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DRAFT

September 2011

OECD GUIDELINE FOR TESTING OF CHEMICALS

Avian Two-generation Toxicity Test in the Japanese Quail

INTRODUCTION

1. The purpose of this guideline is to describe a laboratory testing procedure for

evaluating the impact of pesticides and other chemicals upon avian health and 

reproduction. Data collected from such tests may be used in the assessment of 

risks to birds exposed to pesticides and other chemicals under field conditions. 

These data from a two-generation test are principally intended to evaluate the 

adverse consequence of exposure to a “possible” endocrine disruptor to the 

reproductive viability of F1 birds, but may also provide information to confirm a 

chemical as an endocrine disruptor. 

2. The test is designed for dietary exposure, but also may include exposure through 

drinking water if more appropriate for the chemical. This guideline describes a 

two-generation test. The test monitors health and reproductive parameters in 

Japanese quail (Coturnix japonica) adults and their progeny, including viability of 

second generation offspring. 

3. The number of second generation (F2) 14-day old survivors per F1 generation hen 

is the primary biological endpoint of this test. Additional hormone and 

morphological endpoints are included to assess and confirm potential endocrine 

disruption. 

PRINCIPLE OF THE TEST

4. Birds in the parent (F0) generation are paired at 4 weeks of age, and acclimated 

for 2 weeks prior beginning chemically treated diet. Dosing begins in the F1 

generation at hatch. Parameters for adult (F0 and F1), egg production (F0 and 

F1), and offspring health (F1 and F2) are evaluated by making statistical 

comparisons between treated groups and the control group. The no observed 

effect concentration (NOEC) and the lowest observed effect concentration 

(LOEC) are determined for health, fitness and endocrine parameters.

VALIDITY CRITERIA

5. The test substance concentration in the diet to which birds are exposed should be 

satisfactorily maintained and reported. Losses of 20% or less relative to initial 

concentrations are generally considered acceptable. Higher rates of loss must be 

investigated and explained.
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6. Parental mortality during the two week acclimation should not exceed 3%. When 

possible, all mortalities in the control group should be explained. At least eight 

breeding F1 pairs that have produced eggs must be available in the control group 

at the end of the test period.

DESCRIPTION OF THE METHOD

Test animals

7. The test species is the Japanese quail (Coturnix japonica). 

8. Japanese quail generally start laying eggs at or later than six weeks of age. Once 

egglaying has begun it will take about two to three weeks for birds to reach full 

egg production. Depending on the strain, eggs will be fertile when birds are about 

eight to ten weeks of age. 

9. Birds used in the test should appear healthy and be free of abnormalities or 

injuries that may affect test results. Birds should not receive any dietary 

medications beginning one week prior to the start of the test and continuing until 

the test is terminated. All F0 birds used in a test should be from the same hatch.

Husbandry

10. Temperature, ventilation, and light controlled facilities are needed throughout the 

test. Recommended ventilation is about 8 to 15 air changes per hour. Sound and 

vibrations or other disturbances in the rooms should be kept to a minimum to 

minimize stress.

11. Artificial lighting should approximate the daylight visual spectrum and be 

automatically controlled. The photoperiod for adult birds, from the start of 

acclimation onwards, and the chicks is 16 or 17 hours light and 7 or 8 hours of 

darkness. Birds should be exposed to light intensity of at least 10 lux, measured 

at the level of the feeder. The light intensity is dictated by the amount required 

for daily observations. Light intensity that is too high will encourage aggressive 

behavior.

12. Incubators and hatchers, preferably with automatic temperature and humidity 

controls and an egg-turning device, are necessary. In addition, suitable equipment 

is required to maintain stored eggs within the temperature and humidity ranges 

specified.

13. Suitable pens of appropriate size for adults and for chicks are required. Wire pens 

with slanting floors and egg-catchers or other measures to prevent breakage of 

eggs, are recommended for adults. Pens for both adults and chicks should 

preferably be of stainless or galvanized steel or other inert materials. When using 

wire meshing for the floors, the wire mesh should be of a size sufficient to prevent 

foot injury (trauma or pododermatitis) but large enough to allow excreta to drop 

through. Measures should be taken to minimize spillage of diet (e.g., covering the 

food troughs with a wire grid).
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14. Birds should be placed in breeder pens at four weeks of age, and then housed in 

opposite sex pairs (one male/one female). The parental cages should be 

distributed in such a way that cross-contamination and positional effects are 

avoided. Pen mate aggression can be a problem and measures must be taken to 

minimize injuries, stress and mortality resulting from such aggression. Pairs 

should be reintroduced daily to maintain fertility. 

15. For rearing chicks, the use of rearing pens with thermostatically controlled warm 

compartments or cages, which are free of drafts and have a radiator (e.g., 

ceramics), is recommended. Sufficient space for feeding and drinking must be 

provided, especially during the first week after hatching, to promote the weaker 

animals accessing food and water resources. Chicks should be identified 

individually or by pen of origin. They may be housed in groups of approximately 

equal number, by treatment group. The optimal number of chicks per pen 

depends on pen size. Birds must be separated into pairs (one male/one female) 

when 4 weeks old after sex has been determined.

16. Guidance for housing conditions is provided in Table 1.

Table 1: Housing conditions

Age

(week)

Temperature

(

o

C)

Relative humidity

(%)

Minimum floor area

(cm2/bird)

1 35-38 40-80 50

2 30-35 40-80 75

3-4 23-27 40-80 100

>4 16-27 40-80 625

Note: The acceptability of housing conditions of adults will be evaluated on the basis of 

results of reproductive performance.

Feeding

17. Diet and drinking water are provided ad libitum. The diet should be described 

and should meet the specific nutrient requirements of Japanese quail. The caloric 

contents of the diet must be reported. If the same diet is used for chicks and 

adults, extra calcium should be added for the adult diet. During the treatment 

period, birds are fed basal diet mixed with the test substance at specified 

concentrations.

18. Guidance for recommended nutritional values is provided in Table 2.

Table 2: Recommended nutritional values

Nutrient

Adults (> 4weeks)

Recommended range (%)

Chicks 

Recommended range (%)

Crude protein 27 to 29 27 to 30

Crude fibre 3.5 to 5.0 3.0 to 6.0
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Crude fat 2.5 to 7.0 5.5 to 7.5

Calcium 2.6 to 3.6 0.75 to 1.2

Phosphorous 0.9 to 1.1 0.6 to 1.0

TEST DESIGN

19. Care should be taken to ensure that birds are drinking water as they transition 

from group pens to the breeder pens. An acclimation period of at least two weeks 

is required for the F0 generation prior to the start of the test. The test begins with 

the administration of treated diet (i.e. when birds are 6 weeks old, at the earliest). 

Breeding pairs should be maintained in a 16:8 or 17:7 light:dark photoperiod for 

the duration of the study. Onset of egglaying can take place during the 

acclimation period.

Prior to the start of treatment, individual F0 generation birds are weighed, sexed

(preferably, using genetic techniques), and impartially allocated to pens and 

treatment groups. Birds are paired by known lineage such that males from one 

line are paired with females from another line. Pairs should be observed for signs 

of excessive aggression, and if present the sexes separated except for periodic 

conjugal visits. A translucent separator wall may be used for this purpose in each 

pen. Otherwise, the pairs will be housed together in the same pen. When 

together, signs of excessive aggression should be monitored and the males and 

females separated if injury is imminent. The number of pairs allocated to each 

group (for example 16) should be sufficient to ensure that there are 12 breeding 

pairs in the F0 control group that have successfully produced eggs at the end of 

the treatment period. Eggs are collected from F0 post-treatment week 5 (and 

possibly, week 8 and/or 9) to examine fertility. Broods from the post-treatment 

6th and 7th week are used to establish the first generation offspring (F1). 

Optimally, 6 eggs are set for each parental pair, but a minimum of 40 eggs must 

be set for each test level. If during the 6th and 7th post-treatment weeks, a 

treatment level does not produce at least 40 eggs, that level is dropped from 

further analysis in the test. At approximately two weeks of age F1 control and 

treatment chicks are genetically sexed. At four weeks of age, F1 control and 

treatment group birds are weighed and randomly allocated to pens by pairs with 

male offspring from odd numbered parental pens being paired with females from 

even numbered parental pens and even numbered pen males being paired with 

odd numbered females. The F1 generation breeding birds are maintained until 6 to 

7 weeks post-fertility. Eggs collected from F1 pairs during weeks 1 through 5 

will be used to evaluate fertility. Eggs collected from weeks 6 and 7 post-pairing 

will be used to establish the F2 generation, optimally 6 eggs will be collected 

from each F1 breeding pair. F2 offspring survival will be observed though day 14. 

Observations can be extended to examine timing of sexual maturity in the F2 

generation. 
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Dietary concentrations

20. The concentration of the test substance in the diet is expressed as weight of the 

test substance per unit weight of diet with known water content: mg/kg of diet (or 

weight of test substance per weight of water when birds are exposed through 

drinking water). The concentration should also be expressed as mg/kg body 

weight per bird per day.

21. Dietary concentrations of the test substance should be chosen on the basis of 

toxicological data from a range-finding study or other preceding avian tests. 

Information may also be gained from tests with rodents or other mammals. The 

highest concentration should be a level that is expected to reveal significant 

effects on adult health or reproductive parameters. If no significant effects are 

expected, then the highest concentration to be tested should be the expected 

concentration of the chemical in the environment with the addition of a safety 

factor (e.g., 5X). However, the highest dose administered should not cause 

mortality or other severe signs of parental toxicity that will preclude the 

evaluation of reproductive parameters. The lowest concentration tested should be 

chosen so that no effects are seen on adult health or reproductive parameters. 

Intermediate concentrations should be geometrically spaced between the highest 

and lowest doses. A minimum of four concentrations and a control are required 

for the definitive test.

22. A range finding test can be performed before initiating the main study to aid in 

establishing test concentrations and to evaluate any potential avoidance effects.

Preparation and monitoring of the test diet

23. To prepare the test diets, the appropriate amounts of test substance are mixed into 

the diet. The mixing method should be developed so as to obtain a homogeneous 

distribution of test substance in the diet. The use of a premix is advisable 

24. If necessary, a vehicle of negligible toxicity (e.g., food grade corn oil, water, etc.) 

may be used to ensure a uniform distribution. Acetone may be used to dissolve a 

test substance, provided it is allowed to evaporate before feeding the diet to the 

birds. The amount of vehicle used should be as low as possible and should not 

exceed 2% by weight of the diet. When used, a constant amount of vehicle 

should be added to each test group and the control group diet, in order to keep the 

caloric value of the diet equal between dosage groups.

25. The frequency of diet preparation should be chosen so that degradation or 

volatilization of the test substance in the diet does not allow the actual 

concentration to fall below 80% relative to initial concentration. The frequency 

of diet renewal should not be more than once a day and not less than once a week. 

Residual feed should be visibly inspected for mold and if acceptable, weighed and 

mixed with fresh feed supplied to the birds. It may be desirable to keep all feed 

frozen until use.
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26. Stability of the chemical in feed must be analyzed under typical test conditions 

prior to the start of the test or in parallel with the range-finding study. Stability 

data from previous (mammalian) feeding studies may be used as guidance, if 

available. Stability should be verified during the definitive reproduction test. In 

order to check stability of the test substance under test conditions, feed should be 

sampled and analyzed from the feed hoppers at the end of the first feeding period, 

i.e., before diet in the feeding bowls is renewed, and again at the end of the last 

feeding period. A sufficient number of samples should be taken into account for 

variability.

27. Homogeneity of the test substance in the diet may be evaluated prior to the test or 

samples for homogeneity measurements may be taken at the first mix prepared for 

the study.

28. To verify test concentrations, samples of diets fed to the birds should be taken 

every time new diet is mixed during the treatment period to allow measurement of 

the actual concentration of the test substance.

29. Both F0 and F1 generations are maintained on treated diet until test termination. 

All F2 chicks are maintained on untreated diet.

Measurements and observations

Adults

30. During the treatment period, the parental birds are observed daily to detect any 

overt signs of toxicity or other clinical signs. The following observations are to 

be performed on adults during the study :

• Clinical signs of toxicity and health conditions should be evaluated at least 

once daily, during the acclimation and treatment periods. Observations should 

include mortality and clinical sign of toxicity such as lethargy, depression, 

wing droop, ruffled feathers, lacrimation, feces, etc. Any injuries sustained 

and subsequent treatment should also be recorded. Clearly moribund 

individuals or those otherwise in severe distress should be immediately 

euthanized.

• Food consumption (per pair) should be recorded at least weekly as often as 

food is replaced in the feeders. Any apparent food spillage should be noted.

• Body weights should be determined at least at the start of treatment and at the 

end of the treatment period. There should be no significant difference in mean 

body weight between test groups at the start of the treatment period. If there 

is a significant difference between test groups, birds should be again 

randomized.

• Observations of secondary sex characteristics, e.g. male/rusty or 

female/mottled plumage, production of the first egg or first foam), cloacal 

length, should be recorded beginning on the 4th week and every week after 

until distinctly dimorphic traits are expressed (F0 and F1 generations). If 
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plumage is not distinctly “male” or “female”, note the plumage as 

“ambiguous”. 

Offspring – Eggs

31. Eggs are collected at least once daily, numbered according to pen of origin and 

stored; broken eggs are numbered, recorded and then discarded. The total number 

of eggs laid per pen should be accounted for at the end of the study.

32. The eggs are stored in a cold storage facility, for a maximum of 2 weeks, prior to

setting in the incubator (see Table 3). Cool storage prevents embryo development 

and aids in synchronizing development of embryos of eggs. Before placing the 

eggs in the incubator, they are candled to check for abnormalities and fine cracks 

that can be identified only by candling. After removing all cracked and abnormal 

eggs, the remaining eggs are equilibrated to room temperature and set in the 

incubator. All cracked and abnormal eggs should be recorded. 

Table 3: Conditions for egg storage, incubation and hatching

Temperature (oC) Relative humidity 

(%)

Turning

Storage 13-16 55-75 Optional

Incubation 37.5-37.8 50-70 Yes

Hatching 37-37.5 70-75 No

33. Eggs from F0 and F1 pairs will be collected during the 5th, 6th, and 7th post-pairing 

weeks. Eggs collected during the 5th week will be candled to estimate fertility and 

embryo viability on embryonic days (ED) 8 and 15, respectively. On ED 15, eggs 

will be opened and embryo weight and sex will be recorded. Tissues will be 

collected (see 43 and 44 below) from two ED 15 chicks per pen. In addition, trunk 

blood and thyroid glands will be collected for biochemical analyses from two ED 

15 chicks per pen. If egg production from the 5th week is limited (i.e. < 4 eggs 

with viable ED15 embryos/pen are produced), the priority is as follows: 1st

embryo for histology, 2nd embryo for biochemistry, 3rd embryo for histology, 4th

embryo for biochemistry. If the eggs produced during week 5 are not sufficient 

(e.g. < 3/pair) to estimate fertility, additional eggs can be collected from breeding 

pairs during weeks 8 and 9. F1 birds are maintained on treated diet until test 

termination. Eggs collected from weeks 6 and 7 will be set, artificially incubated, 

and allowed to hatch to establish the F1 generation. All F2 offspring will be 

maintained on untreated diet and survival through day 14 is recorded. 

The onset of sexual maturity (egglaying or foam production) or absence thereof 

can be recorded for F2 hatchlings. The timing of sexual maturity will be 

considered egg or foam production in 90% of the control (i.e. birds maintained on 

an untreated diet) and appropriate behavioral responses among paired birds. 

Observations will extend for an additional week beyond sexual maturity of the 

control groups to estimate potential pubertal delays among treated birds.
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During the study, one egg per pen is collected from odd number pens in odd 

numbered weeks and from even numbered pens in even numbered weeks to 

evaluate eggshell strength and thickness. If egg production is limited during the 

5th (<5 eggs), 6th, and 7th weeks (<7 eggs total for both weeks), forego egg 

collection for shell measurements. Eggshell strength is measured in Newtons 

using a strength tester. The egg is placed on its side on the test stand so the 

compression head will contact the egg at the equator. To determine eggshell 

thickness, each egg will be cut open around the equator and washed out; 

subsequently the shells are left to dry with the membrane intact for at least 48 

hours at room temperature. Shell thickness is measured at a minimum of four 

points around the girth using a calibrated micrometer. Eggshells are measured to 

at least 0.01 mm and the mean value calculated per egg. Eggs other than those 

collected for production of the next generation (post-treatment weeks 6 and 7), 

embryonic survival and fertility (post-treatment week 5, and if necessary posttreatments weeks 8 and 9), and those collected for eggshell thickness and strength 

(one egg/pen/2 weeks), can be discarded after enumeration and examination for 

cracks or abnormalities. Egg yolks from eggs broken during shell measurements 

can be used for biochemical analyses (see 46 and 47 below).

35. Eggs collected from the F0 and F1 generation during the 5th, 6th, and 7th postpairing week will be incubated to assess embryo viability (5th post-pairing week) 

and establish the next generation (6th and 7th post-pairing weeks). Eggs collected 

from the F1 generation during post-pairing weeks 1-4 will also be incubated 

through embryonic day 8 to evaluate fertility. Incubation is best performed when 

eggs are set in an incubator with temperature and humidity control and an 

automatic turning device. From the second day of incubation onwards, eggs 

should be turned at least three times per day. If turned by hand, eggs should be 

turned an odd number of times a day.

36. Fertility and embryo viability are checked by removing the eggs from the 

incubator and candling them. All incubated eggs are candled for fertility after 8 

days of incubation (ED 8) and, for only eggs collected during post-pairing weeks 

5-7, again for viability of embryos approximately 2 days before hatching (ED 15). 

Eggs from post-pairing weeks 6 and 7 will be transferred to a hatcher. During 

candling, eggs should not be allowed to cool to room temperature, since this may 

delay embryonic development. After hatching, chicks should be dry before they 

are taken out of the hatcher. Those chicks that have not hatched within 

approximately 24 hours of the majority of chicks hatching, should be considered 

unhatched. No assistance should be given to chicks during hatching. 

Egg parameters

37. Prior to start of the treatment period, egg production of all F0 generation pairs of 

birds available for the study (including potential replacements) should be 

recorded. These data will be used to exclude pairs that do not lay eggs from 
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further study. Eggs will be discarded after candling unless otherwise indicated 

(see appendix). 

38. During the treatment periods, the following information is recorded:

• Eggs laid

• Eggs cracked

• Eggs broken

• Description of all egg abnormalities

• Eggs set

• Eggshell strength

• Eggshell thickness

• Fertile eggs

• Infertile eggs

• Early viability (ED8)

• Late viability (ED15)

• Eggs that hatch

• Eggs that do not hatch

39. If a pair is removed during the study due to death, injury, etc., data from all 

preceding weeks are evaluated. If data from a pen in which mortality occurred or 

a bird was injured are used in study results, this should be justified.

40. Typical control values for reproductive parameters in Japanese quail are provided 

in Table 4. If reproductive parameters of control birds do not meet these values, 

the test procedure and husbandry conditions should be checked for potential 

problems. 

Table 4: Typical values for reproductive parameters in Japanese quail

Number of eggs laid per hen per day 0.66 – 0.89

Percentage of cracked/broken eggs 0 to 10%

Viability (percentage fertile, live embryos of eggs set) 85 to 96%

Hatchability (percentage hatching of fertile eggs) 70 to 98%

Percentage hatchlings that survive 14 days 85 to 97%

Mean number of 14-day old survivors per hen per day 0.34 – 0.71

Eggshell thickness (mm) 0.19 to 0.22

Chicks

41. The following observations on F1 and F2 chicks are recorded during the treatment 

periods:

• Day of hatch

• Hatching mortality
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• Hatchling morphology and/or abnormalities

• Chick body weight at hatching

• Clinical signs of toxicity/disease

• Number of 14-day-old surviving chicks

• Chick body weight at 14 days after hatching

• Time to reproductive maturation (F2 optional)

• Phenotypic sex

• Genetic sex (F0, F1, F2)

• Mating behavior (F2 optional)

Pathology

42. Adults - All adults that survive to the end of the treatment period are killed as 

humanely as possible. All adult birds are to undergo necropsy and gross 

pathology assessment. Adult birds that die, or are killed during the course of the 

treatment period, will be subjected to the same procedures. Trunk blood will be 

collected for biochemical analyses. The left thyroid gland is to be stored for 

glandular T4 content. Organs including brain (POM, pituitary, pineal gland), 

kidneys, liver, adrenal glands, reproductive organs and associated structures, 

cloacal gland, bursa, and right thyroid gland should preserved in situ using 

Davidson’s fixative for further histopathology. Any morphologic abnormality 

should be recorded at the time of necropsy. Fixed organ (or group of organs) 

weight will be collected when organs are removed for histopathology.

43. Embryos – For each mating pair, two late-stage embryos (ED 15, 2 days prior to 

hatch) that are not allowed to hatch (i.e. from 5th week post-treatment eggs) are to 

undergo necropsy and tissues preserved for histopathology as described above for 

adults. Trunk blood and thyroid glands will be collected from two late-stage 

embryos per pair for biochemical analyses. If a mating pair produces fewer than 

4 eggs with viable late-stage embryos, see section 34 above for sample 

prioritization. 

44. Birds (adults or chicks) which are in severe distress will be killed in extremis. If 

one member of a pair dies or is killed in extremis during the treatment period, the 

other member of the pair is killed as well. 

Biochemical/endocrine measurements and observations

45. Thyroid hormone measurements – Measurements of thyroid function include 

thyroid gland weight, glandular thyroxine (T4) content, and thyroid 

histology/histopathology. Samples will be collected from embryonic and adult 

life stages. Eggs collected from post-treatment week 5 incubated to ED 15, can 

be used for embryonic measurements. After assessing late embryo viability two 

days prior to hatch, the embryo is removed from the shell and necropsied. 

Thyroid glands are dissected. Fixed thyroid glands for histology will be collected 
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from two embryos and unfixed thyroid glands for biochemistry will be collected 

from two embryos for each male-female pair. In addition, during necropsy of the 

adult Japanese quail, thyroid glands are dissected and the left thyroid gland will 

be used for glandular T4 analysis and right gland used for histological assessment 

as above. 

46. Steroid hormone measurements - Non-invasive procedures are preferable for 

periodic measurements of estradiol and testosterone in developing birds, adults, 

and eggs. One approach for measuring steroid hormones in the developing birds 

and adults is through fecal/urate material. Steroids are to be measured from 

fecal/urate material using radioimmunoassay (RIA) starting Day 1 of adult 

pairing, and weekly thereafter. Adults are housed in pairs, and the fecal/urate 

material shall be linked to the pen. Steroid hormones shall be analyzed in trunk 

blood collected from two late-stage embryos per male-female pair, from blood 

samples collected from all animals at necropsy, and from egg yolks resulting from 

eggshell thickness/strength measures.

47. Vitellogenin measurements – Vitellogenin is a protein that is generally controlled 

by estrogen. It is present in females of oviparous species during egg 

development. It is not expressed in normal males due to lack of estrogen, 

however in males that have been exposed to estrogenic contaminants, it is 

expected that vitellogenesis will be stimulated. Vitellogenin is to be measured 

from trunk blood of the late embryos used for steroid and thyroid hormone 

analyses and from adults at necropsy, when appropriate, to evaluate estrogenic 

potential of a test chemical.

Behavioral measurements

48. Males – Initiation of normal male mating behavior is important for successful 

reproduction. Mating behavioral observations will be conducted, preferably 

beginning at the onset of maturation, on males of the F1 (F2, optional) generation. 

Trials shall be performed to assess mating behaviors including time to mount 

attempts, number of mount attempts, and number of successful cloacal contacts. 

Behavioral observations shall be conducted on naive birds raised singly or in 

same-sex pairs. At sexual maturity (i.e. egg laying in females and foam 

production in males), 7 naive male-female pairs will be introduced and behavior 

will be observed during three minute trials for three consecutive days. Trials may 

be truncated after successful mating or if bird aggression becomes an issue. At 

the conclusion of the trial, birds will be returned to their single or same-sex 

housing. The same individuals shall be re-introduced for three consecutive days

or until successful mating is observed. If behavioral observations of the F1 

generation suggest a treatment-related effect, observations may be extended to the 

F2 generation.

Handling-related disturbance during introduction and separation of birds involved 

in behavior observation trials may perturb egg production, and therefore, 
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observational data should be collected from distinct pairs not used in collection of 

other data endpoints (e.g. egg production, fertility, etc.). 

DATA COLLECTION AND REPORTING

Treatment of results

49. Numerical data should be presented in tabular form. All adult health data should 

be recorded per individual bird (food consumption per pair). Since the F0 

parental pair (or pen) is the primary statistical unit and the F1 parental pair (or 

pen) is the secondary unit, all reproductive data should be related by lineage. The 

raw data values should be reported by pen to provide sufficient detail for an 

independent statistical analysis.

50. Measurements of endpoints made on adult birds will be evaluated by comparing 

values obtained from birds in treated groups with values obtained from control 

birds. The following adult health endpoints should be analyzed for both F0 and 

F1 birds, using appropriate statistical procedures:

• Food consumption

• Body weight

• Number of eggs per hen per day

51. Moreover, mortality, clinical observations or pathological findings may be 

evaluated statistically to verify any apparent relationship between adult and 

treatment.

52. The following reproductive data (both continuous and discrete variables) are to be 

evaluated statistically by comparison with the control group using appropriate 

methods:

• Number of fertile eggs as a percentage of eggs set

• Number of early viable embryos as a percentage of eggs fertile

• Number of late viable embryos as a percentage of early viable embryos

• Number of eggs hatched as a percentage of late viable embryos

• Number of 14-day old chicks as a percentage of eggs hatched

• Number of 14-day old chicks as a percentage of eggs set

• Number of 14-day old chicks per hen per day

• Number of cracked eggs as a percentage of eggs laid 

• Number of abnormal eggs as a percentage of eggs laid

• Mean egg shell thickness 

• Mean eggshell strength

• Mean hatchling body weight

• Mean 14-day old chick body weight

• Sex ratio of chicks
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53. The following parameters may be useful to evaluate statistically, depending on the 

findings:

• Number of hatchlings per hen per day

• Number of normal hatchlings as a percentage of total hatchlings

• Number of infertile eggs as a percentage of eggs set

• Number of early embryonic deaths as a percentage of fertile eggs

• Number of late embryonic deaths as a percentage of viable eggs

54. The following endocrine and physiological endpoints (both continuous and 

discrete variables) are to be evaluated statistically by comparison with the control 

group using appropriate methods:

• Fixed weight of adult testes, ovaries, thyroid, adrenals, oviduct, cloacal gland, 

liver

• Histology of thyroid, adrenals, gonads, brain

• Testicular spermatid counts and morphology (optional)

• Gross anomalies of the genital tract

• Feather dimorphism

• Cloacal gland size, 1st appearance of foam

• 1st egg laid

• Mating behavior (time to mount, number of mount attempts, successful 

cloacal contact)

• Fecal/urate steroid hormones (estradiol, testosterone)

• Egg steroid content (estradiol, testosterone)

• Serum steroid hormones (estradiol, testosterone, vitellogenin when 

appropriate)

• Glandular thyroid hormone

55. A “no observed effect concentration” (NOEC) expressed in mg/kg diet and mg/kg 

body weight per day should be determined for all health and reproductive 

parameters evaluated.

Test report

56. The final report should include the following information:

Test substance:

• identification, including chemical name

• batch or lot number

• degree of purity

• chemical stability under the conditions of the test

• volatility
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Test species:

• name of species tested (scientific name), strain or origin, source and age of 

birds at start of test (in weeks)

Test conditions:

• housing conditions (type, size and material of pens for adults and for chicks, 

additional floor covering, adjustments made to pen floors to facilitate egg 

collection and prevent breakage, temperature, humidity, ventilation, 

illumination intensity and photoperiod, water supply); any changes to these 

during the test; measures taken to minimize food spillage

• description of the untreated diet, manufacturer, composition, caloric content of 

diet and carrier, results of periodically performed contaminant and nutrient 

analysis of diet and drinking water contaminant analysis

• test groups (number of concentrations used, nominal concentrations); details 

of type of carrier used and concentration as percentage of diet; test substance 

concentrations must be reported as mg/kg diet and as mg/kg body weight

• specific analytical method used to determine the concentrations of test 

substance in the diet as well as actual values of homogeneity, stability and 

accuracy of preparation in diet under test conditions, as recorded in the study

• description of the type and frequency of the procedure used to prepare the test 

diets; description of the manner of administrating the test diets; storage 

conditions

• description of acclimation, stabilization and pre-treatment procedures, and 

method of assigning pairs to test groups (F0 and F1); arrangement of pens; 

number of pairs per dose group; measures taken, if any, to reduce pen-mate 

aggression

• frequency, duration and methods of observation

• information on the period and conditions of storage of eggs and on the 

incubation method

• method of marking all birds and eggs

Results:

• description of incidence of mortality, indicating the number of dead animals 

and the time of death during the test

• description of all clinical signs of toxicity and other abnormal behavior, 

including time of onset, duration and severity, and number of affected birds 

per test group and in the control group; any injuries sustained and subsequent 

treatment

• macroscopic pathological findings and adult fixed organ weights and results 

of histopathological investigations

• mean body weights of F0 males and F0 females per pen at start of treatment 

and at the end of the F0 treatment period (end of week 6 of treatment); mean 
15

body weights of F1 males and F1 females per pen at pairing and at the end of 

the test; individual weights of the birds that died or were killed during the test

• weekly food consumption per test group during treatment periods and 

extrapolated mean food consumption per pair; and expressed as mg/kg body 

weight per bird per day; an indication of any apparent food spillage

• results of range-finding test (when performed)

• lowest observed effect concentration (LOEC)

• no observed effect concentration (NOEC)

Reproductive Data (tabulated separately for each generation):

• number of eggs laid per hen and per day 

• all eggs should be traceable to their pen of origin

• all egg abnormalities should be described and reported

• numbers of eggs cracked and the percentage of eggs cracked of eggs laid

• eggshell thickness, eggshell strength measurements

• number of fertile eggs and the percentage fertility related to eggs set (F0 

collection weeks 5-7; F1 collection weeks 1-7)

• number of infertile eggs and the percentage infertility related to eggs set (F0 

collection weeks 5-7; F1 collection weeks 1-7)

• number of early viable embryos and the percentage viability related to fertile 

eggs 

• number of early embryonic dead and the percentage related to fertile eggs

• number of late embryonic dead and the percentage related to 8-day viable 

embryos (collection week 5)

• number of normal hatchlings per hen per day and the percentage related to 15-

day viable embryos (collection week 5)

• number of normal hatchlings as a percentage of fertile eggs (collection weeks 

6 and 7)

• number of normal hatchlings as a percentage of eggs set (collection weeks 6 

and 7)

• number of normal hatchlings and the percentage related to total number of 

hatchlings (collection weeks 6 and 7)

• number of 14-day old chicks per hen and per day (collection weeks 6 and 7)

• number of 14-day old chicks as a percentage of (normal) hatchlings 

(collection weeks 6 and 7)

• number of 14-day old chicks as a percentage of eggs set (collection weeks 6 

and 7)

• mean body weights of chicks on the day of hatching and after 14 days per pen, 

per test group

• description of abnormal behavior of chicks, of severe birth defects and their 

general state of health, during the first 14 days after hatching, clinical signs of 

toxicity

• observations of hatchling mortality

• NOEC (and LOEC) per parameter that was evaluated statistically
16

A description of statistical methods used in the analysis of data.

LITERATURE

(1) Report of the SETAC/OECD Workshop on Avian Toxicity testing. (1996). 

Environment Health and Safety Publication - Series on Testing and Assessment: No. 5. 

OECD/GD (96)166, Paris.

(2) OECD Guidelines for the Testing of Chemicals. (1993). Section 2 - Effect on 

Biotic Systems: Test Guideline 206: Avian Reproduction Test (adopted April 1984).

(3) OECD Guidelines for Testing of Chemicals. Proposal for a new test guideline: 

Avian reproduction toxicity test in the Japanese quail or Northern bobwhite. (September 

1999 draft)

(4) Endocrine Disruptor Screening and Testing Advisory Committee: Final Report. 

(1998)
17

APPENDIX

Table 1. Sampling design for the Avian Two-Generation Toxicity Test in the Japanese 

Quail. (ED = embryonic day).

Generation

Timing F0 F1 F2

ED 8 Fertility/embryo viability

ED 15 Fertility/embryo viability

Sample collection (PT wk 5 

eggs)

Trunk blood

Thyroid gland

Histopathology

HATCH Begin treated diet Untreated diet

Week 1 Observations of secondary sex characteristics

Clinical observations (survival)

Behavioral observations

Week 2

Week 3

Week 4 Pairs established

2 week acclimation period*

Daily clinical observations

Pairs established Observations of sexual maturity 

in F2 birds (optional)

Sexual maturity

• Egg production

• Foam production

Sexual maturity

• Egg production

• Foam production

• Mating behavior in 

satellite group

Week 6 Begin treated diet*

All wks 

posttreatment

Daily clinical observations

Daily egg collections

Biweekly egg collection (shell 

thickness/strength)

Weekly food consumption

Daily clinical observations

Daily egg collections

Biweekly egg collection (shell 

thickness/strength)

Weekly food consumption

Set eggs from wks 1-4 to ED8

5 wk posttreatmentEgg collection

• Fertility/embryo data to 

ED15

Egg collection

Fertility/embryo data

to ED15

6-7 wk posttreatmentEgg collection

• Set for F1

Termination (unless extended)

• Trunk blood

• Thyroid gland

• Preserve organs

Egg collection

• Set for F2

Termination

• Trunk blood

• Thyroid gland

• Preserve organs

Sexual maturity (delayed?)‡

Termination

• Trunk blood

• Thyroid gland

• Preserve organs

8-9 wk post 

treatment†

Possible egg collection

Termination

*F0 birds are given two weeks to acclimate after sexual maturity is reached, as indicated 

by the initiation of foam production, egg laying, and mating behavior among pairs. The 

earliest treatment would occur when parental birds are 6 weeks old.

†If egg production during the 5th post-treatment week is not sufficient for evaluating 

fertility and embryonic sample collection, Parental pairs (F0) can be held for an 8th and 

9th treatment week for additional egg collection prior to termination.
18

Table 2. Sample Collection and Apical Endpoints from the Avian Two-Generation Toxicity Test in the Japanese Quail. 

Generation Levels Treatment Sample sizes Endpoints

F0 5 (control + 4 tx) > 6 wks 12 to 16 prs/level

(controls must end with 12 prs)

60 to 80 prs total

Survival

Body wt

Histopathology

Egg parameters 

Fertility 

Biochemistry

F1 5 (control + 4 tx) eggs

embryos

> hatch

Sexual 

maturity

3/pr (biweekly)

(180 to 240 total)

4/pair (2 for histo/2 for biochem)

(240-320 total)

Ideally 6 eggs/pr

(360 to 480 eggs)

Must include 40 /level

7 prs/level

(35 prs total)

Shell thickness

Shell strength

Yolk steroids

Tissue histology

Hormone biochem

Survival

Body wt

Sexual development

Histopath

Egg parameters 

Fertility 

Biochemistry

Mating behavior

F2 5 (control + 4 tx) If extended:

eggs

eggs

embryos

> hatch

Sexual 

maturity

3/pr (biweekly)

(600+ total)

All from wks 1-4

4/pair (2 for histo/2 for biochem)

Ideally 6 eggs/pr

(360 to 480 eggs)

Must include 40/level

(200 total)

7 prs/level*

Shell thickness

Shell strength

Yolk steroids

Fertility (ED 8)

Tissue histology 

Hormone biochem

Survival

Body wt

Sex ratio

Sexual development

Histopath

Egg parameters 

Fertility 

Biochemistry

Mating behavior

*Mating behaviors are observed in F2 generation only if F1 generation displays altered mating behaviors relative to 

controls.
19

Table 3. Weekly schedule of actions by generation for avian 2-generation toxicity study.

week F0 F1 F2

age

egg 

lot action age

egg 

lot action age action

1 set eggs

2

3 0 hatch

4 1

5 2 genetic sex

6 3

7 4 pair

8 5

9 6 treated diet/eggs?

10 7 1 not set

11 8 2 not set

12 9 3 not set

13 10 4 not set

14 11 5 set eggs (to ED15)

15 12 6 set eggs (F1)

16 13 7 set eggs (F1) hatch ED15 (wk 5)

17 14 terminate F0 adults 0 hatch (wk 6)/treated diet

18 1

19 2 genetic sex

20 3

21 4 pair

22 5

23 6 eggs?

24 7 1 set eggs (to ED8)

25 8 2 set eggs (to ED8)

26 9 3 set eggs (to ED8)

27 10 4 set eggs (to ED8)

28 11 5 set eggs (10 ED15)

29 12 6 set eggs (F2)

30 13 7 set eggs (F2) hatch ED15 (wk 5)

31 14 8 terminate F1 adults 0

hatch (wk 6)/untreated 

diet

32 1

33 2 genetic sex

34 3

35 4

36 5

37 6 pair for mating behavior

38 7 terminate F2