic method of the earlier guidelines for
reproduction tests was retained. There have been few
mallard reproduction tests published even in recent years,
although a number of tests have been conducted, classified
as confidential, and submitted to the Agency to support
pesticide registration. Those that have been published
have been written by researchers familiar with bird tests
and many details of protocol have not been included in the
reports. ASTM (1979a) has prepared a draft avian
reproduction method. Although this ASTM protocol is similar
to the Office of Pesticide Programs protocol (US EPA 1978a),
it is designed for a variety of species. Thus, the history
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of avian reproduction test methodology is basically a
history of Patuxent and EPA methods. This guideline
continues the trend because these methods appear to be the
most appropriate for developing data needed to make
regulatory decisions and because no other methods have
become accepted widely.
Even though the basic method is similar for most
investigators, there have been a few points of difference or
controversy. Yet there are very few data to address these
differences and little ongoing research that might resolve
conflicts. To promote uniformity and comparability of
tests, some recommendations and requirements are
standardized in this test guideline. Where such
recommenda.tions and requirements are controversial and are
not sufficiently addressed by published data, it is hoped
that research will be stimulated to resolve questions. If
feasible, conditions and procedures that approximate natural
conditions have been selected in preference to other
options. Controversial points are addressed under Issues in
this support document.
The use of avian reproduction tests in the assessment of
chemical impacts on the environment is based on several
factors. First, birds are an obvious and important
component of the environment. Congress has indicated
repeatedly that birds are worthy of protection by passing
such laws as the Lacey Act of 1900, Migratory Bird Treaty
Act of 1918, Migratory Bird Conservation Act of 1929,
Pittman-Robertson Act of 1937, Fish and Wildlife Act of
1956, Endangered Species Act of 1973, and others. The
United States also has entered into treaties with Great
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Britain and Canada (1916), Mexico (1937), Japan (1974), and
Russia (1976) for the protection of migratory birds. The
people of the United States also have indicated a desire to
protect birds through their support of the Audubon Society,
Nature Conservancy, and other environmental groups.
Sportsmen's organizations support protection of birds,
although their interests often focus heavily on game birds.
Second, birds have a definite economic importance.
Federal and State Agencies spend large sums for the
preservation and propagation of birds. Hunters and
birdwatchers also spend substantial sums in pursuit of their
pastimes. Less measurable, but of definite importance, is
the substantial role of birds in insect control.
Birds have an important ecological role. Insectivorous
birds consume huge quantities of insects and other
invertebrates, many of which are considered pests. Small
mammals and other vertebrates or plants are consumed by
various birds, sometimes to the extent that birds have an
important effect on populations. In turn, birds are
consumed by birds of prey, mammals, and other vertebrates.
Excretory products of birds provide nutrients for plankton
and other microorganisms that in turn are food for larger
organisms. Birds are important in pollination of some plants
and in dispersal of others. Because of their mobility, the
effects of birds are not restricted to specific locations.
Finally, birds are among the more sensitive terrestrial
vertebrates. Because of their high metabolic rate, high
body temperature, and the demands of flight, they require
more energy relative to their size than most other
animals . The energy requirements lead to greater food
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intake and thus to greater toxicant intake when a toxicant
is in or on their food. There are abundant data showing
that some birds, particularly raptors, pelicans, and
waterfowl, are very sensitive in their reproductive
responses to toxic chemicals (e.g., Ratcliffe 1967, Anderson
and Hickey 1972) and that some species have suffered from
drastic population reductions apparently as a result. Avian
reproduction is unique, and no surrogate is adequate to
predict effects on eggshells, the primary mode of impairment
for many chemicals such as DDE (Heath et al. 1969).
Avian reproduction tests are extremely valuable in
assessing the potential population effects of exposure of
birds to toxic chemicals. The route of intake simulates
natural exposure to chemicals on or in the food. Most
physiological effects can be assessed under laboratory
conditions, although many behavioral effects, such as nest
desertion, are difficult to study in the laboratory. A
positive finding of impairment in the laboratory is highly
predictive (qualitatively) in the field when exposure is the
same. However, negative findings in the laboratory may not
preclude adverse effects under field conditions. Thus, some
extrapolation may be made from laboratory to field, but
quantitative extrapolation is risky.
1. Issues
The avian reproduction test guideline and support
document contain some controversial points. Data are
insufficient or absent to support either side of most
points. For other points, there may be data supporting each
side. A number of controversial points have been selected
as potential research projects. Issues are merely
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identified below and are discussed in the appropriate
sections of this document.
o Is productivity from tests run out of normal season
sufficient to evaluate the potential for
reproductive impairment?
o Should methods involving natural incubation of eggs
be incorporated into tests?
o Should the current use of first year birds only be
continued or should proven breeders be used? Would
either age produce results suitable for comparison?
o What carriers, if any, should be used or allowed
for incorporation of the test substance into the
diet?
o Are commercial foods adequate, or is there too much
variation and/or contamination of commercial foods?
o How often should treated diets be mixed? Is there
an advantage in mixing diets with decreasing
concentrations to simulate natural degradation of
test substances?
o What is the optimum number of birds to be tested in
order to attain statistically valid results and
still be cost-effective? Can tests using pairs
only of birds in pens be successfully conducted by
a variety of testing facilities?
o Can more useful results be obtained by testing
enough dose levels to use linear regression
analys is?
o Should outdoor tests be allowed?
o Should medication be allowed as is needed to treat
individual sick birds
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Are the typical values (Section 2.1.6) and test
acceptability criteria (Section 2.1.9) for
productivity and other egg parameters realistic for
a wide diversity of testing facilities, or are they
really only valid for a few?
Test Procedures
1. Test Substance Concentrations
Test substance concentrations in the diet will be
specified in the test rule. These concentrations will be
based upon the properties of the test substance, the lethal
and no-effect levels, if known, or the amount of test
substance known or likely to be found in the environment.
Three concentration levels are specified because, for many
test substances, three levels will allow for a dose-response
regression analysis from which a no- effect level can be
calculated. (See section 2.1.8 on statistics for further
discuss ion) .
2. Controls
Concurrent controls are required for every test to
assure that any observed effects are a result of ingestion
of the test substance and not to other factors. Such other
factors may include environmental factors such as
temperature, lighting, vapors, sensitive or stressed test
birds, etc. If a diluent is used in mixing the diet, this
diluent also is used in the untreated diets in the same
concentration as it occurs in treated diets. In effect,
this results in a diluent, but no completely negative,
control. Diluent choices are based upon their lack of
toxicity (e.g., water, completely evaporated acetone) and it
is not considered necessary, therefore, to have an
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additional negative control when a diluent control is used.
3. Test groups and numbers of animals
A minimum of 8 replicate pens with one drake and three
hens or 20 replicate pens with one drake and one hen is
required for each test concentration. The number of
replicates is needed to achieve a test with statistical
significance. There is enough variation in productivity of
mallard hens that fewer replicates would be very insensitive
to all but the most severe effects.
The recommended arrangement is for use of group pens
containing three females each. This is based upon the
rather aggressive behavior of courting males that could
result in severe stress to the hen, if only one hen were
present. In mallard reproduction tests, Heinz (1976a)
obtained satisfactory reproduction with mallard pairs. He
attributed his success primarily to having a nest box and
flowing water (Heinz, personal communication). Heinz has
pushed strongly for the adoption of his method. Testing of
pairs only is a practice that should lead to statistically
stronger tests and will provide a better indication of
effects on individuals. However, it is offered only as an
option in this guideline because it is unknown at this time
if other researchers will be able to achieve good results.
The alternative arrangement of at least 20 mallard pairs per
level is suggested to accommodate those test facilities that
believe they will have success or that wish to contribute to
the development of methods. A warning is included in the
guideline so that relatively inexperienced investigators
should not expect unqualified success. The additional pens
in pair testing will strengthen notably the statistical
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analysis of the test. This alternative is included because
it is statistically stronger than the more familiar method
of group testing.
4 . Duration of Test
The avian reproduction test lasts approximately 22
weeks. The initial part of the test is an exposure phase
where birds are receiving treated diet. Exposure to treated
diets begins with this phase and allows for the test
substance to act upon the reproductive mechanisms of the
body, and also for birds to accumulate residues of
lipophilic and other test substances. The development of
reproductive capacity actually begins months before egg
laying starts (Wolfson 1964). Therefore, exposure to the
test chemical should be well in advance of egg laying, if
the test is to investigate reproductive effects in general.
The second phase, following lengthening of the
photoperiod, directly brings the birds into readiness for
egg laying., The duration of this phase is dependent upon
the response of the test birds.
The third phase is the egg-laying portion of the test.
This is to be a minimum of eight weeks. The duration of
this phase is based upon two main factors. First, it is
important to determine if egg laying is within normal levels
or if it declines, or otherwise varies, over a period of
time. This yields information on speed of action and has a
role in using the test for assessment purposes. Second, the
eight week period is needed to provide sufficient data for a
strong statistical analysis.
A withdrawal period is optional when impairment has been
detected. This period may provide data on recovery of
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reproductive capacity that could be useful for assessment
purposes.
5. Preparation for reproduction (photoperiod)
Because photoperiod is critical to reproduction, it
should be controlled in indoor tests. Under natural
conditions, photoperiod is lengthening gradually just prior
to the reproductive season. The earlier mallard
reproduction tests were conducted outdoors (Stickel and
Heath 1965, Heath et al. 1969, 1972b, Heath and Spann 1973),
and therefore, used natural photoperiod. More recently,
mallard reproduction tests submitted for pesticide
registration purposes have been conducted indoors and egg
laying has been induced by increasing photoperiod to 16 or
17 hours of light per day. The ASTM (1979a) and US EPA
(1978a) both recommended seven hours of light in the first
phase and 16-17 hours light in the second phase. Without
giving supporting data, both of these protocols stated that
the regime is for maximum egg production, and both specify
at least an option of gradually increasing the length after
egg laying has started. The photoperiod in this guideline
is based on the above data and a lack of any suggested
alternatives.
The dark period in the photoperiod should not be
interrupted, even briefly, except as absolutely necessary.
Kirkpatrick (1955) found that even as little as 15 minutes
interruption on a short day/long night regime caused an
increase in gonadal development of bobwhite, and all birds
exposed to a 60 minute interruption became fully active
sexually. It is highly probable, on the basis of
Kirkpatrick's (1955, 1959) data, that it is the length of
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the dark period rather than the light period that controls
bobwhite reproductive preparation. It is likely that this
holds for mallards also, although no definitive data were
found.
Kirkpatrick (1955) also tested light intensity as a
factor in avian reproduction. Responses of bobwhite were
very similar at intensities of 0.1 to 100 foot candles on a
17 hour light photoperiod. With shorter days, the 0.1 foot
candle intensity did not achieve the same results as 1,10,
and 100 foot candles. These data support the approximate 6
foot candle intensity for bobwhite that was suggested by
ASTM (1979a) and US EPA (1978a). In the absence of other
data, the same light intensity has been specified for
mallards.
6,. Observations and Measurements
a. Adult Birds
Observations of food consumption, body weights, and
signs of toxicity are required. Body weights are required
three times during the test. More frequent body weights
might be informative, but the stress of handling may offset
the collection of data, particularly for laying hens. Food
consumption is to be estimated at frequent intervals. This
will provide data both on test substance ingest ion and on
energy intake for test birds. The latter data will be
particularly helpful in the absence of frequent weighings.
Clinical signs of toxicity contribute substantially to
the analysis of the data, in addition to providing
information on the mode and speed of action. If dose levels
are finely tuned, there should be minimal observed acute
toxicity. However, without adequate preliminary data, one
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or more of the test levels may cause lethal or notable
sublethal effects. Such effects may affect the results of
the test without directly acting on the specific
reproductive parameters being investigated. For example, a
severely stressed bird may not be able to mobilize internal
resources to produce eggs. Signs of toxicity will give
valuable information in evaluating the results of the test.
b. Duckling
Mallard eggs have been incubated in reproduction tests
for as long as 27 days (Heath and Spann 1973). Greenwood
(1975) reported average incubation lengths of 24.6 days for
a wild mallard strain and 25.5 days for a game farm
strain. US EPA (1978a) stated that eggs should be removed
from hatchers on day 27. The meager data available suggest
that 27 days is sufficient and this incubation length has
been selected for this guideline.
Because the pen is the basic unit for statistical
analysis, it is necessary that the eggs and hatchlings be
identified as to pen of origin. This can be done either by
housing all eggs or ducklings from one pen together, or by
individually marking each egg and bird.
Environmental conditions for young ducklings have not
been reported in available reproduction papers (Heath et al.
1969, 1972b, Heath and Spann 1973), nor have the US EPA
(1978a) or ASTM (1979a) reproduction protocols suggested any
housing conditions for ducklings. However, ASTM (1979b) and
US EPA (1978b) dietary test protocols both have recommended
procedures for housing young ducklings. Parameters given in
these references include temperature of 35°C with a lower
temperature outside the cage to provide a gradient,
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galvanized brooder cages maintained indoors, and a
commercial duck starter ration. These conditions have been
included in this guideline.
Observations on ducklings are necessary to determine if
and when toxicity might be expressed in the offspring.
Although most test substances exert their action prior to
hatching, some may affect growth, development, or survival
of ducklings.
c. Eggshell Thickness
The classic example of reproductive impairment in birds
is the eggshell thinning effect of DDE, a metabolite or
degradate of DDT (Tucker and Leitzke 1979). It is important
to measure eggshell thickness because as little as 11%
thinning can have an effect on avian populations (Tucker and
Leitzke 1979). Techniques given in this guideline are
widely used (Heath and Spann 1973, ASTM 1979a, US EPA
1978a), although many papers do not give a full description
of the process. It is particularly important for several
measurements to be made around the girth of mallard eggs, in
order to average out any aberrant single measurements that
could affect results if only one or two measurements are
taken.
d . Typical Observed Values
The typical observed values presented in this guideline
have been taken from the Agency's pesticide guidelines (US
EPA 1975, 1978a) . A number of avian reproduction tests have
been submitted to the Agency for the purpose of pesticide
registration and have been classified as confidential. Most
of the tests that have been conducted during the normal
reproductive season have achieved for control birds the
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typical values as presented in the test guideline. Heath
and Spann (1973) and Heath et al. (1969, 1972b) achieved
these values in control birds insofar as could be determined
from their data.
The values presented in the guideline have been
identified as an issue. It is known that some testing
facilities routinely meet these norms. However, it is not
known if the values are realistic when many testing
facilities are considered.
7. Egg Collection, Storage, and Incubation
Egg storage prior to setting for incubation has been
reported for mallards in only a few papers by one senior
author and has been 16°C and 55% humidity (Heath et al.
1969, 1972b, Heath and Spann 1973). Stromberg (personal
communication) stated that he stored bobwhite eggs at 16°
and at as high a humidity as could be obtained from
evaporating water; this was up to 80%, but at least 50%
relative humidity. He also stated that turning the eggs
daily was important to keep part of the shell and egg from
excessive drying. US EPA (1978a) recommends 16°C and 65%
relative humidity. This guideline specifies 16°C, but only
a range of relative humidity between 55% and 80%, since many
testing facilities do not have the means to control storage
humidity and since this range has produced good viability.
Several papers were found that reported incubation
temperatures and humidities for mallards. Temperatures were
consistently 37.4-37.5°C and humidities ranged from 57-80%
(Heinz 1976a, 1976b, Holmes et al. 1978, Greenwood 1975).
Neither ASTM (1979a) nor US EPA (1978a) specify incubation
temperatures although the latter does recommend 39°C and 70%
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relative humidity during the hatching phase from day 23 to
day 27. Stromberg (personal communication) stated that
99.75°F (37.6°C) and 50-70% relative humidity are standard
commercial poultry incubation temperatures and that all
testing facilities with which he is acquainted use
commercial poultry incubators. A temperature of 37.5°C and
relative humidity of approximately 70% have been specified
in this mallard test guideline.
Candling procedures are a standard practice for
determining eggshell cracks, fertility, and embryo
survival. These procedures have been used in most mallard
tests, but timing has been inadequately reported. US EPA
(1978a) specifies, for mallard, candling on day 0 for
cracks, on day 14 for fertility and on day 21 for embryo
survival. This timing has been adopted in this guideline
because it will permit comparisons of test data with
existing studies in Agency files. At least one major avian
testing laboratory protocol uses the same timing.
US EPA (1978a) recommended moving eggs to a separate
hatcher or incubator for hatching, but gave no reasons.
None of the available published papers mentioned this
procedure. However, Stromberg (personal communication)
stated that this procedure is highly recommended as a means
to minimize disease vectors. This procedure has been
recommended in the guideline.
This test guideline does not consider the effects on
incubation behavior, nest desertion, and care of young.
Techniques for investigating the effects of chemicals on
these behaviors are in their infancy and, at present, are
prohibitively expensive. Agency research has been proposed
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to investigate methods of incorporating natural incubation
into future reproduction guidelines at a reasonable cost.
8. Required Analyses
a. Statistical
The statistical analysis of avian reproduction studies
typically has been analysis of variance (Heath and Spann
1973, Heinz 1976a, US EPA, 1978a, ASTM 1979a) using the
parameters set forth in this guideline. Angular
transformations have been used to a lesser extent (Heath et
al. 197 2b). Regression analysis is a powerful tool that may
be used if there are data at sufficient dose levels to
obtain a dose-response curve. Regression analysis has been
recommended, as an option by ASTM (1979a), and methods
directed toward regression analysis are being developed
(Stromberg, personal communication). It is the opinion of
the author of this test guideline that regression analysis
is a more useful tool than currently typical methods because
it yields a dose response curve and this curve can be used
for extrapolation. Analysis of variance provides only
significance at a particular level and does not lend itself
to extrapolation on a reliable basis. At the present time,
methods and background work have not been developed for a
test oriented primarily towards a dose-response curve. More
dose levels would be needed, possibly with fewer animals per
level. This subject may be included in the Agency's
research projects. In the meantime, this guideline uses
methods appropriate to analysis of variance.
b. Test Substance Concentrations
Samples of treated diets will be analyzed to determine
the actual levels to be used in the test. Analysis will
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help to detect mathematical errors in calculating
concentrations, technicians' errors in mixing diets, and
manufacturers' errors in determining the amount of active
ingredient in a test substance (Heinz, personal
communication). All test substance concentrations will be
analyzed so that, even with only three test concentrations,
dose response curve (if obtained) would be based on measured
concentrations, rather than on nominal levels. A second
analysis about midway through the test will corroborate
initial levels.
c. Basal Diet
Most testing facilities use diets prepared by commercial
feed companies. Some facilities may have a commercial
company prepare a diet to order. Normally, such diets are
supplied with a quantitative list of ingredients, and such a
list should be supplied with the test report. If there are
supplements added to the diet, a list of all supplemental
ingredients also should be submitted. Analysis of
ingredients in the basal diet is important because there are
a number of potential test substances, such as certain
metals, that may interact with components of the diet and
possibly affect the results of a test. A nutrient analysis
will allow for a better evaluation of such results. In
addition, it is possible that dietary deficiencies or
imbalance of ratios of nutrients also could affect the
results. Even though commercial companies normally supply a
nutritionally adequate diet, it is important to know the
components because no rigid requirements exist for the type
and constitution of the diet used.
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9. Acceptability Criteria
Test acceptability is dependent upon following the
mandatory requirements and achieving acceptable control
productivity and survival. When control values do not reach
the typical observed values as discussed in Section 2.3.6,
it is very likely that there are problems with some aspect
of the test. Since reproductive parameters of treated birds
are compared with controls, it is essential that values for
control birds are normal as determined from similar
reproduction studies.
Rather than requiring every parameter to achieve normal
values, it was decided that test acceptability should be
based on the final parameter of 14-day old survivors per
control hen as one criterion. For mallards, the requirement
of 14 survivors per hen over a ten week period was selected
because data in the Agency's pesticide files show that
acceptable tests have achieved and usually surpassed this
level. Heath et al. (1969) reported 16.1 and 16.4 control
survivors per hen for eight week seasons. Heath and Spann
(1973) reported 6.1 and 6.2 control survivors per hen for
four week seasons, which values are equivalent to 15
survivors per hen for a ten week season. Heinz (1979)
achieved an annual average of 46 7-day survivors per control
hen for 3 years, using proven breeders and a season of
unknown length.
Eggshell thickness of control birds is also a criterion
of test acceptability because thin eggshells among control
birds are usually a sign of inadequate diet, which in turn
could affect other aspects of the test. A number of mallard
reproduction tests have had average control eggshell
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thickness of .365 to .393 mm (Heath et al. 1969, 1972b,
Heath and Spann 1973, Heinz 1979). Heath and Spann (1973)
also reported individual extremes of .330 and .420 mm.
Holmes, et al. (1978) reported .343 and .363 mm average
thickness for unfertilized eggs from mated control
females. The Agency's pesticide guidelines (US EPA 1978a)
state that normal eggshell thickness for mallards is .31 -
.33 mm. On the basis of the other published data cited
above and additional reproduction studies in the Agency's
pesticide files, it appears that .31-. 3 3 mm is
unrealistically low. The normal values given in section
(h)(ll) of the test guideline are .35-.39 mm, however, at
the present time, it seems appropriate to lower the
acceptable level to .34 mm. This may be revised pending
receipt of additional data and public comment.
A well conducted test with adult birds should not result
in any but an occasional mortality in control groups, even
though the test is relatively long. This is especially true
since this guideline provides for rearrangement of
incompatible birds during the acclimation period.
Therefore, control mortality in excess of 10% indicates
problems with some aspect of the test.
C. Test Conditions
1. Test Species
a. Selection
The mallard, Anas platyrhynchos L. , is the species to be
tested. The choice of mallard is based on a number of
factors. The mallard has a widespread distribution, not
only in the United States, but also in Eurasia. Such
distribution means that mallards may be exposed to toxic
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substances in the environment regardless of the location of
the toxic substance. As waterfowl, mallards may also be
exposed to toxic substances in the water, in sediments, and
on land. Because the mallard is the most abundant and
widely distributed duck in the northern hemisphere (Bellrose
1976), it is also suitable as a native test species for many
countries belonging to the Organization for Economic
Cooperation and Development (OECD).
Mallards have been one of the more frequently used
native bird species in reproduction tests with toxic
chemicals, but there are still only a few published
experimental studies with the species. DeWitt et al. (1963)
found that mallards were more susceptible to reproductive
impairment from several chemicals than were bobwhite or
pheasant. Heath et al. (1969) showed that mallards are
susceptible to eggshell thinning effects from DDE. The
Agency's confidential pesticide files contain additional
data confirming mallard sensitivity to eggshell thinning and
other reproductive impairment. When reproductive tests are
conducted on both gallinaceous birds and waterfowl, this
combination of species will often indicate impairment that
might not be found if only one species were tested.
The mallard is also amenable to testing in the
laboratory. Mallards can be bred in captivity and are
readily available from commercial sources so that testing of
this species will not deplete wild stocks. There is
sufficient information on the nutritional, habitat, and
behavioral characteristics of natural populations of
mallards in order to meet the basic nutritional and physical
requirements of the species in the laboratory.
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An advantage of reproduction testing with mallards or
bobwhite instead of other birds is that mallard and bobwhite
are typically the two species used in short-term toxicity
tests with birds. There is an advantage in using the same
species in reproductive tests that also can be used in
short-term or field tests. Mallards have been used for
short-term laboratory tests and actual or simulated field
tests from the beginnings of ecological effects testing
(DeWitt 1956) to the present (US EPA 1980b). Thus, the
choice of mallard as a test species facilitates comparisons
of the results from different kinds of tests.
In addition, mallards are generally considered to have a
positive economic value. Although the Agency is charged
with the protection of all species in the environment, the
choice of an economically valuable species for testing is
appropriate to the cost-benefit or risk-benefit analyses
upon which Agency decisions frequently are based. Finally,
there is as good a comparative data base for reproduction
tests with mallards as for any native bird.
If a test is to simulate toxicity to naturally occurring
populations of mallards, then it is important to use birds
that are phenotypically indistinguishable from wild birds.
Since many chemicals act upon specific enzymes, and enzymes
are based on the genetic code, the use of birds
genotypically similar to wild birds would be desirable.
However, the determination of phenotype is a simple
observational process, whereas genotypic determination is
impractical, if possible at all. In addition,
wild birds have a degree of heterogeneity that would not be
typical of any given genotype.
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The necessity for using healthy birds is obvious since
the test is designed to determine toxicity to typical
populations. It is admitted that not all birds occurring in
natural populations are healthy, but the majority of
survivors in natural environments are healthy. The health
of birds is also important in reducing the number of
variables that limit comparisons between tests. There are
several checks in this test guideline that help to ensure
that healthy birds are used. The use of previously untested
birds not selected for resistance and being from disease-
free flocks provides a basically healthy stock. Visual
observations select out abnormal or unhealthy birds from
that stock. A final check on health is based on the birds
ability to survive two weeks immediately preceding
exposure. The 3% maximum mortality during this period
allows for an occasional death that may occur during
acclimation and the time when unfamiliar birds in a pen are
becoming adapted to each other.
The choice of first year birds was made in this
guideline primarily because it facilitates comparisons with
existing data and data that may be generated according to
the US EPA (1978a) protocol for mallard. It also ensures
that birds will be the same age; whereas if no age were
specified, birds in the same test could be various ages.
Use of first year birds also reduces the cost of birds and
ensures that test birds will not have been used in previous
reproduction tests.
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2., Maintenance of Test Species
a. Acclimation
An acclimation period is necessary for birds to become
familiar with the test environment. Ideally, birds will be
maintained in test cages for several months, but this is
impractical for testing facilities that purchase adult
birds. It is also sometimes necessary to alter the
composition of birds in a pen because of excessive
aggression or other incompatibilities. The acclimation
period allows time for rearranging incompitible birds.
b. Diet
There are few data on the detailed nutritional
requirements for mallards. This subject is being
investigated under a current contract and is a proposed
research need. At the present time, a commercial duck
breeder ration is recommended. All known testing facilities
in this country use a commercial ration or a similar but
specially prepared ration made by a commercial company.
There are no known data to support an alternative diet for
mallards. Changes may be made in the future when additional
data become available. The recommendation of a commercial
duck starter ration is made for the same reasons.
Samples of treated diets will be analyzed to determine
the actual test concentrations used in the test.
3. Facilities
Only a few mallard reproduction tests have been reported
in the literature, although a number have been submitted to
the Agency and classified as confidential. As a result, it
is difficult to define optimal conditions and most
conditions have been recommended, rather than required.
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Heath et al. (1969, 1972b) and Heath and Spann (1973) used
outdoor pens with concrete floors for their mallard
reproduction studies. Heath and Spann (1973) specified the
size of pens as 10 X 20 feet; they housed 5 hens and 2
drakes in each pen. Heath, et al. (1969, 1972b) used the
same pens and housed 5 or 6 hens and 2 drakes in each pen.
Heinz (1976b) tested one drake and four hens in 5 x 10 ft.
pens, but in another study tested mallard pairs in one meter
square (10.7 square feet) pens (Heinz 1976a). Further
housing details were not mentioned.
US EPA (1978a, 1980b) and ASTM (1979a) have both
developed protocols for avian reproduction tests.
Rationales for selecting particular conditions were not
spelled out in either protocol. Reasons were apparently
based on experience with reproduction tests that have been
submitted to the Agency for pesticide registration and that
have been classified as confidential. These three protocols
have specified size only as being "adequate". All three
protocols recommend 21°C, 55% relative humidity, and
adequate ventilation. The same conditions have been
recommended in this guideline, since these protocols are the
most widely used and data developed from this guideline will
be comparable with data gathered using all three protocols.
Galvanized metal is the recommended construction
material. This material has been used widely in toxicity
tests (Heath et al. 1972a, Hill et al. 1975) and has been
recommended in ASTM (1979a) and US EPA (1978a) protocols for
reproduction tests. The ASTM (1979a) protocol suggests
stainless steel or perfluorocarbon plastics (e.g., Teflon)
as alternatives. As a relatively non-adsorbent material,
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perfluorocarbon plastics may be used to coat wood or other
materials that might be contaminated by chemicals if used
uncoated. Because of the tendency of wood to sorb
chemicals, uncoated wood may be not be used where it is
likely to become contaminated and come in contact with birds
in subsequent tests.
Pens should be cleaned and sanitized between tests.
Brushing and/or steam cleaning appear to be the most
appropriate since they do not involve the use of chemicals
that could affect subsequent tests. Detergents and bleach
have been used by Denver Wildlife Research Center (Tucker,
personal communication) and Patuxent Wildlife Research
Center (Heinz, personal communication). The use of chemical
disinfectants, such as quaternary ammonium compounds, should
be avoided because of possibility that these compounds can
leave toxic residues. However, the widely used cold
sterilization method with ethylene oxide is acceptable, if
needed for disease control. Pens should not be cleaned
during a test in order to minimize disturbance to the test
birds.
4. Environmental Conditions
a. Temperature (See Section II.C.3)
b. Humidity (See Section II.C.3)
D. Reporting
The information that is required to be reported in
section II.B.8.b is essential to a proper evaluation of the
test reults. These required items are needed (1) to
establish that the test was conducted according to
specifications, (2) to evaluate those conditions and
procedures that could affect the results of the test, and
(3) to supply the Agency with sufficient information to
25
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ES-14
August, 1982
conduct an independent analysis of statistics and
conclusions. The location of the raw data storage will
allow the Agency to find additional information that may
have been left out of the report or that may be needed for
enforcement purposes. The location is necessary because
some chemical companies request the testing facility to keep
these data, while other companies keep their own. The
information is needed in a detailed manner because this
avian reproduction guideline contains few rigid
requirements. Even when minimums or maximums are specified,
it is important to know how much the test may have exceeded
specifications, such as; if test birds were observed more
frequently than required, if the number of test
concentrations exceeded the three levels required, etc.
The information required in section II.B.S.b on Test
Substance Concentration to be available, but not included in
the test report, may be needed if there are serious concerns
about the results or validity of the test. This information
will not normally be needed and therefore is not required
routinely.
III. Economic Aspects
Three laboratories were surveyed to estimate costs for
performing the test outlined in this guideline. The
individual laboratories gave prices of $22,000, $12,650, and
$40,000. The "best estimate" based upon the survey was
$24,883. A cost estimate also was made by separating the
protocol into components and estimating the cost of each
component, including direct labor cost, overhead cost, other
direct cost, general and administrative cost, and fee. The
best estimated final cost, based upon this calculation
26
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ES-14
August, 1982
method, was $24,129, with an estimated range of $12,064 to
$36,193 based on +_ 50% of the best estimate. The calculated
estimate is similar to the best estimate based on the
survey. Marked differences in prices obtained from the
different laboratories may have resulted from a number of
factors, such as nature of the chemical, overhead rates,
outside consultants, automation, marketing strategies, and
other factors as outlined in a cost analysis report by
Enviro Control (1980). The cost estimates were made
assuming that the requirements of the Good Laboratory
Practice Standards, as specified in section (d) of the
Mallard reproduction guideline, are being satisfied.
In a cost analysis of subpart E pesticides guidelines,
laboratories were surveyed in 1978 and in 1980 to determine
the cost of testing (US EPA 1980a). The cited costs did not
differentiate between species, however, the unit cost for an
avian reproduction test was $24,000 in 1978 and $28,000 in
1980.
27
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SS-14
August, 1982
IV. References
Anderson DW, Hickey JJ. 1972. Eggshell changes in
certain North American birds, pp. 514-540 in
Proceedings of the XV International Ornithological
Congress. Voous KH, ed. E.J. Brill, Leiden, 745 pp.
1972.
ASTM 1979a. American Society for Testing and
Materials. Standard practice for conducting
reproduction studies with birds. Draft No. 1
20 pp.
ASTM 1979b.
Materials .
with birds.
American Society for Testing and
Standard practice for conducting LC
Draft No. 1. 13pp.
50
tes ts
Bellrose FC. 1976. Ducks, Geese, and Swans or North
America. Stackpole Books, Harrisburg, Pa. 543 pp.
DeWitt JB. 1956. Chronic toxicity to quail and
pheasants of some chlorinated insecticides. J. Ag. Food
Chem. 4(10): 863-866.
DeWitt JB, Crabtree DG, Finley RB, George JL. 1962.
Effects on wildlife, pp. 4-10 (-i-Tables) in USDI,
Effects of Pesticides on Fish and Wildlife: A Review
Investigations during 1960. Bureau Sport Fish and
Wildl. Circ. No. 143. 52 pp.
of
DeWitt JB, Stickel WH, Springer PF.
studies, Patuxent Wildlife Research
USDI, Pesticide - Wildlife studies:
and Wildlife Service Investigations
1962. Fish and Wildlife Serv. Circ.
Wildlife
pp. 74-96
1963.
Center.
A Review of Fish
during 1961 and
No. 167. 109 pp.
in
Enviro Control, Inc. 1980. Cost analysis methodology
and protocol estimates: ecotoxicity standards.
Rockville, MD: Enviro Control, Inc., Borriston
Laboratories, Inc.
Genelly RE, Rudd RL. 1956. Effects of DDT, Toxaphene,
and dieldrin on pheasant reproduction. Auk 73: 529-539
Greenwood RJ . 1975.
four mallard lines.
Reproduction and development of
Prairie Natur. 7(1): 9-16.
28
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ES-14
August, 1982
Heath RG, Spann JW. 1973. Reproduction and related
residues in birds fed Mirex. pp. 421-435 in Pesticides
and the Environment: A continuing contro~versy.
Symposia Specialists, North Miami. 1973.
Heath RG, Soann JW, Kreitzer JF. 1969. Marked DDE
impairment of mallard reproduction in controlled
studies. Nature 224: 47-48.
Heath RG, Spann JW, Hill EF, Kreitzer JF. 1972a.
Comparative dietary toxicities of pesticides to birds.
U.S. Fish and Wildlife Service, Spec. Rept. Wildl. No.
152. 57 pp.
Heath RG, Spann JW, Kreitzer JF, Vance C. 1972b.
Effects of polychlorinated biphenyls on birds, pp. 475-
485 in Proceedings of the XV International
Ornithological Congress. Voous KH, ed. E.J. Brill,
Leiden, 745 pp. 1972.
Heinz GH. 1976a. Methylmercury: second year feeding
effects on mallard reproduction and duckling behavior.
J. Wildl. Manage. 40(1): 82-90.
Heinz GH. 1976b. Behavior of mallard ducklings from
parents fed 3 ppm DDE. Bull. Env. Contain. Toxic. 16(6):
640-645.
Hill EF, Heath RG, Spann JW, Williams LD. 1975. Lethal
dietary toxicities of environmental pollutants to birds.
U.S. Fish and Wildl. Serv., Spec. Sci. Rept. Wildl. No.
191. 61 pp.
Holmes WN, Cavanaugh KP, Cronshaw J. 1978. The effects
of ingested petroleum on oviposition and some aspects of
reproduction in experimental colonies of mallard ducks
(Anas platyrhynchos). J. Reprod. Fertil. 54(2):335-348.
Kirkpatrick CM. 1955. Factors in photoperiodism of
bobwhite quail. Physiol. Zool. 28:255-264.
Kirkpatrick CM. 1959. Interrupted dark period: tests
for refractoriness in bobwhite quail hens. pp. 751-758
_in_ Withrow RB (ed) . Photoperiodism. Am. Assoc. Advan.
Sci. Publ. No. 55, Washington, D.C.
29
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ES-14
August, 1982
Ratcliffe DA. 1967. Decrease in eggshell weight in
certain birds of prey. Nature 215: 208-210.
Stickel LF, Heath RG. 1965. Wildlife studies-Patuxent
Wildlife Research Center, pp. 3-30 in USDI. Effects of
pesticides on fish and wildlife. Fish and Wildl. Serv.
Circ. No. 226. 77 pp.
Tucker RK, Leitzke JS. 1979. Comparative toxicology of
insecticides for vertebrate wildlife and fish.
Pharmac. Ther. 6: 167-220.
U.S. Environmental Protection Agency. 1975. Protocol
for determining lethal dietary concentration of
chemicals to birds. Federal Register, 40 CFR 162.82
(appendix): 26915. June 25, 1975.
U.S. Environmental Protection Agency. 1978a. Avian
reproduction. Federal Register, 40 CFR 163.71-4:
29729-29730. July 10, 1978.
U.S. Environmental Protection Agency. 1978b. Avian
dietary LC50. Federal Register, 40 CFR 163.71-2:29727-
29728. July 10, 1978.
U.S. Environmental Protection Agency. 1979. Toxic
substances control. Discussion of premanufacture
testing policy and technical issues; request for
comment. Federal Register 44:16240-16292. March 16,
1979.
U.S. Environmental Protection Agency. 1980a. Cost
analysis: Guidelines for registering pesticides in the
United States, Subpart E. Draft. May 1980.
U.S. Environmental Protection Agency. 1980b.
Guidelines for registering pesticides in the United
States. Subpart E. Hazard Evaluation: Wildlife and
Aquatic Organisms. Draft. November 3, 1980.
Wo If son A. 1964. Animal photoperiodism. pp 1-49 in
Giese AC ( ed ) . Photophys iology , Vol. II. Academic
Press, N.Y.
30
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ES-14
August, 1982
Personal Communications:
Gary Heinz (11/16/79)
Patuxent Wildlife Research Center
Laurel, MD.
Richard K. Tucker (11/8/79)
EPA, Office of Toxic Substances
Washington, D.C.
K.L. Stromberg (2/4/80)
Patuxent Wildlife Research Center
Laurel, MD.
31
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EG-18, OECD
August, 1982
DAPHNID CHRONIC TOXICITY TEST
OFFICE OF TOXIC SUBSTANCES
OFFICE OF PESTICIDES AND TOXIC SUBSTANCES
U.S. ENVIRONMENTAL PROTECTION AGENCY
WASHINGTON, D.C. 20460
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Office of Toxic Substances EE-18
Guideline for Testing of Chemicals August, 1982
DAPHNID CHRONIC TOXICITY TEST
(a) Purpose. The proposed Daphnia chronic toxicity
test standard is designed to assess the effects of test
substances on the survival and reproduction of Daphnia as a
representative freshwater invertebrate. The duration of the
test permits the daphnids to be exposed to a chemical from
shortly after birth until well into adulthood. The
organisms are exposed long enough to allow the adults to
produce several broods of progeny. Initiating exposure
shortly after birth allows an assessment of the possible
effects of the test chemical on such processes as
reproduction, maturation, fecundity and growth.
(b) Def initions . The following definitions apply to
this standard:
(1) "Acute lethal toxicity" is the lethal effect
produced on an organism within a short period of time of
exposure to a chemical.
(2) "Confidence limits" are the limits within which, at
some specified level of probability, the true value of a
result lies.
(3) "LC50" is the median lethal concentration, i.e.
that concentration of a chemical in air or water killing 50
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EG-18
January, 1982
percent of a test batch of organisms within a particular
period of exposure (which shall be stated).
(4) "Reference substance" is a chemical used to assess
the constancy of response of a given species of test
organisms to that chemical, usually by use of the acute
LC50. (It is assumed that any change in sensitivity to the
reference substance will indicate the existence of some
similar change in degree of sensitivity to other chemicals
whose toxicity is to be determined).
(5) "Static test" is a toxicity test with aquatic
organisms in which no flow of test solution occurs.
Solutions may remain unchanged throughout the duration of
the test.
(6) "Renewal test" is a test without continuous flow of
solution, but with occasional renewal of test solutions
after prolonged periods, e.g., 24 hours.
(7) "Flow-through test" is a toxicity test in which
water is renewed continuously in the test chambers, the test
chemical being transported with the water used to renew the
test medium.
(8) "Time-response curve" is the curve relating
cumulative percentage response of a test batch of organisms,
exposed to a single dose or single concentration of a
chemical, to a period of exposure.
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EG-13
August, 1982
(9) "Toxicity curve" is the curve produced from
toxicity tests when LC50 values are plotted against duration
of exposure. (This term is also used in aquatic toxicology,
but in a less precise sense, to describe the curve produced
when the median period of survival is plotted against test
concentrations ) .
(10) "Units" all concentrations are given in weight per
volume (e.g., in mg/liter).
( c) Test procedures — (1) Summary of the test. ( A)
Test chambers are filled with appropriate volumes of
dilution water. In the flow-through test the flow of
dilution water through each chamber is then adjusted to the
rate desired. The test substance is introduced into each
test chamber. The addition of test substance in the flow-
through system is done at a rate which is sufficient to
establish and maintain the desired concentration of test
substance in the test chamber.
(B) For the renewal test, the test is started within 30
minutes after the test substance has been added and
uniformly distributed in the test chambers. In the flow-
through test the test begins after the concentration of test
substance in each test chamber of the flow-through test
system reaches the prescribed level and remains stable. At
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BG-18
August, 1982
the initiation of the test, daphnids which have been
cultured or acclimated in accordance with the test design,
are randomly placed into the test chambers. Daphnids in the
test chambers are observed periodically during the test,
immobile adults and offspring produced are counted and
removed, and the findings are recorded. Dissolved oxygen
concentration, pH, temperature, the concentration of test
substance, and other water quality parameters are measured
at specified intervals in selected test chambers. Data are
collected during the test to determine any significant
differences (P _<_ 0.05) in immobilization and reproduction as
compared to the control.
(2) [Reserved]
(3) Range-finding test. (i) General A range-finding
test should be conducted to establish test solution
concentrations for the definitive test.
(ii) Introductory information for range-finding test.
(A) Prerequisites:
(±) Water solubility.
(_2_) Vapor pressure.
(B) Guidance information:
(JJ Structural formula.
(2) Purity of the substance.
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BG-18
August, 1982
(J_) Methods of analysis for the quantification of the
substance in water.
(_4_) Chemical stability in water and light.
(_5_) n-octanol/water partition coefficient.
(_6_) Results of a test on biodegradabili ty.
(C) Qualifying statement. For chemicals with low
solubility under test conditions, it may not be possible to
quantitatively determine the EC50.
(iii) Methods for range-finding test. (A) Definitions
and units:
(_!_) 24 hour EC50. The concentration (based upon
nominal concentration) calculated to have immobilized 50
percent of the daphnids by 24 hours exposure. (If another
definition is used, this shall be reported, together with
its reference).
(_2_) Immobilization. Those animals not able to swim for
15 seconds after gentle agitation of the test container are
considered to be immobile. (If another definition is used,
this shall be reported, together with its reference).
(B) Reference substances. In the course of the acute
immobilization phase a reference substance may occasionally
be tested for EC50 with the test compound as a means of
assuring that the laboratory test conditions are adequate
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BG-18
August, 1982
and have not changed significantly. An example of such a
useful reference substance is K^Cr^O^.
(C) Conditions for the validity of the range-finding
test:
(JJ The mortality in the controls should not exceed 10
percent at the end of the test.
(_2_) The oxygen concentration at the end of the test
shall be > 70 percent of the air saturation value at the
temperature used.
(_3_) Test Daphnia should not have been trapped at the
surface of the water, at least in the control.
(_4_) If conducted, the results with the reference
compound should be within the normal range for the
laboratory conducting the test.
(_5_) If the EC50 is not calculable due to an inadequate
number of intermediate response levels, it is acceptable to
merely report the highest concentra-tion causing complete
immobility, provided that the concentration factor between
d os es was _<_ 1.8.
(D) Performance of the range-finding test;
(_1_) Equipment which will come into contact with the
test solutions should be glass. This glassware should be
cleaned with solvents known to remove previously tested
chemicals.
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EC-IS
August, 1982
(_2_) Any water, either reconstituted or natural water,
can be used, provided that it will sustain growth and
reproduction of Daphnia without signs of stress.
(_3_) At least 20 animals should be used at each test
concentration, preferably divided into four batches of five
animals each.
(_4_) At least 2 ml of test solution should be provided
for each animal.
(_5.) The test temperature should be between 18 and 22°C,
and for each test it should be constant within +_ 0.5°C.
(_6_) A light-dark cycle is optional.
(1_) The concentrations should be formulated in a
geometric series, preferably without using any solvents. If
solvents, solubilizing agents, emulsifiers, etc., have to be
used, they should be commonly used adjuvants and not be
toxic in themselves at the levels used. Neither should they
have a synergistic or antagonistic effect on the toxicity of
the substance tested. In no case should the concentration
of an organic solvent exceed 0.1 ml/1.
(_8_) The test solution should be prepared before
intrododuction of the daphnids.
(_9_) The test solutions should not be aerated.
(10) The daphnids shall not be fed during the test.
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EG-18
August, 1982
(11) The highest concentration to be tested should not
exceed 1.0 g/1.
(12) Concentrations sufficient to lead to zero and 100
percent immobilization and the 24 hour EC50 should be tested
together with a control.
(13) The pH and the oxygen concentration of the blank
and all the test concentrations should be measured at the
beginning and the end of the test. The pH of the test
solutions should not be modified.
(14) Volatile compounds should be tested in completely
filled, closed containers, large enough to prevent lack of
oxygen.
(4) Definitive test. (i) General. The results of the
range-finding test are used to determine, with judgement,
the concentration levels to be used in the definitive
test. It is suggested that this reproduction test be
carried out using a geometrical concentration series of at
least five concentrations with an interval of at least 10,
starting at approximately the 24 hour EC50 concentration and
ending at 1/100 of the 24 hour EC50. If necessary, lower
concentrations are to be tested.
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EG-18
August, 1982
(ii) Introductory information for the definitive
test. (A) Prerequisites:
(JJ Water solubility.
(_2) Vapor pressure.
(_3_) Chemical stability in water and light.
(_4_) Results of a test on biodegradabili ty.
(_5_) 24 hour EC50 or the highest concentration producing
no immobility and the lowest concentration causing complete
immobility.
(B) Guidance information:
(I) Structural formula.
(_2_) Parity of the substance.
(_3_) _n-octanol/water partition coefficient.
(C) Recommendations. (_!_) Instead of a two week test
in which three batches of young should be born per female, a
test of three or four weeks may be preferred in order to
obtain a Tiore thorough judgement of the influence of the
test substance on mortality and reproduction. In this
period approximately six to nine batches of young should be
born per female.
(_2_) It is recommended that a statistical test (such as
an analysis of variance) be used to determine whether the
test replications can be analyzed together.
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EG-18
August, 1982
(iii) Criteria for a valid definitive test. (A)
Control mortality should not exceed 20 percent at the end of
the test.
(B) The oxygen concentration shall have been > 70
percent of the air saturation value throughout the test.
(C) The pH for the controls and for at least the most
concentrated solutions shall be known throughout the test.
The deviation from the initial value at the beginning of the
test should be _<_ 0.3 units.
(D) The first young should have been born in the
controls after a maximum of nine days.
(E) The average cumulative number of young per female
in the controls after three broods, should be > 20 at a
temperature of 20° +_ 0.5°C.
( iv) Definitive test procedures. (A) At least
40 animals should be used at each test concentration,
preferably divided into four batches of ten animals each.
The test concentrations are made up in a geometric series,
and if possible, without any solvents.
(B) Every test shall include controls consisting of the
same dilution water, conditions, procedures and daphnids
from the same population (culture container), except that
none of the chemical is added.
10
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EG-18
August, 1982
(C) The minimum duration of the test is 14 days, in
which at least three broods of the Fl generation shall have
appeared in the controls. If this is not the case, the test
shall be continued until the third brood in the control is
complete. If desired, the test can be continued for a total
period of three to four weeks, even if three broods are born
within three weeks.
(D) The live and dead daphnids of the "parental"
generation (P) are counted and the dead specimens removed.
This should preferably be carried out daily, but at least
every two days, e.g. Monday, Wednesday and Friday.
(E) The presence of eggs in the brood pouch, males or
epphipia shall be recorded. The condition and size of the
parent generation should be visually compared with the
controls.
(F) When the parental animals are about seven days old,
the first young daphnids emerge from the brood pouch. After
this, a new batch appears every two to three days. These
batches are called "broods" of the Fl generation.
(G) The newborn young of the Fl generation are counted
at least every two days (Monday, Wednesday and Friday) and
their estimated condition (based on visual examination) is
11
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BG-18
August, 1982
recorded. After counting and examination, the young are
poured away. The presence of eggs from which no young have
emerged (on the bottom of the test vessel) is recorded.
(5) [Reserved]
(6) Analytical measurements. (i) De ionized water
should be used in making stock solutions of the test
substance. Standard analytical methods should be used
whenever available in performing the analyses. The
analytical method used to measure the amount of test
substance in a sample shall be validated before beginning
the test by appropriate laboratory practices. An analytical
method should not be used if likely degradation products of
the test substance, such as hydrolysis and oxidation
products, give positive or negative interferences which
cannot be systematically identified and corrected
mathematically.
(ii) Samples of the test substance should be taken at
the beginning and during the test. The actual concentration
shall not drop below 80 percent of the nominal concen-
tration. Aeration of the test solutions is permissible,
unless this would cause the actual concentration of the test
substance to drop below 80 percent of the nominal
concentration.
12
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EG-18
August, 1982
(iii) The oxygen concentration in all test solutions
shall be checked once every 48 hours (if desired, every
Monday, Wednesday and Friday).
(iv) The pH of the controls and of at least the most
concentrated solution shall be checked before and after each
renewal in the renewal test and once every 48 hours in a
flow-through test.
( d ) Test conditions — (1) Test species — ( i )
Selection. Daphnia magna less than 24 hours old at the
beginning of the test, laboratory bred, free from known
diseases and with a known history (breeding method,
pretreatme nt) are used in this test. Other Daphnia species
may be used provided that the relevant re-production
parameters are comparable to those of Daphnia magna.
(ii) Feeding. The daphnids should be fed at least
daily during the definitive test. In the chronic daphnid
test, food (in any quantity) of any kind that meets the
criteria of reproduction for validity of the test, is
acceptable. Overloading of the test system with food should
be avoided in order to minimize sorption of the test
substance. Log-phase, unicellular green algae are generally
suitable.
13
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EG-18
August, 1982
(2) Facilities—(i) Apparatus. (A) Normal laboratory
apparatus and equipment should be used. Equipment which
will come into contact with the test solutions should
preferably be all glass. This glassware should be cleaned
with solvents known to remove previously tested chemicals.
(B) This reproduction test should not be carried out in
a static test system; either a renewal or flow-through
system shall be used. The renewal period should be guided
by the chemical analysis and (if applicable) the oxygen
level in the test solution. The solutions shall be renewed
at least once every 48 hours (if desired, on Monday,
Wednesday and Friday) .
(C) Volatile compounds should be tested in completely
filled closed containers, large enough to prevent the oxygen
concentrations from falling below 70 percent of the
saturation value. An almost-closed, flow-through system may
also be used. When more than 20 percent of the test
compound would be lost through volatility, the test should
be carried out either in a flow-through system or in an
enclosed container of sufficient size to ensure that the
oxygen level does not fall below 70 pecent of the saturation
value.
14
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EG-18
August, 1982
(ii) Cleaning. If the renewal scheme is used, the
glassware shall be emptied and food residues removed at
renewal. It is recommended that the glassware be rinsed
with de ionized water and kept as a coded series for the
following renewal. Each test unit therefore has two vessels
which are used alternately. If flow-through systems are
used, these should be cleaned twice a week.
(iii) Dilution water. Surface or ground water,
reconstituted water, or dechlorinated tap water is
acceptable as dilution water if daphnids will survive and
reproduce in it for the duration of the culturing,
acclimation, and testing periods without showing signs of
stress. The quality of the diluton water should be constant
and should meet the following specifications:
Substance Maximum Concentration
Particulate matter 20 mg/1
Total organic carbon or 2 mg/1
chemical oxygen demand 5 mg/1
Un-ionized ammonia 20 ug/1
Res idual chlorine 1 ug/1
Total organophosphorus pesticides 50 ng/1
Total organochlorine pesticides
plus polychlorinated biphenyls (PCBs) 50 ng/1
or organic chlorine 25 ng/1
15
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EG-18
August, 1982
(3) Test parameters — (i) Carriers . If solvents,
solubilizing agents, emuls if iers, etc., have to be used,
they should be commonly used adjuvants and should not
themselves be toxic at the concentrations used. They should
also not interact to alter the toxicity of the substance
under test. In no case should the concentration of an
organic solvent exceed 0.1 ml/1.
(ii) Dissolved oxygen. The oxygen concentration shall
be > 70 percent of the air saturation value throughout the
test.
(iii) Lighting. A light-dark cycle is necessary for
the definitive test; 8 hours darkness and 16 hours light are
recommended .
( iv) Loading . At least 40 ml of test solution should
be provided for each animal in the definitive test.
(v) Temperature. The test temperature should be
between 18 and 22°C, but for each test it should be
constant, wi thin jf 0 .5°C.
(e) Reporting (i) Test substance information:
(A) chemical designation.
(B) additional designations, e.g. trade name.
(C) empirical formula.
(D) manufacturer.
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EG-18
August, 1982
(E) batch number.
(F) degree of purity.
(G) date of sampling.
(H) water solubility.
(I) vapor pressure.
(J) biodegradability.
(K) chemical stability in water and daylight.
(L) _n«octanol/water partition coefficient.
(ii) General information:
(A) Source of Daphnia, any pretreatment, breeding
method (including source, kind and amount of food, feeding
frequency), species identification and method of
verif ication.
(B) Name and address of the testing laboratory, name of
the person responsible for carrying out the test (study
director).
(C) Name and address of sponsor.
(D) Dates of testing.
(E) Description of the test method or reference to the
method used.
(iii) Conditions of testing:
(A) Carriers and/or additives used and their
concentrations. If it is observed that the stability or
17
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0G-18
August, 1982
homogeneity of the test substance cannot be maintained, then
care should be taken in the interpretation of the results
and note made that these may not be reproducible.
(B) Dilution water: source and chemical and physical
characteristics including at least hardness, pH, Ca/Mg
ratio, Na/K ratio, alkalinity.
(C) Test temperature.
(D) Light quality, intensity and periodicity.
(E) All measurements of pH and oxygen level made during
the test, preferably in tabular form.
(F) Results and date of test performed with reference
compound if available.
(G) Description of test vessels: volume of solution,
number of test organisms per vessel, number of test vessels
per concentration, conditioning of the test vessels, the
introduction of the test substance in the dilution water.
(H) In case of renewal, the renewal procedure and
scheme. In case of flow-through, the test substance
delivery system, the flowrate, periodicity of cleaning and
technique used.
(I) If measured, the actual concentrations of the test
substance and the dates of measurement.
(J) Number and percentage of daphnids that showed any
18
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EG-18
August, 1982
adverse effect in the controls and in each treatment at each
observation period and a description of the nature of the
effects observed, e.g. immobilization, mortality in tabular
form.
(K) Description or reference to statistical procedures
applied.
(L) Any other effects differentiating organisms in
tests and controls.
(iv) Specific range-finding and definitive test
information: (A) For the 24 hour EC50 (acute
immobilization) phase also report:
(JJ The 24 hour EC50 perferably with 95 percent
confidence limits, either by computation or graphically, and
the method applied. The probit method is recommended.
(_2_) If possible, the slope of the concentration
response curve with its 95 percent confidence limits.
(_3^ The highest tested concentration producing no
immobile daphnids.
(_4_) The lowest tested concentration producing 100
percent immobile daphnids.
(_5_) Any other effect observed and the concentration at
which it occurred.
19
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EG-18
August, 1982
(B) For the reproduction phase also report:
(_!_) The EC50 ( immobilization) and LC50 values as far as
possible at 24 hours, 48 hours, 96 hours, 7 days, 14 days
and at the end of the test, preferably with 95 percent
confidence limits, either by computation or graphically, and
the method applied. For the determination, a probit method
should be used.
(_2_) The length of time for the appearance of the first
brood for each concentration
(_3_) The number of young alive in each test vessel on
given days when counts were made (the minimum requirement is
for counts at 48 hour intervals on Mondays, Wednesdays and
Fridays) .
(_4_) The number of dead young in each test vessel on
given days when counts were made.
(_5_) Source, kind and amount of food; feeding frequency.
(6_) If the recommended concentration scheme was
followed and no effectss on reproduction are detected, then
the results may be reported as being greater than the
highest concentration tested.
(_7_) For each of the observed effects a statistical
analysis of the homogeneity of replicate results for each
concentration should be made. If homogeneity is found, it
20
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EG-18
August, 1982
should be determined through an appropriate statistical
analysis, whether a significant difference exists between
the control and the test concentrations.
(_8_) The highest concentration tested at which no
significant difference is found compared to the controls
with respect to mortality, reproduction and other observed
effects.
(_9_) The lowest concentration tested with significant
difference compared to the controls.
(10) Any other parameter can be reported at the option
of the study director.
21
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EG-19, OECD
August, 1982
ALGAL ACUTE TOXICITY TEST
OFFICE OF TOXIC SUBSTANCES
OFFICE OF PESTICIDES AND TOXIC SUBSTANCES
U.S. ENVIRONMENTAL PROTECTION AGENCY
WASHINGTON, D.C. 20460
-------
Office of Toxic Substances EG-19
Guideline for Testing Chemicals August, 1982
FRESHWATER ALGAE ACUTE TOXICITY TEST
(a) Purpose. (1) A unicellular green alga is used as
a model system initially to estimate the concentration of a
chemical which could affect the primary production of
plants. Because regeneration times for unicellular algal
species are measured in hours, this relatively short test
can assess effects over several generations. Results allow
the assessment of effects on initial organisms from short
term exposures and give an indication of the effect on algal
populations .
(2) Many different protocols for algal tests are
available. This growth test is easy to perform and gives
reproducible results with the recommended species. This
Test Guideline can be adapted for other algal species. If
such an adaptation is used, a description of the method
should be provided with the test report.
(b) Def initions . The following definitions apply to
this guideline:
(1) "EC-X" means the experimentally derived chemical
concentration that is calculated to effect X percent of the
test criterion.
(2) "Growth rate" means an increase in biomass or cell
numbers of algae per unit time.
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EG-19
August, 1982
(3) "Inhibition" means any decrease in the growth rate
of the test algae compared to the control algae.
(4) "Limited water-soluble substances" means chemicals
which are soluble in water at less than 1,000 mg/1.
(5) "Readily water-soluble substances" means chemicals
which are soluble in water at a concentration equal to or
greater than 1,000 mg/1.
(c) Test procedures — (1) Summary of the test. (A)
The procedures for the preparation of the algal suspension,
the stock solution of the test chemical, and the test media,
are dependent on the solubility of the chemical and
modifications in the testing procedure may be necessary due
to the chemical's solubility in water. For chemicals with
low solubility under test conditions, it may not be possible
to quantitatively determine the EC-50.
(B) For purposes of the test, algae are grown in
Erlenmeyer flasks in an environmentally controlled growth
chamber. The test is started when 50 ml of algal suspension
(IxlO4 or 2xl04 cells/ml) and 50 ml of the appropriate test
chemical dilutions are placed in the flasks. Algal growth
is measured at 24 hour intervals for at least 96 hours. A
Coulter Counter, counting chamber, or other appropriate
instruments may be used to determine cell density. The data
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EG-19
August, 1982
are used to define the concentration-response curve and the
time-growth curve, and to calculate the EC-50 and the no-
effect level (EC-0) for the chemical.
(2) [Reserved]
(3) Range-finding test. A range-f inding test should be
conducted to establish test chemical concentrations for the
definitive test.
(4) Definitive test. (i) A definitive test is used to
determine time-growth and concentration-response curves, as
well as the EC-50 and no effect level (EC-0) of the test
chemical. The testing method will vary slightly depending
upon whether the chemical is readily water-soluble, of
limited solubility, or volatile.
(ii) Criteria for a valid definitive test. (A) Algae
in the control flasks should exhibit log phase growth within
43 hours of test initiation and should produce a standing
crop of at least 105 cells/ml in 96 hours.
(3) At 96 hours, one test concentration should show no
significant decrease in growth rate and one concentration
should show greater than a 50 percent decrease in growth
rate relative to the control.
(C) The pH of each test solution should be measured
before use and, if necessary, adjusted to 7.5 _+_ 0.2 using
HC1 or NaOH.
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August, 1982
(D) The test chemical concentration should be
determined before and after the test when practical.
(E) No more than 20 percent of the test chemical should
be lost by volatilization. If more is lost (or is likely to
be), the test should be conducted in closed flasks with a
resulting lower standing crop.
(iii) Test procedures dependent on solubility.--(A)
Readily water-soluble chemicals. When readily water-soluble
chemicals are tested, the following procedures are
recommended :
(_1) A stock solution of the readily water-soluble
chemical should be prepared with micropore-filtered (0.45 urn
pore size) medium. The concentration of the stock solution
should be twice as high as the highest concentration to be
used in the test. From this stock solution at least five
dilutions should be made. The dilutions should be in a
geometric series with a ratio of 10 or n/10 (where n =
number of dilutions). A minimum of five concentrations
should be used such that the highest concentration results
in at least 50 percent growth inhibition and the lowest
concentration shows no significant difference (a = 0.05)
from the control.
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EG-19
August, 1982
(2) The algal suspension for testing readily-soluble
chemicals should have a density of approximately 2x104
cells/ml of medium. The cell density should be quantified
prior to beginning the test.
(_3_) The test should be carried out in triplicate (i.e.
3 replicate flasks per concentration). The test begins by
transferring 50 ml of algal suspension to 250 ml Erlenmeyer
flasks. Then, in sequence of increasing concentrations, a
50 ml volume of each prepared dilution is added to the
appropriate flask. The control flasks receive 50 ml of
medium. The flasks are then gently shaken and placed in the
test chambers. The algal concentration in samples from each
flask is determined at intervals of at least 24, 48, 72 and
96 hours after the start of the test and the number of cells
or biomass (dry weight) per ml is calculated for each
s ample.
(_4_) A fluorimeter or spectrometer can be used to
calculate cell number or biomass, but will not provide
precise measurements at the start of the test and at 24
hours. If a Coulter Counter, or spectrophotometer, is used
to enumerate algae at the beginning of the test, 100 ml of
medium should be used to determine the background.
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33-19
August, 1982
(B) Limited water-soluble chemicals. When chemicals
with limited water-solubility are tested, the following
modifications of the above procedure are recommended:
(_1_) A stock solution for chemicals with limited water-
solubility should be prepared with a suitable organic
solvent. This stock solution should be 10^ times as
concentrated as the highest concentration to be tested and
the amount of solvent necessary to dissolve the chemical
should not exceed 0.1 ml/1 at the highest chemical
concentration used.
(2) An algal suspension for testing chemicals with
limited water-solubility should have a density of
approximately 10^ cells/ml. One hundred (100) ml of this
algal suspension (10^ cells/ml) are placed in each flask and
10 ul of the various dilutions of test chemical and solvent
are added. Ten (10) ul of solvent are added to the solvent
control flasks. Otherwise the test should be conducted as
described for readily-soluble chemicals.
(iv) Test procedures dependent on volatility. When
volatile chemicals are tested, the following modification of
the procedures in Sec( 4) (iii) (A) and (B) is recommended.
The test is performed as described above (depending on the
chemical's solubility in water) except that 250 ml conical
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August, 1982
flasks with ground glass stoppers should be used. These
flasks are; filled as described above with the algal
suspension and test chemical solution. For each measurement
interval the complete contents of the flasks should be
used. Therefore, it is necessary to use a sufficient number
of flasks to allow for this destructive sampling with time.
(5) Test results, (i) The results of the measurements
should be tabulated. Growth curves resulting from the
experiment should be drawn on semi-logarithmic paper (Figure
1).
BIOMASS
0.5N«o|—
0.2SN<«) —
0.125 N(«) —
0.00 N<»>
TIME
Figure 1. Theoretical example of a result of a toxicity test with algae. The growth curve
is plotted on a simple logarithmic scale. The figures give the concentrations of
the test compound in weight per volume. The broken lines show the
measurements. N(oo) is the maximum cell density that can be achieved.
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EG-19
August, 1982
(ii) The reported chemical effects on algae should
include both the EC-50 value and the no-effect level ( EC-
0). A number of methods have been described for determining
the EC50 and ECO. Two examples are provided.
(A) EC-50/ and EC-0 determined from the specific
maximum growth rate (u). The mean value of the three
replicate measurements and the 95 percent confidence limits
should be calculated for the 24, 48, 72, and 96 hour
sampling times, and plotted as in Figure 1. Two values are
selected from the log-linear portion of the curve and the
specific growth rate (u) is calculated according to the
following formula:
10910 Nl
u = N?
0.434 (t2-t1)
where:
N-^ = the lower cell number chosen in the log phase at t]_
N2 = the higher cell number chosen in the log phase at
t2
t = time in hours
u = maximum specific growth rate
0.434 = coefficient to convert log^g data to loge
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EG-19
August, 1982
The percent reduction of the growth rate compared to
the control can be calculated:
"% inhibition" = 1 - u(tox) x 100
u(b)
where:
u(tox) = the growth rate in the presence of the chemical
u(b) = the growth rate in the control
The calculated percentages ("% inhibition") are plotted
against the log concentration as shown in Figure 2.
100
% INHIBITION
1
50
LOG CONCENTRATION
Figure 2. Theoretical example of the relation between the logarithm of the
concentration of the chemical and the percentage inhibition.
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EG-19
August, 1982
The highest concentration showing no difference from the
control (a) and the EC50 (b) can then be read from this
graph.
(B) EC-50 determined from the mean relative growth
rate. (JL_) The mean relative growth rate (RGR) during log
phase growth for each culture can be calculated as follows:
RGR = I0*?] on? - logi nftT
t2 - tl
where:
n-^ = number of cells/ml at t^
T\2 = number of cells/ml at t2
t = time (hours)
(2) The RGR of the three control replicates can be
calculated and its 95 percent confidence limits
determined. Treatment RGR values that are greater than the
control upper 95 percent confidence limit indicate algal
stimulation and should be ignored. Those above the control
RGR but within the 95 percent confidence interval should be
assumed to equal the control mean. Treatment RGR values
that are less than the lower 95 percent confidence limit of
the control should be used in the following calculations:
The low treatment RGR values should be expressed as a
percentage of the control RGR. The percentage values
10
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BG-19
August, 1982
including the highest concentration of test compound at
which growth is 100 percent that of control (zero inhibition
and the EC--0) and the lowest concentration with a value of 0
percent (100 percent inhibition) should be included in an
appropriate statistical analysis to determine the EC-50
value and its 95 percent confidence limits.
(2_) Interpretation of results. Algal populations
rapidly regenerate themselves upon removal of stress;
consequently, concentrations that produce effects need
careful interpretation.
(6) [Reserved]
(d ) Test conditions — (1) Test Species — ( i)
Selection (A) It is recommended that the algae used be a
fast-growing species that is convenient for culturing and
testing. The following freshwater species are considered
sui table:
(_1) Selenastrum capricornuturn.
(_2^) Scenedesmus quadricauda.
(_3^ Chlorella vulgaris.
(B) Axenic cultures are recommended and are highly
desirable when testing biodegradable compounds. However,
pure monocultures of algae are required.
11
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EG-19
August, 1982
(ii) Stock culture. (A) The stock cultures are algal
cultures that are regularly transferred to fresh medium to
act as initial test material. Cultures that are not used
regularly should be streaked out on sloped agar tubes. The
tube cultures should be transferred to fresh medium at least
once every two months.
(B) The stock cultures should be grown in Erlenmeyer
flasks containing the appropriate medium (volume about 100
ml). When the algae are incubated at 20°C with continuous
illumination, a weekly transfer is recommended. An amount
of "old" culture is transferred with sterile pipettes into a
flask of fresh medium, for an approximate 100 fold
dilution. The growth rate of a species can be determined
from the growth curve. If this is known, it is possible to
estimate at what density the culture should be transferred
to new medium. This should be done before the culture
reaches the senescent phase.
( iii) Selection of test algae. (A) Algae used in a
test should be in an exponential growth phase, with a cell
density of at least 104 cells/ml. If it is not possible to
use the stock culture directly for testing (due to lack of
or excessive cell growth) it may be necessary to pre-culture
the algae prior to use.
12
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EG-19
August, 1982
(B) To pre-culture the algae, 2 liters of sterile
nutrient solution are added to a 3 liter Erlenmeyer flask.
Sufficient algal suspension from the stock culture is added
to give an algal concentration of approximately 104 cells/ml
(+_ 25 percent). The flasks should be continuously
illuminated with fluorescent light, and the temperature
maintained at 20°C. Cultures should be shaken by hand at
least once every day; this is particularly important for
non-motile species. The cell concentration in the culture
should be determined daily so that the desired concentration
may be obtained for testing.
(C) Other methods of culturing may also be used. Some
algae can be grown rapidly in shake cultures. Bubbling with
air containing additional carbon dioxide may also accelerate
growth. Furthermore, under the culturing conditions
described above, the pre-culture,for the algal species
recommended in this test, should be optimal for inoculation
of test flasks after four to five days. If the algal
cultures contain deformed or otherwise abnormal cells (e.g.
clumped, chlorotic), they should not be used for the test.
(2) Facilities — (i) Apparatus. (A) In order to avoid
contamination with bacteria and other algae, all stock
culture maintenance operations should be carried out under
13
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EG-19
August, 1982
sterile conditions. Possible contamination should be
checked by suitable methods.
(B) Testing and culturing of algae should be done in an
environmentally controlled cabinet or chamber capable of
maintaining a temperature of 20°C _+_ 2°C and continuous
illumination of approximately 300 to 400 uE/m2 sec.
(C) In addition to normal laboratory apparatus and
equipment for algal testing, a counting apparatus (e.g.
Coulter Counter, counting chamber, fluorometer,
spectrophotometer, colorimeter) to determine cell numbers is
also necessary.
(D) The following apparatus and equipment are necessary
for algal culturing:
(1_) Incubators or climate rooms capable of maintaining
temperature and light at the recommended levels.
(_2_) Filtering apparatus, accompanying membrane filters,
(0.45 urn) and 5 liter flask.
(_3_) Inoculation needle.
(_4_) Sterile graduated pipettes.
(_5_) pH-meter.
(_6_) Culture tubes (150 x 18 mm) with sponge and/or
metal caps.
14
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EG-19
August, 1982
(_7_) Erlenmeyer flasks, 200 ml, with ground glass
stoppers, and a 3 liter flask.
(ii) Dilution water. Redistilled water should be used
for preparation of stock, chemical, and nutrient media
solutions .
(3) Test parameters — (i) Carriers . If solvents, are
used, they should not themselves be toxic at the
concentrations used and should not affect algal growth. In
no case should the concentration of an organic solvent
exceed 0.1 ml/1 in the highest concentration.
(ii) Lighting. Algae should be kept under continuous,
uniform illumination of approximately 300 to 400 uE/m2
sec. The light source should be fluorescent lights.
(iii) Lo ad i ng . For readily water-soluble and volatile
chemicals, an algal suspension containing approximately 2 x
104 cells/ml should be used. For chemicals with limited
water solubility the algal density should be approximately
104 cells/ml.
(iv) Temperature. The temperature for culturing and
testing algae should be 20°C +_ 2°C.
15
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EG-19
August, 1982
(v) Nutrient media. The recommended media for adequate
growth of algae is shown in Table 1.
TABLE I MEDIUM FOR FRESHWATER ALGAE
Nutrient salts
Amount
K2HP04.3H20
CaCl2.2H20
Na2C03.10H20
Fe3-citrate
Citric acid monohydrate
Trace element solution
(see below)
Redistilled water made
up to
PH
0.5
0.33
0.052
0.035
0.054
0.006
0.006
1.0
9
g
g
g
g
g
g
ml
1.0 liter
7.7 + 0.3
Trace Element Solution:
H3B03
MnCl2.4H20
ZnCl2
CuS04.5H20
(NH4)6M07.4H20
Redistilled water made up to
2.90
1.81
0.11
0.08
0.018
1.0
g
g
g
g
g
liter
16
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EG-19
August, 1982
All nutrient solutions can be prepared as concentrated stock
solutions using the salts shown in Table 1 and stored in the
dark and cold. These solutions should be sterilized by
filtration or by autoclaving. The medium is prepared by
adding the correct amount of stock solutions, or the
nutrients salts directly, to sterile distilled water, to
give the final concentrations listed. For solid medium, 1.5
to 2 percent agar can be added. Other media may be
necessary if species other than those above are used.
(e) Reporting. In addition to a description of the
type of test and method, the report submitted to EPA should
include the following information.
(1) For the chemical tested: Manufacturer, empirical
formula, batch number and its degree of purity, chemical
characterization, (e.g. trade name), and physical
properties;.
(2) For the test organisms: Origin of innoculum,
laboratory culture and strain number, and method of
cultivation (including whether cultures aerated and/or
shaken) .
(3) For the test conditions: Date of beginning and end
of the test and its duration, temperature, light intensity
and light quality in the growth chamber, type of test flask
17
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EG-19
August, 1982
(and if closed or open), initial pH of test solution, what
carrier is used and how much, concentrations of test
chemical, and the counting method.
(4) For the results: A tabulation of cell number or
biomass per ml for each flask at each sampling period, the
plotted time-growth curves for each concentration and the
concentration-effect curve, the EC-50 value and the highest
concentration showing no statistical growth inhibition (EC-
0) and the statistical methods used to calculate them, and
other observed effects, e.g. algicidal vs. algistatic
effects, clumping or chlorosis of cells.
(5) For the laboratory performing the test: The name of
the person responsible for carrying out the test (study
director) as well as the name of the person carrying out the
test, the name and address of the testing laboratory, and
the date and signature of the person responsible for the
test.
18
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EG-20, OECD
August, 1982
FISH ACUTE TOXICITY TEST
OFFICE OF TOXIC SUBSTANCES
OFFICE OF PESTICIDES AND TOXIC SUBSTANCES
U.S. ENVIRONMENTAL PROTECTION AGENCY
WASHINGTON, D.C. 20460
-------
Office of Toxic Substances EG-20
Guideline for Testing Chemicals August, 1982
FISH ACUTE TOXICITY TEST
(a) Purpose. This guideline will be used in developing
data on the acute toxicity of chemical substances and mixtures
("chemicals") to fish subject to environmental effects test
regulations under the Toxic Substances Control Act (TSCA) (Pub.L.
94-469, 90 Stat. 2003, 15 U.S.C. 2601 et. seg.). The United
States Environmental Protection Agency (EPA) will use data from
these tests in assessing the hazard of a chemical to the
environment.
(b) Def initions . The definitions in section 3 of the
Toxic Substances Control Act (TSCA) and in Part 792—Good
Laboratory Pracice Standards apply to this test guideline. The
following definitions also apply:
(1) "Acute toxicity" is the discernible adverse effects
induced in an organism within a short period of time (days) of
exposure to a chemical. For aquatic animals this usually refers
to continuous exposure to the chemical in water for a period of
up to four days. The effects (lethal or sub-lethal) occurring
may usually be observed within the period of exposure with
aquatic organisms.
(2) "Acute lethal toxicity" is the lethal effect
produced on an organism within a short period of time of exposure
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EG-20
August, 1982
to a chemical.
(3) "Confidence limits" are the limits within which, at
some specified level of probability, the true value of a result
lies.
(4) "LC50" is the median lethal concentration, i.e.,
that concentration of a chemical in air or water killing 50
percent of a test batch of organisms within a particular period
of exposure (which should be stated).
(5) "Static test" is a toxicity test with aquatic
organisms in which no flow of test solution occurs. (Solutions
may remain unchanged throughout the duration of the test).
(6) "Semi-static test" is a test without flow of
solution, but with occasional batchwise renewal of test solutions
after prolonged periods (e.g., 24 hours).
(7) "Flow-through test" is a toxicity test in which
water is renewed constantly in the test chambers, the chemical
under test being transported with the water used to renew the
test medium.
(8) "Time-response curve" is the curve relating
cumulative percentage response of a test batch of organisms,
exposed to a single dose or single concentration of a chemical,
to a period of exposure.
(9) "Toxicity curve" is the curve produced from toxicity
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EG-20
August, 1982
tests when LC50 values are plotted against duration of
exposure. (This term is also used in aquatic toxicology, but in
a less precise sense, to describe the curve produced when the
median period of survival is plotted against test
concentrations.).
(10) "Units" all concentrations are given in weight per
volume (e.g., in mg/liter).
(c) Test procedures — (1) Summary of the test. (i) The
aqueous solubility and the vapor pressure of the test chemical
should be known prior to testing. The structural formula of the
test chemical, its purity, stability in water and light,
_n-octanol/water partition coeffecient, and pKa value should be
known. The results of a biodegradability test and the method of
analysis for the quantification of the chemical in water should
also be known.
(ii) The fish are exposed to a range of test substance
concentrations preferably for a period of up to 96 hours.
Mortalities are recorded at 24, 48, 72 and 96 hours and the
concentrations which kill 50 percent of the fish (LC50) are
determined where possible.
(iii) The maximum concentration tested producing no
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August, 1982
mortality and the minimum concentration tested producing total
mortality should be recorded.
(iv) For chemicals with limited solubility under the
test conditions, it may not be possible to determine an LC50.
(2) [Reserved]
(3) Range-finding test. It may be necessary to perform
a range-finding test prior to a definitive test. It provides
information about the range of concentrations to be used in the
definitive test.
(4) Definitive test. (i) Fish should be exposed to at
least five concentrations spaced by a constant factor not
exceeding 1.8. A control and solvent control, when appropriate,
should also be tested.
(ii) Stock solutions of the required strength are
prepared by dissolving the appropriate amount of the test
substance in the required volume of dilution water. The pH value
of the stock solution should be adjusted to the pH value of the
dilution water unless there are specific reasons not to do so.
The test should be carried out without adjustment of pH if there
is evidence of marked change in the pH of the solution, and it is
advised that the test be repeated with pH adjustment and the
results reported. This pH adjustment should be made in such a
way that the stock solution concentration is not changed to any
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August, 1982
significant extent and that no chemical reaction or physical
precipitation of the test compound is caused. NCI or NaOH should
be used to adjust the pH.
(iii) Stock solutions of substances of low aqueous
solubility may be prepared by ultrasonic dispersion or, if
necessary, by use of organic solvents, emulsifiers or dispersants
of low toxicity to fish. When such auxiliary substances are
used, the control fish should be exposed to the same
concentration of the auxiliary substance as that used in the
highest concentration of the test substance. The concentration
of such auxiliaries should not exceed 0.1 ml/1.
( iv) The chosen test concentrations are prepared by dilution
of the stock solution.
(v) For test to be valid, the following criteria apply:
(A) If it is observed that the stability or homogeneity of
the test substance cannot be maintained, then care should be
taken in the interpretation of the results and a note made that
these results may not be reproducible.
(B) The mortality in the controls should not exceed 10
percent at the end of the test.
(C) The dissolved oxygen concentration should have been >60
percent of air saturation throughout the test.
(D) There should be evidence that the concentration of the
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August, 1982
substance being tested has been satisfactorily maintained (e.g.,
within 80 percent of the nominal concentration) over the test
period.
(5) Test results. (i) The fish are inspected after 24, 48,
72 and 96 hours. Fish are considered dead if touching of the
caudal peduncle produces no reaction. Dead fish are removed when
observed, and mortalites are recorded. Observations after the
first three hours and six hours are desirable.
(ii) Records are kept of visible abnormalities (e.g., loss
of equilibrium, swimming behavior, respirtory function,
pigmentation, etc.).
(iii) The cumulative percentage mortality for each
recommended exposure period should be plotted against
concentration on logarithmic-probability paper. A line is then
fitted by eye to these points and the concentration corresponding
to the 50 percent response point is read off. This is the LC50
for the appropriate exposure period. Median lethal
concentrations also can be calculated using standard procedures
given in any of the references cited in section (f). Confidence
limits (p=0.95) for the calculated LC50 values can be determined
using the standard procedures. The LC50 value should be rounded
off to two significant figures.
(iv) Where the data obtained are inadequate for the use of
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August, 1982
standard methods of calculating the LC50 (because most of the
results are for either no deaths or total mortality, where a
dilution ratio of 1.8 has been used) then the highest
concentration causing no deaths and the lowest concentration
producing 100 percent deaths should be used to determine the LC50
(this being taken as being the geometric mean of these two
concentrations).
(6) [Reserved]
(d) Test conditions — (1) Test species — (i) Selection.
(A) One of several species may be used, the selection being at
the discretion of the testing laboratory. It is suggested that
the species used be selected on the basis of such important
practical criteria as: their ready availability throughout the
year, their ease of maintenance, their convenience for testing,
and any economic, biological or ecological factors which have
bearing. The fish should be in good health and free from any
apparent malformation. If other species fulfilling the the above
criteria are used, the test method should be adapted in such a
way as to provide suitable test conditions.
(B) Examples of fish recommended for testing and their size
are given in Table 1.
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EG-20
August, 1982
TABLE 1—RECOMMENDED SPECIES FOR ACUTE TESTING
Recommended species Recommended
total length
(cm)
Brachydanio rerio (Teleostei, 2.0 _+_ 1.0
Cyprinidae) (Hamilton-Buchanan)
Zebra-f ish
Pimephales promelas (Teleostei, 2.0 _+ 1.0
Cyprinidae) Fathead minnow
Cyprinus carpio (Teleostei, 3.0 _+_ 1.0
Cyprinidae) (Linne 1758)
Common carp
Oryzias latipes (Teleostei, 2.0 _+ 1.0
Poeciliidae) (Schlegel 1850)
Red killifish
Poecilis reticulata (Teleostei, 2.0 _+ 1.0
Poeciliidae) (Peters 1859)
Guppy
Lepomis macrochirus (Teleostei, 2.0 _+ 1.0
Centrarchidae) (Linnaeus 1758)
Bluegill
Salmo gairdneri (Teleostei, 5.0 _+ 1.0
Salmonidae) (Richardson 1836)
Rainbow trout
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BG-20
August, 1982
(ii) Collection or acquisition. The fish mentioned above
are easy to rear or are widely available throughout the year.
They are capable of being bred and cultivated either in fish
farms or in the laboratory under disease- and parasite-controlled
conditions so that the test animal will be healthy and of known
parentage.
(iii) Holding and acclimation. (A) Fish should be held for
at least 12 to 15 days before testing. All fish should be
maintained in water of the quality to be used in the test for at
least seven days before they are used.
(B) Coldwater fish should be held in tanks containing at
least 300 1 of water while warmwater fish should be held in tanks
containing at least 100 1.
(C) The temperature of the holding water should be the same
as that used for testing. The dissolved oxygen concentrations
should be maintained above 80% of the air saturation value. A 12
to 16 hour photoperiod should be used.
(D) All fish should be fed three times per week or daily
until 24 hours before the test is started.
(E) A batch of fish is acceptable for testing if the
percentage mortality over the seven day period prior to testing
is less than five. If the mortality is between 5 and 10 percent
acclimation should continue for seven additional days. If the
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EG-20
August, 1982
mortality is greater than 10 percent, the entire batch of fish
should be rejected.
(2) Test facilities — (i) Apparatus . An oxygen meter,
equipment for determination of water hardness, adequate appartus
for temperature control, test tanks made of chemically inert
materials and other normal laboratory equipment are needed.
(ii) Dilution water. (A) Drinking water (dechlorinated if
necessary), good quality natural water, or reconstituted water,
with a total hardness of between 50 and 250 mg/1 (as CaCO)^ and
with a pH of 6.0 - 8.5 are preferred.
(B) Reconstituted water should be prepared from deionized
water or distilled water with a conductivity _<_ 10 Scm~ . One
hundred liters of reconstituted water can be prepared by adding
2.5 1 of the following solutions to a tank and bringing the
solution to volume with deionized water:
11.76g CaCl2 ' 2H20/1
4.93g MgS04 • 7H20/1
2.59g NaHC03/l
2.59g KC1/1
The sum of the calcium and magnesium ions in this solution is 2.5
mmol/1. The proportion of Ca:Mg-ions is 4.13 and of Na:K-ions is
10:1. The acid capacity of this solution is 0.8 mmol/1.
(C) The dilution water should be aerated until oxygen
10
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EG-20
August, 1982
saturation is achieved and then stored for about two days without
further aeration before use.
(3) Test parameters. (i) Constant conditions should be
maintained as far as possible throughout the test and, if
necessary, semi-static or flow-through procedures should be used.
(ii) The preparation and storage of the test material, the
holding of the fish, and all operations and tests should be
carried out in an environment free from harmful concentrations of
dust, vapors, and gases and in such a way as to avoid cross-
contamination. Any disturbances that may change the behaviour of
the fish should be avoided.
(iii) The following parameters are important:
(A) Dissolved oxygen. The dissolved oxygen concentrations
should be at least 60 percent of the air saturation value.
(B) Light. A 12 to 16 hour photoperiod should be used.
(C) Loading. A maximum loading of 1.0 g/1 for static and
semi-static tests is recommended; for flow-through systems a
higher loading can be acceptable.
(D) Temperature. Test temperatures of 15 _+ 2°C for rainbow
trout and 22 +_ 2°C for carp are recommended. The other
recommended species should be tested at 23 _+_ 2°C. The
temperature should be maintained within +_ 1°C of the selected
test temperature throughout the test period.
11
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EG-20
August, 1982
(E) Feeding. The fish should not be fed during the test.
(e) Reporting . (1) The sponsor should submit to the EPA
all data developed by the test that are suggestive or predictive
of toxicity.
(2) In addition to the reporting requirements prescribed in
Part 792--Good Laboratory Practice Standards the reported test
data should include the following:
(i) Details of the test procedures used (e.g. static, semi-
static, flow-through, aerated, etc.).
(ii) Information about the test organism (scientific name,
strain, supplier, any pretreatment, etc).
(iii) The concentrations tested.
(iv) The number of fish in each test chamber and the loading
rate.
(v) The methods of preparation of stock and test solutions.
(vi) The dissolved oxygen concentrations, pH values,
temperature, total hardness of the test solutions measured each
24 hours and any other available information on water quality.
(vii) Any available information on the concentrations of the
test chemical in the test solutions.
(viii) The maximum concentration causing no mortality within
the period of the test.
(ix) The minimum concentration causing 100 percent mortality
12
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EG-20
August, 1982
within the period of the test.
(x) The cumulative mortality in each concentration according
to the recommended observation times.
(xi) The LC50 values (based on nominal concentrations) at
each of the recommended observation times (with 95 percent
confidence limits, if possible).
(xii) A graph of the concentration-mortality curve at the
end of the test.
(xiii) The statistical procedures used for determining the
LC50 values.
(xiv) The mortality of the control animals.
(xv) Any incidents in the course of the test which might
have influenced the results.
(xvi) Any abnormal responses of the fish.
(xvii) A statement that the test was carried out in
agreement with the prescriptions of the Test Guideline given
above (otherwise a description of any deviations occuring).
(f) References.
(1) APHA. 1975. American Public Health Association,
American Water Works Association, Water Pollution Control
Federation. Standard methods for the examination of water and
wastewater, 14th ed. New York: American Public Health
Association.
13
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EG-20
August, 1982
(2) Committee on Methods for Toxicity Tests with Aquatic
Organisms. 1975. Methods for acute toxicity tests with fish,
macroinvertebrates and amphibians. Corvallis, Oregon: U.S.
Environmental Protection Agency. EPA-660/3-75-009.
(3) Finney AJ. 1978. Statistical methods in biological
assay. Weycombe: U.K. Griffin Ltd.
(4) Litchfield JT, Wilcoxon F. 1947. A simplified method
of evaluating dose-effect experiments. J. Pharm. Exp. Ther.
96: 99-1113.
(5) Peltier W. 1978. Methods for measuring the acute
toxicity of effluents to aquatic organisms. Cincinnati, Ohio:
U.S. Environmental Protection Agency. EPA-600/4-78-012.
(6) Sprague JB. 1969. Measurement of pollutant toxicity to
fish. I: Bioassay Methods for Acute Toxicity. Water Research
3: 794-821.
(7) Stephan CE. 1977. Methods for calculating an LC50.
In: Mayer FL, Hamelink JL. eds. Aquatic Toxicology and Hazard
Evaluation. ASTM STP 634. American Society for Testing and
Materials, pp. 65-84.
(8) Tabata K. 1972. Quality control of Japanese rice fish
for TLm-test. Water and Effluent 14: 1297-1303.
14
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EG-21, OECD
August, 1982
FISH BIOCONCENTRATION TEST
OFFICE OF TOXIC SUBSTANCES
OFFICE OF PESTICIDES AND TOXIC SUBSTANCES
U.S. ENVIRONMENTAL PROTECTION AGENCY
WASHINGTON, D.C. 20460
-------
Office of Toxic Substances EG-21
Guideline for Testing Chemicals August, 1982
FISH BIOCONCENTRATION TEST
(a) Purpose. This guideline is to be used for
assessing the propensity of chemical substances to
b ioconcentrate in fish. This guideline describes a
bioconcentration test procedure for the continuous exposure
of fish to a test substance in a flow-through system. The
United States Environmental Protection Agency (EPA) will use
data from this test in assessing the hazard a chemical may
present to the environment.
(b) Eief initions . The definitions in section 3 of the
Toxic Substances Control Act (TSCA) and in Part 792—Good
Laboratory Practice Standards are applicable to this test
guideline. The following definitions also apply:
(1) "Bioconcentration" is the increase in concentration
of test material in or on test organisms (or specified
tissues thereof) relative to the concentration of test
material in the ambient water.
(2) "Bioconcentration factor (BCF)" is the ratio of the
test substance concentration in the test fish (Cg) to the
concentration in the test water (C ) at steady-state.
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EG-21
August, 1982
(3) "Depuration or clearance or elimination" is the
process of losing test material from the test organisms.
(4) "Depuration rate constant (k2) ^s tne
mathematically determined value that is used to define the
depuration of test material from previously exposed test
animals when placed in untreated dilution water, usually
reported in units per hour.
(5) "Steady-state or apparent plateau" is a condition
in which the amount of test material being taken up and
depurated is equal at a given water concentration.
(6) "Uptake (u)" is the process of sorbing test
material into and/or onto the test organisms.
(7) "Uptake phase" is the time during the test when
test organisms are being exposed to the test material.
(8) "Uptake rate constant (k]_)" is the mathematically
determined value that is used to define the uptake of test
material by exposed test organisms, usually reported in
units of li ters/gram/hour .
( c ) Test procedures — (1) Summary of the test. ( i)
The test compounds' water solubility, n-octanol/water
partition coefficient and stability in water (hydrolysis,
photolysis and microbial degradation) should be known prior
to testing. The 24 and 96 hour LCSO's for the fish species
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EG-21
August, 1982
to be used in the study should also be known. These data
should be based on test substance concentrations measured
during a flow-through acute toxicity test.
(ii) The procedure proposed is applicable to organic
chemicals that are not readily degradable in a microbial
degradation test, relatively stable in the aquatic
environment and soluble in water at <1 mg/1.
(iii) Before any biological experiments are carried
out, the analytical method for the particular substance
should be tested. It should be shown experimentally on both
water and organisms that the recovery as well as the
reproducibility are satisfactory. Blank samples (of water,
solvents, etc.) should regularly be analyzed to ensure that
no contamination occurs. The detection level should be
determined and no quantification should be based on signals
which are less than 2.5 times the instrument noise.
Organisms and water samples should be removed in such a way
that no contamination or losses by adsorption occur.
(iv) This Guideline describes a procedure for
characterizing the bioconcentration potential of chemicals
in aquatic biota. Parameters used to characterize the
bioconcentration potential include the uptake rate constant
(k]_), the depuration rate constant (k2)' and the steady-
state bioconcentration factor, BCF (k-L/k2).
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BG-21
August, 1982
(v) Each of three separate groups of test organisms of
the same species is exposed to a different concentration of
the test material in water; 0, x, and lOx mg/1, where x is
defined by analytical and toxicological boundaries. The
duration of the uptake phase (3 hrs to 30 days) and
depuration phase (6 hrs to 60 days) varies according to the
time required to reach the desirable percent of steady-state
which is roughly estimated before the test starts. During
both phases of the test, organisms and water are
periodically removed from the test chambers and analyzed for
the test material.
(vi) The uptake rate constant, depuration rate
contant(s), bioconcentration factor, and their confidence
limits are calculated from the model that best describes the
measured concentrations of test material in the organisms
and water at any point in time.
(2) [Reserved]
(3) [Reserved]
(4) Definitive test—(i) Test solution preparation.
The test material should be added to the dilution water with
minimal use of solvents or other carriers. Several systems
adaptable to flow-through tests have been described for
saturation of water with relatively insoluble test materials
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EG-21
August, 1982
(Chadwick and Kugeragi 1968, Borthwick et al. 1977, Veith
and Comstock 1975). Acetone, dime thylf ormamide , ethanol,
methanol, and triethylene glycol are the solvents
recommended for use in preparing stock solutions. The
concentration of solvent in any test solution should not
exceed 0.1 ml/liter in flow-through tests.
(ii) Exposure concentrations. Test fish should be
exposed to two or more concentrations of test material in
water under flow-through conditions. As a guidance, the
highest concentration should be less than one-tenth of the
threshold or incipient LC50 for the test species and at
least 10 times higher than the detection limit in water and,
if possible, each exposure concentration should differ from
another by a factor of ten.
(iii) Test duration—(A) Estimation of the uptake
phase. (_1) As a guideline, the statistically optimum
duration of the uptake phase (u) is near the midpoint of an
uptake curve plotted on semi-log paper, or u = 1.6/k2' ^ut
not more than 3.0/k2, which is equivalent to 95 percent of
steady-state (Reilly et al. 1977). A ore-test estimate of
^2 maY be obtained from:
(_i_) A test with the same compound and a different
species.
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EG-21
August, 1982
( ii) A test with a similar material.
( i i i ) The results of a preliminary range-finding test.
(iii) Water solubility data.
log k2 = o.43 log(s) - 2.11
where (s) is the aqueous solubility in ppm.
( iv ) _n-octanol/ water partition coefficient data,
log k2 = -0.414 log (PQW) + 0.122
where (PQW) is the n-octanol/water partition
coeff icient.
(_2_) The duration of the uptake phase (u) for a test
material with log Pow = 3 would be:
log k2 = 0.414 (3) + 0.122 = -1.12
,k2 = 0.0759
u = 1.6/0.0759 = 21 hours
Similarly, for a test material having a log Kow = 6, the
duration of the uptake phase (u) would be:
log k2 = -0.414 (6) + 0.122 = -2.362
k2 = 0.0043
u = 1.6/0.0043 = 372 hours (16 days)
(B) Estimation of the depuration phase. Two times u is
usually sufficient time for about 95 percent removal of the
body burden (tj/2 = 0.69/k2), but several biological or
analytical factors may suggest equally acceptable
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EG-21
August, 1982
guidelines. Some compounds follow more complex
uptake/depuration behavior than a simple two compartment (Cw
and Cf), two parameter (k]_ and k2) model. For these
compounds, longer depuration periods are advisable. On the
other hand, the depuration time will most likely be
restricted by the lower limit of analytical detection for
fish.
(iv) Sampling schedule. (A) As a guideline, no fewer
than four uptake sampling times and five depuration sampling
times should be spaced throughout the duration of the
experiment, according to the following fractions of the total
time (Tt): first at 0.0278 Tt, second at 0.0556 Tt, third
at 0.1111 Tt, fourth at 0.2222 tfc, fifth at 0.3333 Tt (this
is the optimum change-over time), sixth at 0.5000 Tt,
seventh at 0.6667 Tt, eighth at 0.8333 Tt, and ninth at
1.000 Tfc. Table I contains examples of acceptable sampling
schedules for bioconcentration tests with test materials
with a log PQW = 3.0 and Log Pow = 6.0.
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83-21
August, 1982
TABLE I—ACCEPTABLE SAMPLIN3 SCHEDULES FOR BICCONCENTRATION TESTS
Test
Phase
Uptake
Depuration
Terminate
Action log
Start test
Add fish
Trans f er
fish to
untreated
water
test
Sampling^
Pow = 3
Hours
-1
0
0.5
1.0
1.5
2
4
7
13
20
24
30
40
50
60
Times
Log Pow = 6
Days
-2
-1
0
0.5
1.0
1.4
2.8
5.7
11
17
20
26
34
43
51
No. water
samples
1
2
2
2
2
2
2
2
2
2
2
2
2
2
2
28
No. fish
samples
4
4
4
4
6
4
4
4
6
40
^ Samples taken after a minimum of 3 tank volumes have been delivered.
(v) Sampling procedures. (A) It is advisable to
analyze both water and organism samples as soon as possible
after they have been collected to prevent degradation or
loss of test material and to determine approximate uptake
and depuration rate constants as the test proceeds. If
samples cannot be analyzed immediately, it is sometimes
appropriate to extract the test material into a solvent,
rendering it inert or easier to store until it can be
analyzed .
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EG-21
August, 1982
(B) Water samples should be obtained by siphoning
through glass tubing from the most central point in the test
tank. The sample vessel and siphon should be rinsed with
the test solution before collecting the sample.
(C) Water samples are best collected directly into
glass vessels of appropriate volume from which the test
material can be extracted or analyzed. These vessels might
include separatory funnels in the case of organic compounds,
or scintillation vials for radioactive test materials.
(D) If significant amounts of particulate matter are
present in the water sampled, a second sample should be
taken and analyzed after centrifuging to determine whether
test material was adsorbed on the particulate matter rather
than dissolved.
(E) Water samples containing highly persistent test
materials can be stored frozen in plastic containers for
later analysis. Care should be exercised to avoid use of
containers which could sorb or contaminate samples. With
most organic test materials, and especially those tending to
degrade easily, a better practice is to extract them from
the water and store them under refrigeration in solvent in
tightly sealed glass vials.
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EG-21
August, 1982
(F) When removing test organisms for analysis, they
should be netted or trapped in a random manner with as
little disturbance as possible. If two or more test
material concentrations are present, separate nets should be
used for each concentration. Organisms should be rinsed
with dilution water if accompanied by extraneous matter,
blotted dry, and killed by pithing the brain with a
dissecting needle or by severing the spinal cord above the
opercular region with scissors. They should then be
individually weighed and a record made to permit association
of the weight with the sample.
(G) Fish may be analyzed as whole fish or as portions,
e.g., edible portion (muscle), viscera, remaining carcass,
etc. Specific organs may also be analyzed if sufficient
biomass is available. If results based on body portions are
desired, after the fish is killed it should be eviscerated,
taking care not to puncture any parts of the visceral
portion which could leak body fluids and possibly cause
contamination of the remaining portions. The edible portion
or muscle may be removed with a scalpel, blotted dry, and
weighed before storing or analysis. The remaining carcass
should be weighed before being stored or analyzed. It is
necessary to record data for each portion for each
10
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August, 1982
individual fish so that whole body data can be reconstructed
based on the sum of the parts.
(H) After weighing, the sample is usually ground or
homogenized to promote extraction of test material or to
enhance solution of the tissue. Procedures for grinding,
extraction, separation of impurities, determination of lipid
content, etc., are described in the U.S. Food and Drug
Administration's Pesticide Analytical Manual (1975) or the
U.S. Environmental Protection Agency's Manual of Analytical
Methods for the Analysis of Pesticide Residues in Human and
Environmental Samples (1974).
(I) When determining the bioconcentration of test
materials which concentrate in lipids, it is often desirable
to determine the percent of the total tissue weight made up
by lipids. Results between samples are frequently less
variable when based on lipid weight rather than on total
weight (Reinert 1970).
(J) Organism samples can be wrapped in acetone-rinsed
foil, placed in glass jars and frozen if they are not to be
analyzed immediately.
(5) Test results. (i) Most bioconcentration data can
reasonably be described with a simple two-compartment/two-
parameter model as shown by a straight line depuration
11
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August, 1982
profile plotted on semi-log paper. If the depuration
profile does not appear to be a straight line, then more
complex models can be employed (Blau et al. 1975). Typical
variations from the simple model include a third parameter
to describe the rate of metabolism of the parent compound or
two additional parameters to describe redistribution of the
parent compound within the body of the fish. If the best
model is in question, it may be worthwhile to estimate
parameters for the models in question and to compare the
likelihood index of each model according to statistical
tests (Blau et al. 1975).
(ii) Graph paper method for depuration rate constant.
Plot each concentration of the test material found in fish
at each sampling time on semi-log paper. The slope of that
line is K:
tcfi
10
*2 " * 1
* t-<2
kj units • t~l
[t—1
12
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August, 1982
(iii) Graph paper method for uptake rate constant.
Given ^21 calculate k^ as follows:
k . . __5«_*2
j^ _ __^ .• •.«•
Cw (1-e 2 )
The value of Cf is read from the smooth uptake/depuration
curve near the uptake raid-point on serai-log paper.
(iv) Computer method for calculating uptake and
depuration rate constant. The preferred means for obtaining
the bioconcentration factor and k^ and k2 rate constants is
to use nonlinear parameter estimation methods on a digital
computer. Two such programs are BIOFAC (Dow) and NONLIN
(Proctor and Gamble). These programs find values for k^ and
k2 given a set of sequential time concentration data and the
model:
-k,t
r •
S*
-k2tc t_ <. t
13
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EG-21
August, 1982
This approach provides standard deviation estimates of
k-^ and k^/ and BIOFAC statistically weights the analytical
and biological variation of the fish concentration data.
These and other non-linear parameter estimation programs are
readily available for most computers accepting the Fortran
IV language or can be made available from a time-sharing
service bureau; they are currently being used by many
bioconcentration testing laboratories.
(v) Validity of the test results. (A) Scientific
judgement rather than rigid criteria should be exercised in
accepting or rejecting bioconcentration test results.
(3) Calculated BCF values based on an octanol/water
partition coefficient have a very wide confidence margin
(greater than jf 100 percent), but the quality of the value
may be better (narrower confidence margin) than an
experimental value from a poorly designed study. Generally,
the confidence margins for well designed studies approach _+
20 percent. Acceptable bioconcentration data should be
reported with confidence margins.
(C) Other criteria for judging the quality of
bioconcentration data include the following guidelines:
(_1_) Percent mortality or adverse effect in control or
treated organisms (suggested guideline, 10 percent).
14
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August, 1982
(2) Percent effect of dose on uptake/depuration rate
constants (suggested guideline 20 percent).
(_3_) Percent variation in Cw (suggested guideline, 20
percent) except for the initial dip that may approach 50
percent during the first few days of exposure.
(_4_) Temperature and dissolved oxygen should not vary
nore than _+_ 1°C and +_ 3 mg/liter.
(_5_) The importance of actually visualizing an apparent
plateau has been a subject of recent debate. It is
suggested that 80 per-cent of steady-state (^-1/^2^ in anv
tissue with a confidence margin of +_ 20 percent is more than
sufficient to estimate high quality rate constants for
compounds with BCF <10,000. For compounds with BCF >10,000
it may be desirable and acceptable to terminate the uptake
phase after a few days not to exceed 28 days even though <
80 percent: of steady-state was reached.
(_6_) A clearly defined uptake/depuration profile is an
indicator of high quality bioconcentration data.
(6) Analytical measurements (i) Prior to analyzing
fish or water for the test substance, control samples should
be spiked with several different concentrations of the test
substance and then analyzed. Final values of GW and Cf
should be corrected for recoveries and background.
15
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EG-21
August, 1982
(ii) Analytical detection limits of test substance in
both fish and water should be determined before the
bioconcentration test begins and should be documented in the
protocol. As a guideline/ the limit of detection may be
defined as a signal 2.5 times higher than the background
noise level.
( iii) If possible, results reported as "not detected at
the limit of detection" should be minimized by pre-test
method development and experimental design. These results
cannot be used for rate constant calculations. The units Cw
and Cg should both be expressed either as ppm or ppb.
(d) Test conditions — Test species—•(!) Selection.
(i) The procedures regarding selection of which species to
test, their source, handling, holding, disease treatment,
acclimation, and quality assurance prior to and during
testing should be those given in Committee on Methods for
Toxicity Tests with Aquatic Organisms (1975).
(ii) The freshwater fish species used most frequently
in bioconcentration tests have been rainbow trout, bluegill,
and fathead minnows. The most commonly used marine fish
have been spot, sheepshead minnows, silvers ides, shiner
perch, English sole, staghorn sculpin and 3-spine
sticklebacks. These species are more readily available than
most others and can be obtained in convenient sizes.
16
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August, 1982
(2) Facilities — (i) Construction materials.
Construction materials and commercially purchased equipment
that may contact any water into which test organisms are
placed should not contain any substances that can be leached
or dissolved by the water. Glass, $316 stainless steel, and
pertluorocarbon plastics should be used whenever possible to
minimize leaching, dissolution, and sorption. Some will be
more suitable than others for use with specific test
materials. Unplasticized plastics, cast iron, and concrete
can be used for holding and acclimation tanks and in the
water supply system. Rubber, copper, brass, galvanized
metal and lead should not come into contact with dilution
water, stock solutions, effluent samples or test solutions.
(ii) Toxicant delivery system. (A) One of several
toxicant delivery systems can be used successfully,
including the proportional diluter (Lemke et al. 1977).
Diluters are accurate over extended periods of time, are
relatively trouble-free, and have fail-safe provisions.
However, proportional diluters often require that
laboratories have more than 8 feet of headroom. A small
chamber to promote mixing of test material-bearing and
dilution water should be used between the diluter and test
chambers for each concentration. Design alterations, such
17
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EG-21
August, 1982
as modification to deliver duplicates of only two or three
concentrations, are easy to make (Jarvinen et al. 1977).
(B) Pump systems are relatively simple to understand
and use, require little space, and can be quite accurate.
Some investigators have found metering pumps to maintain
less variable test concentrations than piston operated
pumps.
(C) The performance of the toxicant delivery system
should be checked before and during each test. This should
include determination of the flow rate through each test
chamber and measurement of either the concentration of
toxicant in each test chamber or the volumes delivered by
each portion of the delivery system. The general operation
of the toxicant delivery system should be checked daily
during the test.
(D) The flow rate through the test chambers should be
at least five volume additions per 24 hours, but should take
into account the size of the test chamber, the size of the
test organisms and the loading. It is usually desirable to
construct the metering system so that it can provide at
least ten volume additions per 24 hours. The flow rates
through the test chambers should not vary by more than 20
percent from any one test chamber to any other or from one
time to another within a test.
18
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EG-21
August, 1982
(iii) Test chambers. Each of the height and width
dimensions of the test chamber should be at least 1.5 times
the largest horizontal dimension of the test organism. A
minimum volume of one liter per fish is crowded but
satisfactory for fish up to 15 g; large volumes to fish
weight ratios are preferred to minimize the initial Cw dip
and to help maintain the dissolved oxygen concentration.
(iv) Cleaning. Metering systems, test chambers, and
equipment used to prepare and store dilution water, stock
solutions, and test solutions should be cleaned before
use. New equipment should be washed with detergent and
rinsed with water, pesticide-free acetone, water, acid (such
as 5 percent concentrated nitric acid), and twice with tap
or other clean water. At the end of every test, all items
that are to be used again should be immediately emptied,
rinsed with water, cleaned by a procedure appropriate for
removing the test material (e.g., acid to remove metals and
bases; detergent, organic solvent, or activated carbon to
remove organic compounds), and rinsed twice with tap or
other clean water. Acid is useful for removing mineral
deposits, and 200 mg of hypochlorite/liter is useful for
removing organic matter and for disinfection. A solution
containing 200 mg hypochlorite per liter is conveniently
19
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EG-21
August, 1982
prepared by adding 6 ml of liquid household chlorine bleach
to 1 liter of water. However, acid and hypochlorite should
not be mixed because hazardous fumes may be produced.
Metering systems and test chambers should be rinsed with
dilution water just before use.
(v) Dilation water—(A) General requirements. (JJ An
adequate supply of dilution water that is acceptable to the
test organisms and to the purpose of the test should be
available. A minimum criterion for an acceptable dilution
water is that healthy test organisms will survive in it for
the duration of acclimation and testing without showing
signs of stress, such as discoloration or unusual
behavior. A better criterion for an acceptable freshwater
dilution water is that test organisms will survive, grow,
and reproduce satisfactorily in it.
(^) If the dilution water is or is prepared from
dechlorinated water, it should be shown that in fresh
samples of the dilution water either (_i_) the concentration
of residual chlorine is less than 3 mg/liter or (ii) Acartia
tons a, mys id shrimp, oyster larvae, or first ins tar daphnids
can survive for 48 hours without food. The dilution water
should be assayed for the selected test material.
20
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HG-21
August, 1982
(B) Fres hwater. Because daphnids are more sensitive to
many toxicants than most other readily available freshwater
aquatic animals, water in which first ins tar daphnids will
survive for 48 hours without food is probably acceptable for
most short-term tests with freshwater animals. Water in
which daphnids will survive, grow, and reproduce
satisfactorily should be an acceptable dilution water for
longer tests with freshwater animals.
(C) Estuarine and marine water. Because Acartia tons a,
mys id shrimp, and oyster larvae are more sensitive to many
toxicants than most other estuarine and marine aquatic
animals, water in which they will survive for 48 hours
without food is probably acceptable for most short-terra
tests with estuarine and marine animals. Water in which
Acartia tonsa or mys id shrimp will survive, grow, and
reproduce satisfactorily should be an acceptable dilution
water for longer tests with estuarine and marine animals.
(e) Reporting . In addition to the reporting
requirements prescribed in Part 792--Good Laboratory
Practice Standards, the test report should include t.ie
following information:
21
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HG-21
August, 1982
(1) A detailed description of the test material,
including its source, lot number, composition (identity and
concentration of major ingredients and major impurities),
known physical and chemical properties, and identity and
concentration of any carriers (solvents) or other additives
used;
(2) The source of the dilution water, its chemical
characteristics, and a description of any pre-treatment;
(3) Detailed information about the test organisms,
including scientific name and how verified (and strain for
salmonids when appropriate), weight (wet, blotted dry),
standard length of fish, height of bivalve molluscs, age,
life'stage, source, history, observed diseases, treatments,
acclimation procedure, and food used;
(4) A description of the experimental design and
metering system;
(5) Description of tissue and water samples analyzed,
and methods used to obtain, prepare, and store them;
(6) Methods used for, and results (with standard
deviation) of all chemical analyses of water quality and
concentration of test material in tissue and water,
including validation studies and reagent blanks;
22
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EG-21
August, 1982
(7) The steady-state bioconcentration factor, the
uptake and depuration rate constants, the confidence
(_+_ standard deviation) and the method of computations/data
analysis;
(8) Anything unusual about the test, any deviation from
these procedures, and any other relevant information.
(f) References
(1) Blau GE, Neely WB, Branson DR. 1975.
Ecokinetics: a study of the fate and distribution of
chemicals in laboratory ecosystems. AICHE Jour. 21:854-861»
(2) Borthwick PW, Tagatz ME, Forester J. 1977. A
gravity-flow column to provide pesticide-laden water for ;.
aquatic bioassays. Bull. Environm. Contam. Toxicol. 13:183-
187.
(3) Chadwick GC, Kugemagi V. 1968. Toxicity
evaluation of a technique for introducing dieldrin into
water. J. Water Pollut. Control Fed. 40: 76-82.
(4) Committee on Methods for Toxicity Tests with
Aquatic Organisms. 1975. Methods for acute toxic ity tests
with fish, macroinvertebrates, and amphibians. Corvallis,
Oregon: U.S. Environmental Protection Agency. EPA-660/3-
75-009.
23
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EG-21
August, 1982
(5) Jarvinen AW, Hoffman MJ, Thorslund TW. 1977.
Toxicity of DDT food and water exposure to fathead
minnows. Duluth, Minnesota: U.S. Environmental Protection
'Agency; EPA-600/3-76-114.
(6) Lemke AE, Brungs WA, Halligan BJ. 1978. Manual
for construction and operation of toxicity testing
proportional diluters. Duluth, Minnesota: U.S.
Environmental Protection Agency. EPA-600/3-78-072
'(17).' Re illy PM, Bajramovic R, Blau GE, Branson DR,
_Sau$rh,Qff MW. '1977. Guidelines for the optimal design of
experiments to estimate parameters in first order kinetic
models. Can. J. Chem. Eng. 55: 614-622.
(8) Reinert RE. 1970. Pesticide concentrations in
Great Lakes fish. Pest. Monit. J. 3(4): 233-240.
(9) US EPA. 1974. Analysis of human or animal adipose
tissue. In: Thompson JF, ed. Analyses of Pesticide
Residues in Human and Environmental Samples.
(10) USEPA. 1975. U.S. Food and Drug
Administration. Pesticide Analytical Manual. Vol. 1.
Rockville, Md.
(11) Veith GD, Cornstock VM. 1975. Apparatus for
continuously saturating water with hydrophobic organic
chemicals. J. Fish. Res. Board Can. 32: 1849-1851.
24
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50272 -101
REPORT DOCUMENTATION
PAGE
1. REPORT NO.
EPA 560/6-82-002 Part 2
4. Title and Subtitle
Environmental Effects Test Guidelines
7. Author(s)
9. Performing Organization Name and Address
Office of Pesticides and Toxic Substances
Office of Toxic Substances (TS-792)
United States Environmental Protection Agency
401 M Street, S.W.
Washington. D.C. 20460
12. Sponsoring Organization Name and Address
3. Recipient's Accession No.
PB82-232992
5. Report Date
August, 1982
8. Performing Organization Rept. No.
10. Project/Task/Work Unit No.
11. Contract(C) or Grant(G) No.
(C) ,
(G)
13. Type of Report & Period Covered
.Annual.
14.
IS. Supplementary Notes
16. Abstract (Limit: 200 words)
These documents consitute a set of 21 environmental effects test guidelines (and,
in some cases, support documents) that may be cited as methodologies to be used
in chemical specific test rules promulgated under Section 4(a) of the Toxic ^
Substances Control Act (TSCA). These guidelines cover testing for invertebrate
toxicity, aquatic vertebrate toxicity, avian toxicity, phytotoxicity^and,^^
bioconcentration. The guidelines will be published in loose leaf form'" ana1"
updates will be made available as changes are dictated by experience and/or
advances in the state-of-the-art.
17. Document Analysis a. Descriptors
b. Identifiers/Open-Ended Terms
c. COSATI Field/Group
18. Availability Statement
Release unlimited
19. Security Class (This Report)
Unclassified
20. Security Class (This Page)
Unclassified
21. No. of Pages
22. Price
(See ANSI-Z39.18)
See Instructions on Reverie
OPTIONAL FORM 272 (4-77)
(Formerly NTIS-35)
Department of Commerce
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