United States

Environmental Protection 

Agency

Office of Chemical Safety

and Pollution Prevention

(7101)

EPA 712-C-023

January 2012

Ecological Effects

Test Guidelines

OCSPP 850.2300:

Avian Reproduction

Test
i

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.regulations.gov 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 Reregistration 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 concentrationresponse 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 
Page 3 of 25

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 penreared. 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.
Page 4 of 25

(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.
Page 5 of 25

(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 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.

(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 
Page 6 of 25

test substance concentration should be free of biological effects (capturing the noobserved effect concentration (NOEC)). The highest nonlethal concentration may 

be estimated from the concentration-response data generated in an avian dietary 

toxicity (LC50) 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)(1) and (j)(2) of this 

guideline) for short grass and the highest pesticide label rate (in lbs a.i./acre) —

accounting for multiple applications that occur in a season. For example, at 1 lb 

a.i./acre applied twice in a season (i.e., 2 lb a.i./acre) and using the Kenaga

nomogram for short grass (1 lb 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.
Page 9 of 25

(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 (oC) 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 

oC 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 oC ± 1 oC and approximately 70% relative 

humidity.

(C) Brooder pens. For hatchlings, a temperature gradient in the brooder 

pen from approximately 35 oC to 22 oC 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 
Page 10 of 25

(μmol/m2/s) with a minimum of 10 lux (0.2 μmol/m2/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 

-15oC 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.
Page 11 of 25

(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 
Page 12 of 25

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.
Page 13 of 25

(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)(1) of this 

guideline, the index j = 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, bwj, 

for pen j 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 t1 = day 0; t2 = 

day 28 and t1 = day 14).

bwj  bwjt2 bwjt1 Equation 1

where:

bwjt = male (or female) body weight in pen j at time t, where t1 is test 

initiation; and t2 is test termination.
Page 14 of 25

(ii) Average hatchling weight. The response measure for hatchling weight is the 

average hatchling weight per pen which is calculated using Equation 2.

j

NH

k

k j

j NH

HATWT

HATWT

j



 1 Equation 2

where:

k = index number of a hatchling in pen j from 1 to NHj;

NHj = total number of hatchlings in pen j; and

HATWTkj = body weight of hatchling k in pen j.

(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.

j

HS

k

k j

j HS

SURVWT

SURVWT

j



 1 Equation 3

where:

k = index number of a 14-d old hatchling in pen j from 1 to HSj;

HSj = total number of 14-d old surviving hatchlings in pen j; and

SURVWTkj = body weight of 14-d old surviving hatchling k in pen j.

(iv) Average egg shell thickness. The response variable for egg shell thickness is 

the average thickness per pen which is calculated using Equation 4.

j

m

k

k j

j m

THICK

THICK

j



 1 Equation 4

where:

k = index number of egg shell thickness measurement in pen j from 1 to 

mj;

mj = total number of eggs with shell thickness measured in pen j; and
Page 15 of 25

THICKkj = average shell thickness of egg k in pen j., 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., t1, t2, 

etc.) during the course of the test is calculated and plotted to assess effects on the 

pattern of food consumption.





















term

t jt

jt

j

m

FOOD

TFOOD

1

Equation 5

where:

t = index of weekly and biweekly measurements of food consumption, 

with t=1 being week 1 of the study and t=term being the test or exposure 

termination week;

FOODjt = total food consumption in pen j at time t;

mjt = number of adult birds in pen j at time t;

(vi) Proportion of uncracked eggs. The proportion of uncracked eggs is 

calculated using Equation 6.

 

j

j j

j EL

EL EC UE 

 Equation 6

where:

ELj = total number of eggs laid in pen j; and

ECj = number of eggs cracked in pen j.

(vii) Proportion of normal eggs. The proportion of normal eggs is calculated 

using Equation 7.

 

j

j j j

j EL

EL EC EA NE  

 Equation 7

where:

ELj = total number of eggs laid in pen j;

ECj = number of eggs cracked in pen j; and
Page 16 of 25

EAj = number of irregular or abnormal eggs in pen j.

(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 17 of 25

  

C

C X

I

100 %

 Equation 8

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.
Page 18 of 25

Table 1.―Reproductive Response Variables To Evaluate

Measured response variables

Number of eggs laid per pen (ELj)

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 (ELj – ECj)/(ELj)

Proportion of eggs set of eggs laid per pen (ESj/ELj)

Proportion viable embryos of eggs set per pen (VEj/ESj)

Proportion of live embryos of viable embryos per pen (LEj/VEj)

Proportion of normal hatchlings of eggs laid per pen (NHj/ELj)

Proportion of normal hatchlings of eggs set per pen (NHj/ESj)

Proportion of normal hatchlings of live embryos per pen (NHj/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 (HSj/NHj)

Average egg shell thickness per pen (THICKj)

Average hatchling body weight per pen (HATWTj)

Average 14 day-old survivor body weight per pen (SURVWTj)

Adult male body weight gain per pen ( ) bwj

Adult female body weight gain per pen ( ) bwj

Total food consumption per adult bird per pen (FOODj)
Page 19 of 25

(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 Test birds should be exposed for at least 10 weeks prior to the onset of 

egg laying and for at least 8 weeks, preferably 10 weeks, following the 

onset of laying 

Temperature 15 to 30oC for adults; A gradient between approximately 22oC and 

35oC for hatchlings

Light quality Lights should emit a spectrum simulating that of daylight

Light intensity 10 - 65 lux (0.2 to 1.2 μmol/m2/s)

Photoperiod Variable (see text)

Humidity Approximately 45 to 70%

Pen size See the most recent standards of good husbandry including 

references provided in this guidance document.

Number of pens per 

concentration level

16 pens per test concentration and control are preferred.

Test species Northern bobwhite and mallard (additional species may tested as an 

option)

Age of test organisms 16 weeks or slightly older at study initiation

Number of birds per 

concentration level

Paired design (one male and one female) is preferred

Number of concentration levels Minimum of three, plus a control group

Administration of test substance Through diet 

Measures of Effect 

(Measurement Endpoints)

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.
Page 20 of 25

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)(1) 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.
Page 21 of 25

(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.
Page 22 of 25

(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 lbs 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.
Page 23 of 25

(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 24 of 25

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., J.E. Nellesson and T.G. Pfleeger. 1994. Literature review and 

evaluation of the EPA food-chain (Kenaga) nomogram, an instrument for estimating 
Page 25 of 25

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 Reregistration 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. USDA, Science and Education Administration, Beltsville, MD 20705.