&EPA
United States
Environmental Protection
Agency
Office of Chemical Safety .-DA -,.~ „ noo
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and Pollution Prevention . on.0
^7101j January 2012
Ecological Effects
Test Guidelines
OCSPP 850.2300:
Avian Reproduction
Test
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NOTICE
This guideline is one of a series of test guidelines established by the United States
Environmental Protection Agency's Office of Chemical Safety and Pollution Prevention
(OCSPP) for use in testing pesticides and chemical substances to develop data for
submission to the Agency under the Toxic Substances Control Act (TSCA) (15 U.S.C. 2601,
et seq.), the Federal Insecticide, Fungicide and Rodenticide Act (FIFRA) (7 U.S.C. 136, et
seq.), and section 408 of the Federal Food, Drug and Cosmetic (FFDCA) (21 U.S.C. 346a).
Prior to April 22, 2010, OCSPP was known as the Office of Prevention, Pesticides and Toxic
Substances (OPPTS). To distinguish these guidelines from guidelines issued by other
organizations, the numbering convention adopted in 1994 specifically included OPPTS as
part of the guideline's number. Any test guidelines developed after April 22, 2010 will use
the new acronym (OCSPP) in their title.
The OCSPP harmonized test guidelines serve as a compendium of accepted scientific
methodologies and protocols that are intended to provide data to inform regulatory decisions
under TSCA, FIFRA, and/or FFDCA. This document provides guidance for conducting the
test, and is also used by EPA, the public, and the companies that are subject to data
submission requirements under TSCA, FIFRA, and/or the FFDCA. As a guidance
document, these guidelines are not binding on either EPA or any outside parties, and the
EPA may depart from the guidelines where circumstances warrant and without prior notice.
At places in this guidance, the Agency uses the word "should." In this guidance, the use of
"should" with regard to an action means that the action is recommended rather than
mandatory. The procedures contained in this guideline are strongly recommended for
generating the data that are the subject of the guideline, but EPA recognizes that departures
may be appropriate in specific situations. You may propose alternatives to the
recommendations described in these guidelines, and the Agency will assess them for
appropriateness on a case-by-case basis.
For additional information about these test guidelines and to access these guidelines
electronically, please go to http://www.epa.gov/ocspp and select "Test Methods &
Guidelines" on the left side navigation menu. You may also access the guidelines in
http://www.requlations.qov grouped by Series under Docket ID #s: EPA-HQ-OPPT-2009-
0150 through EPA-HQ-OPPT-2009-0159, and EPA-HQ-OPPT-2009-0576.
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OCSPP 850.2300: Avian reproduction test.
(a) Scope—
(1) Applicability. This guideline is intended to be used to help develop data to submit to
EPA under the Toxic Substances Control Act (TSCA) (15 U.S.C. 2601, et seq.), the
Federal Insecticide, Fungicide, and Rodenticide Act (FIFRA) (7 U.S.C. 136, et seq.), and
the Federal Food, Drug, and Cosmetic Act (FFDCA) (21 U.S.C. 346a).
(2) Background. The source material used in developing this harmonized OCSPP test
guideline include the OPPT guidelines under 40 CFR 797.2130 Bobwhite Reproduction
Test and 797.2150 Mallard Reproduction Test; the OPP 71-4 Avian Reproduction Test
(Pesticide Assessment Guidelines Subdivision E); OECD 206, Avian Reproduction Test,
and the Pesticide Reregi strati on Rejection Rate Analysis: Ecological Effects.
(b) Purpose. This guideline is designed to develop data on the reproductive effects on the
northern bobwhite (Colinus virginianus) and mallard (Anas platyrhynchos) of chemical
substances and mixtures ("test chemicals" or "test substances") subject to environmental effects
test regulations. This guideline prescribes specific guidance for the testing of northern bobwhite
and mallard, which are the Agency's preferred test species. The Agency will use these and other
data to assess the chronic hazard and risks to birds that these chemicals may present through
environmental exposure.
(c) Definitions. The definitions in the OCSPP 850.2000 guideline apply to this test guideline.
In addition, the following more specific definitions apply, which refer specifically to the
production and quality of eggs and subsequent development of these eggs through hatching and
up to the point where young are 14 days old:
14-day-old survivors are birds that survive for 2 weeks following hatch.
Cracked eggs are 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 measures of embryo development.
Eggs set refers to all eggs placed under incubation, i.e. total eggs produced 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.
Eggshell thickness refers to the thickness of the shell and the membrane of an 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 these several measured points in millimeters (mm).
Hatchlings, normal refers to embryos that mature, pip the shell, and liberate themselves
from the eggs on day 23-25 of incubation for northern bobwhite and days 25-28 of
incubation for mallards.
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Live 18-day embryos or 21-day embryos for northern bobwhite and mallards,
respectively refers to embryos that are developing normally after 18 or 21 days of
incubation for northern bobwhite and mallards, respectively. This is determined by
candling the eggs.
Viable embryos (or fertile eggs) refers to eggs in which fertilization has occurred and
embryonic development has begun. This is determined by candling the eggs 11 days
after incubation has begun for northern bobwhite and 14 days for mallards. It is difficult
to distinguish between the absence of fertilization and early embryonic death. The
distinction can be made by breaking open eggs that appear infertile and examining
further. This distinction is especially important when a test substance induces early
embryo mortality.
(d) General considerations—
(1) Summary of the test. Adult birds are administered the test substance continuously in
their daily diet prior to the onset of breeding and continuing for an extended period after
egg laying has been initiated by photostimulation. Eggs are collected, marked, stored,
and subsequently incubated through hatching. Offspring are maintained on a clean diet
for a period of approximately two weeks after hatching. Effects on adult birds, embryos,
and hatchlings are monitored throughout the exposure period in order to assess the
potential reproductive impact of the test substance. The no observable effect
concentration (NOEC) for each of the monitored effects is determined and the most
sensitive of these endpoints is used as the overall reproductive NOEC for the test.
(2) General test guidance. The general guidance in OCSPP 850.2000 applies to this
guideline except as specifically noted herein.
(3) Range-finding test. Unless the approximate NOEC for the most sensitive
reproductive endpoint is known already, a range-finding test should be conducted to help
determine the concentrations to be used in the definitive test. If a range-finding test is
performed, a six week dietary exposure period may provide information helpful in
determining appropriate test concentrations for the definitive test.
(4) Definitive test. The objective of the definitive test is to determine the concentration-
response relationships for avian reproductive parameters from dietary exposure to a test
substance, and to determine the NOEC for reproduction. The definitive test consists of a
minimum of three dietary concentrations of the test substance, plus a control. The dietary
levels are confirmed by chemical analysis under test conditions. A list of reproductive
response variables that are evaluated to determine a reproductive NOEC are in Table 1.
A summary of test conditions is provided in Table 2 and validity elements for an
acceptable definitive test in Table 3.
(e) Test standards—
(1) Test substance. The substance to be tested should be technical grade unless the test
is designed to test a specific formulation, mixture, or end-use product. For pesticides, if
more than one active ingredient constitutes a technical product, then the technical grade
of each active ingredient should be tested separately, in addition to the combination, if
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applicable. The OCSPP 850.2000 guideline lists the type of information that should be
known about the test substance before testing and discusses methods for preparation of
the test substance in the diet for use in testing. The Agency should be contacted prior to
testing with nanomaterials.
(2) Test duration. The definitive test consists of three phases following acclimation to
test facilities. The initial phase begins with exposure of treatment groups of adult birds to
diets containing the test substance and is typically 6 to 8 weeks long. After the initial
phase, the light/dark photoperiod is manipulated to bring the hens into laying condition
during the second phase. This second (photostimulation) phase ends with the onset of
egg-laying and is typically 2 to 4 weeks long. Unless otherwise specified, test birds
should be exposed for at least 10 weeks prior to the onset of egg laying. The final phase
begins with the onset of laying and lasts for at least 8 weeks, preferably 10 weeks. A
withdrawal study period may be added to the test if reduced reproduction is observed and
the test substance is bioaccumulative. The withdrawal period, if used, need not exceed 3
weeks.
(3) Test organisms—
(i) Species. These test protocols and standards describe tests specific to using the
northern bobwhite (Colinus virginianus (L.)) an upland game bird, and the
mallard (Anas platyrhynchos (L.)) for a waterfowl. Test birds should be pen-
reared. The Agency will consider alternative species on a case-by-case basis.
(ii) Source. Birds may be reared in the laboratory or purchased from a breeder.
For a satisfactory test, all control and experimental birds used in a test should be
from the same source, breeding population, and strain. Purchased birds should be
certified as disease-free or as bred from disease-free stocks. Rearing stock and/or
test birds should be obtained only from sources that have met the requirements for
"U.S. Pullorum-Typhoid Clean" classification under paragraph (j)(6) of this
guideline. Birds should be obtained only from sources whose colonies have
known breeding histories. Steps should be taken to prevent inbreeding. 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 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 heterogeneity of naturally occurring birds.
(iii) Age. Adult test birds used are those approaching their first breeding season
and are at least 16 weeks old. All test birds should be the same age within one
month.
(iv) Acclimation. Test birds should be acclimated to test facilities and untreated
basal diet for at least 2 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.
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(v) Health status. All birds should have a health observation period of at least 2
weeks prior to selection for treatment. Birds used in the test should be in apparent
good health. Birds should not have been selected in any way for resistance to
toxic substances. Birds are not used for testing under the following conditions.
(A) They are deformed, abnormal, sick, or injured.
(B) More than 3 percent (3%) of either sex of a population of birds
becomes debilitated during the health observation period.
(C) Birds were used in a previous test, either in a control or test substance
treatment group, or they are offspring of birds used in a test substance
treatment group in a previous test. However, offspring of birds used as a
control in a previous test are acceptable. Control offspring may be reared
and used in another test as adults.
(vi) Care and handling. 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.
(vii) Diet and feeding—
(A) Adult birds. 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 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
ingredients) of the diet should be included with the test report.
(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.
(viii) Water. 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. Antibiotics or other medication
should not be used in the water of breeding birds. Bacitracin, or one of its forms,
may be added to the drinking water of young birds, if necessary.
(4) Administration of test substance. The test substance is administered in the diet with
ad libitum feeding. Any test diet remaining in feed trays should be discarded before fresh
test diet is provided.
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(i) Preparation of diet treatments.
(A) The test substance should be mixed into the diet in a manner that will
ensure even distribution of the test substance throughout the diet. 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. If possible, the test substance should
be added to the diet without the use of a vehicle or diluent. If a diluent is
needed, the preferred diluent is reagent 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 arable (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.
(B) 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 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 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 20%. If the test
substance is known or found to be volatile or labile to the extent that 20%
or more loss occurs within 1 week, then test substance diets should be
prepared (freshly or from frozen concentrate) at a frequency that will
prevent more than 20% loss of test substance. The Agency should be
contacted prior to testing with nanomaterials.
Sampling frequency and analysis to confirm dietary test substance
concentrations and stability are conducted at a minimum as described in
paragraph (e)(9)(i) of this guideline.
(ii) Treatment concentrations. Test concentrations of the test substance should
be based on measured or calculated residues expected in the diet, unless otherwise
specified. There are at least three test substance treatment groups and a control
group. One test substance concentration should be equal to or greater than an
actual or expected field residue exposure level. One test substance concentration
should indicate a reproductive effect, (capturing the Least Observed Effect
Concentration (LOEC)) or be greater than 5,000 mg/kg-diet or higher, and one
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test substance concentration should be free of biological effects (capturing the no-
observed effect concentration (NOEC)). The highest nonlethal concentration may
be estimated from the concentration-response data generated in an avian dietary
toxicity (LCso) test or from a range-finding test. For pesticides, if the expected
environmental concentration exceeds 5,000 mg/kg-diet, then the estimated
environmental field concentration should be used in place of the 5,000 mg/kg-diet
concentration. For pesticides, reasonable upper bound expected environmental
field residue exposure level can be estimated using the Kenaga nomogram as
modified by Fletcher et al. (see references in paragraphs (j)0) and G)(2) of this
guideline) for short grass and the highest pesticide label rate (in Ibs a.i./acre) —
accounting for multiple applications that occur in a season. For example, at 1 Ib
a.i./acre applied twice in a season (i.e., 2 Ib a.i./acre) and using the Kenaga
nomogram for short grass (1 Ib a.i./acre—240 mg/kg) the upper bound residue
level is 480 mg a.i./kg-diet. This conservatively assumes no degradation of the
parent between applications.
(5) Controls.
(i) Every test includes a concurrent control treatment. The control birds are from
the same breeding population as the test substance treatment groups and are kept
under the same experimental conditions as the test substance treatment groups.
The test procedures are the same for control and treated birds, except that no test
substance is added to the diets of control birds. If a vehicle or diluent is used in
preparation of the test diets, the same diluent is added to the diets of control birds
in the highest concentration used for the test diets.
(ii) For a satisfactory test, the following values for response variables in controls
should be met or at least approached at test termination. There is likely to be a
problem with test procedures or conditions that should be investigated and
corrected when these values are not met.
(A) Eggs laid. Normal values for both northern bobwhite and mallards are
29 to 61 eggs per hen for a 10 week egg laying period.
(B) Eggs cracked. Normal values for northern bobwhite are 0 to 7.0% of
eggs laid. Normal values for mallards are 0 to 4.0% of eggs laid.
(C) Fertility (viable embryos). Normal fertility values for northern
bobwhite and mallards are 80 to 100% of eggs set.
(D) Live 18-d or 21-d northern bobwhite and mallard embryos,
respectively (as a percentage of viable embryos). Normal values for
northern bobwhite are 97 to 100%. Normal values for mallards are 94 to
100%.
(E) Hatchability (percentage of 18-d or 21-d northern bobwhite and
mallard embryos, respectively that hatch). Normal values for northern
bobwhite are 85 to 100%. Normal values for mallards are 52 to 100%.
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(F) Percentage of eggs set that hatch. Normal values for northern
bobwhite are 71 to 95%. Normal values for mallards are 44 to 92%.
(G) 14-day-old survivors of eggs hatched. Normal values for northern
bobwhite are 77 to 100%. Normal values for mallards are 94 to 100%.
(H) Eggshell thickness. Normal average values for northern bobwhite are
0.20 to 0.24 mm. Normal values for mallards are 0.316 to 0.372 mm.
(6) Number of test organism and replicates.
(i) The experimental unit for this test is the pen. All control and treatment birds
should be randomly distributed to pens from the same population. For northern
bobwhite and mallard, each of the test substance groups and the control group
consist of a minimum of 16 replicate pens. Each pen contains one male and one
female. The use of 20 replicate pens in the control group may yield a test with
greater statistical power.
(ii) An alternative arrangement of birds may consist of multiple female birds
(typically two) and one male bird in each pen. For this arrangement, each pen is
considered a replicate. Productivity should be calculated on a per hen basis, with
an average given for each pen. Either arrangement is acceptable if productivity
reaches the definitive values given in (e)(5)(ii)(A) of this guideline. Because the
behavioral interactions of birds in the two arrangements are 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.
(iii) 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. Birds should be marked
with leg bands.
(7) Facilities, apparatus and supplies. Normal laboratory equipment and supplies, and
items especially listed in (e)(7)(i) through (e)(7)(vi).
(i) Facilities. Pens should be kept indoors in order to better control lighting,
temperature, humidity, and other factors. Outdoor pens should only be used
during the normal breeding season.
(ii) Breeding pens or cages—
(A) Size. The Agency recognizes that minimum cage size
recommendations are evolving over time. The use of a certain cage size,
as with any husbandry parameter, should result in control birds with no
overt signs of stress (e.g., reproductive results are within test validity
elements reported in this guideline). Northern bobwhite and mallards
should be housed in breeding pens or cages of adequate size conforming to
good husbandry practices (see the most recent standards of good
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husbandry including, but not limited to, references provided in this
guidance document).
(B) Construction materials. The preferred construction materials are
stainless steel, galvanized sheeting, and wire mesh. For enclosed cages,
floors and external walls may be wire mesh; and ceilings and common
walls solid sheeting. Wire mesh for floors should be fine enough so as to
not interfere with normal movement of the birds. Open-topped pens may
be constructed of the same materials for the side walls and wire mesh or
concrete for the floor. 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 a pen. Wood should not be used for floors or
lower sides of pens unless it has been coated with a nonadsorbent material
such as perfluorocarbon plastic (e.g. Teflon), or unless the wood is
replaced between tests.
(C) Cleaning.
(1) Pens should be disassembled (if feasible) and cleaned
thoroughly between tests. Any used floor litter is discarded.
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. The floor composition will dictate methods
used to clean the floor. 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.
(2) During the test, pens should be cleaned when necessary.
However, care should be taken to keep disturbance to a minimum
as birds are not to be removed from cages during cleaning.
(iii) Egg storage, incubators and hatchers. Storage and incubator equipment of
sufficient size to store all eggs laid over a two week period and incubate all eggs
generated during the study. All eggs should be set after candling for incubation in
a commercial incubator. Storage equipment and incubators should be able to
maintain stable temperature and humidity conditions. Stored and incubated eggs
are turned daily. If incubators are not equipped to turn eggs automatically, they
will need to be turned daily by hand. Eggs are removed to a separate incubator or
hatcher on day 21 for northern bobwhite and day 24 for mallard. Forced draft
incubators or hatchers should be used.
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(iv) Candling. Candle lamps to check all eggs at set observation times for: fine
cracks in egg shells; infertile eggs; and dead embryos.
(v) Brooder pens. After hatching, chicks or ducklings are 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 controlled device.
(vi) Cleaning. All materials that will come in contact with the test organisms and
test substance should be cleaned before use. Cleaning procedures should be
appropriate to remove known or suspected contaminants.
(8) Environmental conditions—
(i) Temperature and humidity—
(A) Adult birds. Temperature and humidity should be controlled during
the study. The recommended temperature level for adult birds is 15 to 30
degrees Celsius (°C) with approximately 45 to 70% relative humidity.
(B) Eggs—
(1) Storage period. All eggs are collected daily, marked
according to the pen from which collected, and stored at 13 to 16
°C and 55 to 80% relative humidity. Storage in plastic bags may
improve uniformity of hatching. Stored eggs should be turned
daily. Stored eggs are set weekly or every other week for
incubation.
(2) Incubation period. During the incubation period, eggs should
be maintained at 37.5 °C ± 1 °C and approximately 70% relative
humidity.
(C) Brooder pens. For hatchlings, a temperature gradient in the brooder
pen from approximately 35 °C to 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.
Humidity should be approximately 70%.
(ii) Lighting and photoperiod—
(A) Adults.
(1) 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 65 lux (for sunlight, equivalent to
approximately 1.2 micromoles per square meter per second
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(|imol/m /s) with a minimum of 10 lux (0.2 |imol/m /s) at the level
of the birds.
(2) 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 Table 3 of this guideline is acceptable as long as birds
are exposed to treated diets a minimum of 10 weeks prior to the
onset of laying. Regardless of the methods selected, lighting
should be controlled carefully, preferably by automatic timers. A
15 to 30 minute transition period between the light and dark
periods is desirable. In addition, it is important during the initial
phase to not interrupt the dark period unless absolutely necessary.
(3) A suggested photoperiod regime would consist of maintaining
birds under a photoperiod for 7 or 8 hours of light during the initial
phase. At the end of the initial phase, the photoperiod may be
increased to 16 to 17 hours of light per day. The photoperiod may
be maintained at this level for the remainder of the study, or it may
be increased each week by 15 minutes per day.
(B) Chicks and ducklings. Lighting should be on a diurnal basis (e.g. 16
hours of light, 8 hours of dark, with a 15-30 minute transition at dawn and
dusk, but other lighting regimes are acceptable).
(iii) Ventilation. Good ventilation should be maintained. Suggested ventilation
rates are 10 to 15 changes per hour.
(9) Observations-
(i) Measurement of test substance. Samples of treated diets should be analyzed
to confirm proper dietary concentrations of the test substance under actual test
conditions. The analytical method used to determine test substance
concentrations shall be validated before beginning the test, as described in OCSPP
850.2000. During the exposure period, analyses should be conducted on
representative samples of test feed taken from feeders of all test concentrations at
the beginning of the exposure period, midway through the test (10 to 12 weeks
later), and at the end of the exposure period. 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.
(ii) Contaminants in feed. Diets should be analyzed periodically to identify
background contaminants such as heavy metals (e.g., arsenic, cadmium, lead,
mercury, and selenium) and persistent pesticides, especially chlorinated
insecticides. A broader pesticide screen to include other chemicals (e.g.,
organophosphorus pesticides) may be useful.
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(iii) Basal diet composition. A nutrient analysis of the basal diet should be
included with the test report. The analysis should include percentages by weight
of protein, fat, fiber, ash, calcium, and phosphorus. In addition to these analyzed
components, a list of expected amounts of vitamins, minerals or other
supplements should also be reported. Most commercial feed companies provide
both the analysis and the list of supplements on the label.
(iv) Environmental conditions—
(A) Temperature. Temperature should be recorded at least weekly at the
same time of day. 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 to 4 centimeters (cm) above the floor of
the cage.
(B) Humidity. Humidity should be monitored on a constant basis in at
least one representative location.
(v) Measures of effect. All calculations and formulae provided below assume
each pen (replicate) consisted of one male bird and one female bird. If an
alternative design is used (e.g., one male bird and two female birds per pen),
formulae will need to be adjusted accordingly; consultation with a statistician is
recommended. The measurement interval for the determination of the NOECs
should commence at onset of exposure of adults and finish at the end of the
exposure period (typically 8-10 weeks after egg laying starts). All outcomes from
eggs laid during the exposure period will be included in the statistical analysis. If
a withdrawal period is used, separate summary data and statistics should be
calculated based solely on data obtained from the withdrawal period. A
statistician should be consulted for statistical analysis to compare data from the
exposure and withdrawal periods.
(A) Adult birds—
(1) Body weight and food consumption. 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. Taking 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 biweekly throughout the rest of the study.
(2) Signs of toxicosis. Observations on adult birds should be made
at least once a day. or other signs of toxicity should be described
and recorded by date or day of test.
(3) Gross pathology. Gross pathological examinations should be
conducted on all birds that die during the test period, and for all
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survivors at the end of the test. At a minimum, the examination
should include the GI tract, liver, kidneys, heart, reproductive
organs, and spleen. The subcutaneous fat and muscles should also
be examined for evidence of deterioration. It is preferred that a
sufficient number of samples of two or more tissues (e.g. muscle,
fat) be analyzed for test substance residues unless it can be
demonstrated that the elimination rate is less than 24 hours.
(B) Eggs—
(1) Eggs laid. All eggs laid should be collected daily, counted and
marked according to the pen from which collected, and stored.
Storage in plastic bags may improve uniformity of hatching.
Stored eggs should be turned daily. Eggs should be removed daily,
marked, and stored until there is a sufficient quantity for
incubation.
(2) Cracked eggs, egg shell thickness, and eggs set.
(a) At weekly or biweekly intervals, eggs should be
removed from storage and be candled to detect eggshell
cracks. All eggs should be candled at day 0 for cracks and
all cracked eggs are counted and discarded. Except for
eggs with cracked shells and those eggs removed for
eggshell thickness measurements, all eggs should be set
after candling for incubation and the number of eggs set
recorded.
(b) Once every 2 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 shell 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 millimeter (mm)
units.
(3) Fertility and early death of embryos. Eggs should be
candled again on day 11 for northern bobwhite or day 14 for
mallards of incubation to determine fertility and early death of
embryos.
(4) Embryo survival. A final candling should be done on day 18
for northern bobwhite or day 21 for mallards to measure embryo
survival. Eggs should be removed to a separate incubator or
hatcher on day 21 for northern bobwhite or day 24 for mallard.
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(C) Chicks and ducklings—
(1) Number of hatchlings. Count the number of embryos that pip
shell, and embryos that liberate themselves. Hatching will
normally be complete by the end of day 24 for both species. By
day 24 or 27 of incubation, the hatched bobwhite chicks and
ducklings, respectively, should be removed from the hatcher or
incubator. Chicks or 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.
(2) Signs of toxicosis or abnormalities. Chicks or 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.
(3) Body weight of hatchlings. Each chick or duckling is weighed
individually upon hatching. An average hatchling weight for each
pen is calculated.
(4) Body weight of 14-d-old survivors. Each chick or duckling is
weighed individually on day 14. An average hatchling weight for
each pen is calculated.
(f) Treatment of results—
(1) Response variable calculation. For all equations in paragraph (f)(l) of this
guideline, the index7 = 1 to the total number of pens per treatment group (typically 16).
(i) Adult body weight gain. The change in adult body weight (males and
females are tracked separately) between test initiation and test termination, AbWj,
for pen7 is the measure used in this test guideline to evaluate the inhibitory effects
of the test substance on adult growth. The change in adult body weight (male or
female), assuming one bird of each sex per pen, is calculated using Equation 1.
Additionally, the change in adult body weight during the course of the test prior to
the onset of laying (day 0 to 14, 14 to 28, 28 to 42, etc.) is calculated and plotted
to assess effects on the pattern of growth (e.g., t2 = day 14 and tl = day 0; t2 =
day 28 and tl = day 14).
AbWj = bwjt2 - bwjtl Equation 1
where:
bwjt = male (or female) body weight in pen 7 at time t, where tl is test
initiation; and t2 is test termination.
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(ii) Average hatchling weight. The response measure for hatchling weight is the
average hatchling weight per pen which is calculated using Equation 2.
NH ,
^HATWTk
Equation 2
where:
k = index number of a hatchling in peny from 1 to NHf,
= total number of hatchlings in peny; and
= body weight of hatchling k in peny.
(iii) Average 14-d survivor weight. The response measure for 14-d old survivor
weight is the average survivor weight per pen which is calculated using Equation
3.
HS,
Y.SURVWT,
SURVWTj = k=l /w Equation 3
where:
k = index number of a 14-d old hatchling in peny from 1 to HSf,
HSj = total number of 14-d old surviving hatchlings in peny; and
SURVWTkj = body weight of 14-d old surviving hatchling k in peny.
(iv) Average egg shell thickness. The response variable for egg shell thickness is
the average thickness per pen which is calculated using Equation 4.
THICKj = *=i /„ Equation 4
where:
k = index number of egg shell thickness measurement in peny from 1 to
mj'>
ntj = total number of eggs with shell thickness measured in peny; and
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= average shell thickness of egg k in pen /, measured as
described in paragraph (e)(9)(v)(B)(2)(b) of this guideline.
(v) Total food consumption per adult per pen. Total food consumption per
adult bird between test initiation and test termination, TFOODj, for pen j is the
measure used in this test guideline to evaluate adversion or inhibitory effects of
the test substance on consumption of food by adults. The total food consumption
per adult bird per pen is calculated using Equation 5. Additionally, the weekly (or
biweekly after the onset of laying) food consumption rate per adult (e.g., tl, t2,
etc.) during the course of the test is calculated and plotted to assess effects on the
pattern of food consumption.
%% FOOD .t\ ,_
TFOOD = Y - - Equation 5
J
t=\
where:
t = index of weekly and biweekly measurements of food consumption,
with t=l being week 1 of the study and t=term being the test or exposure
termination week;
FOODjt = total food consumption in pen7 at time t;
mjt = number of adult birds in pen7 at time t;
(vi) Proportion of uncracked eggs. The proportion of uncracked eggs is
calculated using Equation 6.
UE}. = (EL] ~ EC]/EL Equation 6
where:
ELj = total number of eggs laid in pen/ and
EQ = number of eggs cracked in pen/
(vii) Proportion of normal eggs. The proportion of normal eggs is calculated
using Equation 7.
Anr (EL,-EC,-EA,)/ _ .. _
NEj = ^ } } ]YFj Equation 7
where:
ELj = total number of eggs laid in pen/
= number of eggs cracked in pen/ and
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j = number of irregular or abnormal eggs in pen/
(viii) Other proportions. Eight additional proportions are calculated as new
variables for each pen: proportion of eggs set per eggs laid (ESj/ELj); proportion
of viable embryos per eggs laid (VEj/ELJ); proportion of live 18-d-old northern
bobwhite or 21-d-old mallard embryos per viable embryos (LEj/VEJ)', proportion
of normal hatchlings per eggs set (NHj/ESj)', proportion of hatchlings per live 18-
d-old northern bobwhite or 21-d-old mallard embryos (NHj/LEj); proportion of
hatchlings per eggs laid (NHj/ELj); proportion of 14-d-old survivors per eggs set
(HSj/ESj)', proportion of 14-d-old survivors per hatchlings (HSj/NHJ).
(2) Descriptive statistics—
(i) Environmental conditions.
(A) Calculate descriptive statistics (mean, standard deviation, minimum,
maximum) for temperature, relative humidity, and light intensity during
the three exposure phases (initial, photostimulation and laying) for adults,
and for chicks or ducklings in the brooder pens.
(B) Calculate descriptive statistics (mean, standard deviation, minimum,
maximum) for temperature, and relative humidity during egg storage and
incubation.
(ii) Dietary test substance concentrations. Calculate descriptive statistics
(mean, standard deviation, coefficient of variation, minimum, maximum) by
treatment level of the test substance concentration in the diet.
(iii) Basal diet. Calculate descriptive statistics (mean, standard deviation) of the
percentages by weight of protein, fat, fiber, ash, calcium, and phosphorus.
(iv) Reproductive response variables. For each treatment group including the
control, calculate and plot summary statistics (mean, median, minimum,
maximum, first quartile, and third quartile) for each reproductive response
variable in Table 1. Additionally, calculate the standard deviation, coefficient of
variation, standard error of mean, and 95% confidence interval of mean for each
treatment group including the controls.
(3) Percent inhibition—
(i) Inhibitory effects. Except for the two response variables, number of cracked
eggs and number of irregular and abnormal eggs, all other response variables are
expected to exhibit increasing inhibition or reduction in the measured response
with increasing test substance concentration in the diet. For all response variables
in Table 1 the percent inhibition (%I) as compared to the control at each test
substance concentration is calculated using Equation 8.
Page 16 of 25
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(c-x)(ioo) «-*•«,
%/ = -i £—/ Equation 8
C
where:
C = the control mean treatment response value (e.g. number of eggs laid);
and
X = the test substance treatment mean response value (e.g. number of eggs
laid). Stimulation or a greater response in the test substance treatment
than the control is reported as negative %I.
(ii) Stimulatory effects. For the response variables number of cracked eggs and
number of irregular and abnormal eggs, the interest is in the increase or
stimulation of these events with increasing test substance concentrations rather
than in their reduction or inhibition. The percent stimulation or increase is also
calculated using Equation 8 except stimulation is reported as negative values of
%I. Negative %I values indicate an increased or stimulatory effect over the
control response. If working with negative numbers is confusing, the analyst may
find multiplying the %I value by -1 reduces confusion when discussing the
increase in cracked eggs and irregular or abnormal eggs with increased test
substance concentration.
(4) NOEC. A NOEC and LOEC are determined for each of the reproductive response
variables in Table 1 using appropriate statistical methods. All methods used for statistical
analysis should be described completely. Experimental units (replicates) are the
individual pens within each treatment level. The overall study NOEC and LOEC are the
lowest values (most sensitive) of all response variables considered.
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Table 1.—Reproductive Response Variables To Evaluate
Measured response variables
Number of eggs laid per pen (EL])
Number of irregular or abnormal eggs per pen (EAj)
Number of cracked eggs per pen (ECj)
Number of eggs set per pen. (ESj)
Number of viable embryos per pen (VEj)
Number of live embryos (18-day-old northern bobwhite or 21 -day-old mallard embryos) per pen (LEj)
Number of normal hatchlings per pen (NHj)
Number of 14 day-old survivors per pen (HSj)
Calculated response variables
Proportion of uncracked eggs per pen (EL/- ECj)/(ELj)
Proportion of eggs set of eggs laid per pen (ES/ELj)
Proportion viable embryos of eggs set per pen (VE/ES^
Proportion of live embryos of viable embryos per pen (LE/VEj)
Proportion of normal hatchlings of eggs laid per pen (NH/ELJ)
Proportion of normal hatchlings of eggs set per pen (NH/ESj)
Proportion of normal hatchlings of live embryos per pen (NH/LEj)
Proportion of 14 day-old survivors of eggs set per pen (HSj/ESj)
Proportion of 14 day-old survivors of normal hatchlings per pen (HS/NHj)
Average egg shell thickness per pen (THICK])
Average hatchling body weight per pen (HATWTj)
Average 14 day-old survivor body weight per pen (SURVWTj)
Adult male body weight gain per pen
Adult female body weight gain per pen
Total food consumption per adult bird per pen (FOOD])
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(g) Tabular summary of test conditions. Table 2 lists the important conditions that should
prevail during this test. Meeting these test conditions will greatly increase the likelihood that the
completed test will be acceptable or valid.
Table 2.—Summary of Test Conditions for Avian Reproduction Test
Test duration
Temperature
Light quality
Light intensity
Photoperiod
Humidity
Pen size
Number of pens per
concentration level
Test species
Age of test organisms
Number of birds per
concentration level
Number of concentration levels
Administration of test substance
Measures of Effect
(Measurement Endpoints)
Test birds should be exposed for at least 1 0 weeks prior to the onset of
egg laying and for at least 8 weeks, preferably 10 weeks, following the
onset of laying
15 to 30°C for adults; A gradient between approximately 22°C and
35°C for hatchlings
Lights should emit a spectrum simulating that of daylight
10-65 lux (0.2 to 1 .2 umol/nf /s)
Variable (see text)
Approximately 45 to 70%
See the most recent standards of good husbandry including
references provided in this guidance document.
16 pens per test concentration and control are preferred.
Northern bobwhite and mallard (additional species may tested as an
option)
1 6 weeks or slightly older at study initiation
Paired design (one male and one female) is preferred
Minimum of three, plus a control group
Through diet
NOEC for each reproductive parameter (see Table 1) and feed
consumption
(h) Test validity elements. This test would be considered to be unacceptable or invalid if one or
more of the conditions in Table 3 occurred. This list should not be misconstrued as limiting the
reason(s) that a test could be found unacceptable or invalid. However, except for the conditions
listed in Table 3 and in OCSPP 850.2000, it is unlikely that a study will be rejected when there
are slight variations from guideline environmental conditions and study design unless the control
organisms are significantly affected, the precision of the test is reduced, the power of a test to
detect differences is reduced, and/or significant biases are introduced in defining the magnitude
of effect on measurement endpoints as compared to guideline conditions. Before departing
significantly from this guideline, the investigator should contact the Agency to discuss the reason
for the departure and the effect the change(s) will have on test acceptability. In the test report, all
departures from the guideline should be identified, reasons for these changes given, and any
resulting effects on test endpoints noted and discussed.
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Table 3.—Test Validity Elements for Avian Reproduction Test
1. Birds were not randomly assigned to treatment and control pens.
2. More than 10% of the control birds died or became moribund during the test.
3. The average number of eggs laid per hen in the control group was less than 29 for northern bobwhite
or mallard.
4. The number of viable embryos in the control group was less than 80% of the eggs set for northern
bobwhite or mallard.
5. The number of 18-d-old northern bobwhite and 21-d-old mallard embryos of eggs set in the control
group was less than 97% for northern bobwhite or less than 94% for mallard, respectively.
6. The number of normal hatchlings in the control group was less than 85% of the viable embryos for
northern bobwhite or less than 52% of the viable embryos for mallard.
7. The number of normal hatchlings in the control group was less than 71% of the eggs set for northern
bobwhite or less than 44% of the eggs set for mallard.
8. The number of 14 day old survivors in the control group was less than 77% of the normal hatchlings for
northern bobwhite or less than 94% of the normal hatchlings for mallard.
9. The average eggshell thickness in the control group is less than 0.20 mm for northern bobwhite or
0.316 mm for mallards.
10. There are greater than 13% cracked eggs in the control group.
(i) Reporting—
(1) Background information. Background information to be supplied in the report
consists at a minimum of those background information items listed in paragraph (j)0) of
the OCSPP 850.2000 guideline.
(2) Guideline deviations. Provide a statement of the guideline or protocol followed.
Include a description of any deviations from the test guideline or any occurrences which
may have influenced the results of the test.
(3) Test substance.
(i) Identification of the test substance: common name, IUPAC and CAS names,
CAS number, structural formula, source, lot or batch number, chemical state or
form of the test substance, and its purity (i.e. for pesticides, the identity and
concentration of active ingredient(s)), radiolabeling if any, location of label(s),
and radiopurity.
(ii) Storage conditions of the test chemical or test substance and stability of the
test chemical or test substance under storage conditions if stored prior to use.
(iii) Methods of preparation of the test substance and the treatment concentrations
used in the range-finding and definitive tests.
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(iv) If a vehicle (solvent) is used to prepare the test diet provide: the name and
source of the vehicle, the nominal concentration(s) of the test substance in the
vehicle in stock solutions, and the vehicle concentration(s) used in the test
substance diet and control diet.
(v) Storage conditions and stability of the test substance concentration in the
treated diets throughout the duration of the study.
(4) Test organisms.
(i) Scientific and common name of the species test.
(ii) Age of birds (in months) at test initiation.
(iii) History of the birds used in the test: source, name of supplier, batch or lot
number.
(iv) Description of housing for stock birds during pre-test observation and
acclimation: type, size, material of pens, and loading (number of birds per pen).
(v) Description and documentation of any acclimation performed.
(vi) Description of pre-test observation period: date, duration, feeding regime, and
environmental conditions (temperature, humidity, photoperiod, light intensity).
(vii) Results of health observations during pre-test observation period: occurrence
and rate of mortality, occurrence, type and rate of sickness and injuries.
(5) Test system and conditions. Provide a description of the test system and conditions
used in the definitive test, and any preliminary range-finding test.
(i) Date of test initiation, duration of test, and for adults the duration of initiation,
photostimulation, and egg-laying phases and withdrawal phase if applicable.
(ii) Description of housing for adult birds used in range-finding and definitive
tests: type, size, and material of pen.
(iii) Detailed description of the basal diet: source, composition, diluents (if used),
supplements (if used), and a nutrient analysis of the basal diet.
(iv) Exposure regime to test diet and watering during exposure phases and
withdrawal phase if appropriate.
(v) Frequency and method of determining food consumption per pen and any
repellancy or food palatability issues.
(vi) Description of any medication given during the test: type, frequency, and an
explanation of how it was administered and justification of why it was given.
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(vii) The number of test substance and control treatments and the nominal test
substance dietary concentrations (in mg a.i./kg-diet for pesticides).
(viii) Statement about the selection basis or source for the highest dietary test
substance concentration such as the highest expected environmental concentration
based on the highest pesticide label rate (in Ibs a.i./acre), accounting for multiple
applications that occur in a season, or actual highest measured test substance
residue levels.
(ix) Number of replicate test pens used per test substance concentration and
control treatments and number of birds of each sex per pen at test initiation.
(x) Description of arrangement of pens to prevent cross-contamination.
(xi) Methods of assigning birds to test pens, including method of randomization.
(xii) Method of marking all adult and hatchling birds and eggs.
(xiii) Methods and frequency of environmental monitoring performed during the
initiation, photostimulation, and laying phases for air temperature, humidity, and
light intensity.
(xiv) Test substance dietary residue sampling methods and frequency to document
homogeneity and stability of the test substance in the diet throughout the study
duration.
(xv) Egg collection interval from pens.
(xvi) Description of storage housing for eggs: type, size, placement in bags if
applicable.
(xvii) Interval for removing eggs for egg shell thickness determinations and the
number of eggs used per pen for egg shell thickness
(xviii) Duration of an egg in storage and frequency of setting eggs.
(xix) Frequency of turning eggs in storage and during incubation.
(xx) Times (days) eggs were candled and the purpose for which they were
candled.
(xxi) Time (days) when eggs were transferred to a hatcher.
(xxii) Time (days) when hatched eggs were removed and counted.
(xxiii) Methods and frequency of environmental monitoring performed during the
storage and incubation of eggs: air temperature and humidity.
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(xxiv) For the definitive test, a description of all analytical procedures and
additionally the accuracy of the method, method detection limit, and limit of
quantification.
(xxv) Frequency, duration, and types of observations conducted on adult birds
during initial, photostimulation, and laying phases and withdrawal phase if
appropriate, and on eggs, and hatchlings.
(6) Results.
(i) Environmental monitoring data results (air temperature, humidity and light
intensity) for adults in the initial, photostimulation, and laying phases and the
withdrawal phase if appropriate; for hatchlings; and for chicks or ducklings in
tabular form (provide raw data for measurements not made on a continuous
basis), and descriptive statistics (mean, standard deviation, minimum, maximum).
(ii) Egg storage and incubation temperature, and relative humidity results (provide
raw data for measurements not made on a continuous basis), and descriptive
statistics (mean, standard deviation, minimum, and maximum).
(iii) Mean, standard error, minimum and maximum test substance concentrations
(mg a.i./kg-diet for pesticides) by observation time and treatment level and an
evaluation of the results in terms of documenting the stability of the test substance
concentration in the diet throughout the duration of the study.
(iv) Tabulation of number of eggs laid, number of irregular or abnormal eggs,
number of cracked eggs, number of eggs set, number of viable embryos, number
of live embryos, number of normal hatchlings, and number of 14-day old
survivors by treatment, observation week, and pen. Descriptive statistics (mean,
standard deviation, standard error, 95% confidence interval, median, first and
third quartiles, minimum, maximum) and plot of these effects (mean, median, first
and third quartiles, minimum, maximum) by treatment level. Percent inhibition
calculations as compared to control. Provide sufficient raw data for performance
of an independent statistical analysis.
(v) A tabulation of percentage of eggs set of eggs laid, viable embryos of eggs set,
live 18-day old embryos, 14-day old survivors of eggs laid, and 14-day old
survivors of hatchlings. Descriptive statistics (mean, standard deviation, standard
error, 95% confidence interval, median, first and third quartiles, minimum,
maximum), plot of these effects (mean, median, first and third, minimum,
maximum) by treatment level, and a tabulation of the %I. Provide sufficient raw
data for performance of an independent statistical analysis.
(vi) A tabulation of egg shell thickness by treatment and pen and calculated
average egg shell thickness per pen. Descriptive statistics (mean, standard
deviation, standard error, 95% confidence limits, median, first and third quartiles,
minimum, maximum) of average egg shell thickness, plot of the mean, median,
first and third quartiles, minimum and maximum average egg shell-thickness by
Page 23 of 25
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treatment level, and a tabulation of the %I. Provide sufficient raw data for
performance of an independent statistical analysis.
(vii) Description of any signs of intoxication or any other abnormal behavior in
adult birds, including time of onset, duration, severity (including death), and
numbers affected (including accidental deaths or injuries), and any remissions.
(viii) List of the number of the birds for which necropsies were performed and
details of the necropsies.
(ix) For young birds, description of any signs of toxicosis or any other abnormal
behavior, including time of onset, duration, severity (including death), and
numbers affected (including accidental deaths or injuries), and any remissions
during first and second week after hatching.
(x) Tabulation of body weights of adult male and female birds by pen and
observation time (provide raw data) and the body weight gain for each sex by pen
between test initiation and termination. Descriptive statistics of body weight gain
for each sex (mean, standard deviation, standard error, 95% confidence limits,
median, first and third quartiles, minimum, and maximum) by treatment level,
plot of mean, median, first and third quartiles, minimum and maximum, and a
tabulation of the %I. Tabulation and plot of the mean change in adult body
weight by observation interval and treatment level during the course of the test
prior to the onset of laying (day 0 to 14, 14 to 28, 28 to 42, etc.).
(xi) Tabulation of body weights of surviving young at 14 days of age by pen, and
observation time (provide raw data), descriptive statistics (mean, standard
deviation, minimum, and maximum), plot of mean, median, first and third
quartiles, minimum and maximum, and a tabulation of the %I.
(xii) For adult birds and chicks or ducklings, the food consumption data results
and the calculated food consumption per bird by observation time and pen in
tabular form. Descriptive statistics (mean, standard error) and plots of food
consumption per bird by treatment level and observation time to evaluate patterns
in food consumption throughout the duration of the study. Descriptive statistics
(mean, standard deviation, median, first and third quartiles, minimum, and
maximum) and plots of these for total food consumption per bird by treatment
level.
(xiii) Description of statistical method(s) used for NOEC and LOEC
determination, including software package, and the basis for the choice of
method.
(j) References. The following references should be consulted for additional background
material on this test guideline.
(1) Fletcher, J.S., I.E. Nellesson and T.G. Pfleeger. 1994. Literature review and
evaluation of the EPA food-chain (Kenaga) nomogram, an instrument for estimating
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pesticide residues on plants. Environmental Toxicology and Chemistry 13(9): 1383-
1391.
(2) Hawkins, P., Morton, D.B., Cameron, D., Cuthill, I, Francis, R., Freire, R., Gosler,
A., Healy, S., Hudson, A., Inglis, I, Jones, A., Kirkwood, J., Lawton, M., Monaghan, P.,
Sherwin, C., and Townsend, P., 2001. Laboratory birds: refinements in husbandry and
procedures. Laboratory Animals. 35(1): 1-163. October, 2001
(3) Hoerger, F. and E.E. Kenaga. 1972. Pesticide residues on plants: correlation of
representative data as a basis for estimation of their magnitude in the environment. In F.
Coulston and F. Corte, eds. Environmental Quality and Safety: Chemistry, Toxicology
and Technology, Volume 1. Georg Theime Publishers, Stuttgart, Germany, pp 9-28.
(4) National Research Council of the National Academies. 2010. Guide for the Care and
Use of Laboratory Animals. The National Academies Press, Washington, D.C.
(5) Organization for Economic Co-operation and Development, 1984. TG-206, Avian
Reproduction Test, adopted April 1984.
(6) U.S. Environmental Protection Agency, 1982. Pesticide Assessment Guidelines
Subdivision E, Hazard Evaluation: Wildlife and Aquatic Organisms. Office of Pesticide
and Toxic Substances, Washington, D.C. EPA-540/9-82-024, October 1982.
(7) U.S. Environmental Protection Agency, 1994. Pesticide Reregi strati on Rejection
Rate Analysis: Ecological Effects. Office of Prevention, Pesticide and Toxic Substances,
Washington, D.C. EPA 738-R-94-035, December, 1994.
(8) U.S. Department of Agriculture, 1979. National Poultry Improvement Plan, Report
No. 2., in Directory of Participants Handling Waterfowl, Exhibition Poultry, and Game
Birds. USD A, Science and Education Administration, Beltsville, MD 20705.
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