United States Prevention, Pesticides EPA712-C-96-141
Environmental Protection and Toxic Substances April 1996
Agency (7101)
&EPA Ecological Effects Test
Guidelines
OPPTS 850.2300
Avian Reproduction Test
'Public Draft"
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INTRODUCTION
This guideline is one of a series of test guidelines that have been
developed by the Office of Prevention, Pesticides and Toxic Substances,
United States Environmental Protection Agency for use in the testing of
pesticides and toxic substances, and the development of test data that must
be submitted to the Agency for review under Federal regulations.
The Office of Prevention, Pesticides and Toxic Substances (OPPTS)
has developed this guideline through a process of harmonization that
blended the testing guidance and requirements that existed in the Office
of Pollution Prevention and Toxics (OPPT) and appeared in Title 40,
Chapter I, Subchapter R of the Code of Federal Regulations (CFR), the
Office of Pesticide Programs (OPP) which appeared in publications of the
National Technical Information Service (NTIS) and the guidelines pub-
lished by the Organization for Economic Cooperation and Development
(OECD).
The purpose of harmonizing these guidelines into a single set of
OPPTS guidelines is to minimize variations among the testing procedures
that must be performed to meet the data requirements of the U. S. Environ-
mental Protection Agency under the Toxic Substances Control Act (15
U.S.C. 2601) and the Federal Insecticide, Fungicide and Rodenticide Act
(7U.S.C. I36,etseq.).
Public Draft Access Information: This draft guideline is part of a
series of related harmonized guidelines that need to be considered as a
unit. For copies: These guidelines are available electronically from the
EPA Public Access Gopher (gopher.epa.gov) under the heading "Environ-
mental Test Methods and Guidelines" or in paper by contacting the OPP
Public Docket at (703) 305-5805 or by e-mail:
guidelines@epamail.epa.gov.
To Submit Comments: Interested persons are invited to submit com-
ments. By mail: Public Docket and Freedom of Information Section, Office
of Pesticide Programs, Field Operations Division (7506C), Environmental
Protection Agency, 401 M St. SW., Washington, DC 20460. In person:
bring to: Rm. 1132, Crystal Mall #2, 1921 Jefferson Davis Highway, Ar-
lington, VA. Comments may also be submitted electronically by sending
electronic mail (e-mail) to: guidelines@epamail.epa.gov.
Final Guideline Release: This guideline is available from the U.S.
Government Printing Office, Washington, DC 20402 on The Federal Bul-
letin Board. By modem dial 202-512-1387, telnet and ftp:
fedbbs.access.gpo.gov (IP 162.140.64.19), or call 202-512-0135 for disks
or paper copies. This guideline is also available electronically in ASCII
and PDF (portable document format) from the EPA Public Access Gopher
(gopher.epa.gov) under the heading "Environmental Test Methods and
Guidelines."
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OPPTS 850.2300 Avian reproduction test.
(a) Scope—(1) Applicability. This guideline is intended to meet test-
ing requirements of both the Federal Insecticide, Fungicide, and
Rodenticide Act (FIFRA) (7 U.S.C. 136, et seq.) and the Toxic Substances
Control Act (TSCA) (15 U.S.C. 2601).
(2) Background. The source material used in developing this har-
monized OPPTS test guideline are 40 CFR 797.2130 Bobwhite Reproduc-
tion Test and 797.2150 Mallard Reproduction Test; OPP 71-4 Avian Re-
production Test (Pesticide Assessment Guidelines, Subdivision E—Hazard
Evaluation; Wildlife and Aquatic Organisms) EPA report 540/09-82-024,
1982; and OECD 206, Avian Reproduction Test.
(b) Purpose. This guideline is designed to develop data on the repro-
ductive effects on the bobwhite and mallard of chemical substances and
mixtures subject to chronic environmental effects test regulations. The
Agency will use these and other data to assess the reproductive effects
on birds that these chemicals may present to the environment. The pre-
requisite data for the study are water solubility, vapor pressure, and avian
dietary LC50 of the test substance. Guidance information on the substance
include the structural formula, purity, w-octanol/water partition coefficient,
methods for quantification of the test substance in the diet, and its stability
in water, light, and in the diet.
(c) Definitions. The definitions in section 3 of the Toxic Substances
Control Act (TSCA) and 40 CFR Part 792—Good Laboratory Practice
Standards apply to this test guideline.
(1) In addition, the following definitions apply generally to this guide-
line:
Acclimation is physiological and behavioral adaptation to environ-
mental conditions (e.g. housing and diet) associated with the test proce-
dure.
Basal diet is the untreated form of the diet, such as the diet obtained
from a commercial source.
Photoperiod is the light/dark periods in a 24 h day. This is usually
expressed in a form such as 17 h light/7 h dark or 17L/7D.
Test substance is the specific form of a chemical or mixture of chemi-
cals that is used to develop the data.
(2) The following definitions refer specifically to the production and
quality of eggs and the subsequent development of these eggs up to the
point where young are 14 days old.
Eggs cracked. Eggs determined to have cracked shells when inspected
with a candling lamp. Fine cracks cannot be detected without using a can-
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dling lamp and if undetected will bias data by adversely affecting embryo
development. Values are expressed both as number of eggs per pen per
test and as a percentage of eggs laid by all hens during the test.
Eggs laid. This term refers to the total egg production during the
test, which normally includes 10 weeks of laying. Values are expressed
as numbers of eggs per pen per season (or test).
Eggs set. All eggs placed under incubation, i.e. total eggs minus
cracked eggs and those selected for analysis of eggshell thickness. The
number of eggs set, itself, is an artificial number, but it is essential for
the statistical analysis of other development parameters.
Eggshell thickness. The thickness of the shell and the membrane of
the egg at several points around the girth after the egg has been opened,
washed out, and the shell and membrane dried for at least 48 h at room
temperature. Values are expressed as the average thickness of the several
measured points in millimeters.
14-day-old survivors. Birds that survive for 2 weeks following hatch.
Values are expressed both as a percentage of hatched eggs and as the
number per pen per season (test).
Hatchability. Embryos that mature, pip the shell, and liberate them-
selves from the eggs on day 23 or 24 of incubation for bobwhite and
days 25, 26, or 27 of incubation for mallards. Values are expressed both
as number of hatchlings per pen per test and as percentage of viable em-
bryos (fertile eggs).
Live 18-day embryos or 21-day embryso for bobwhite and mallards,
respectively. Embryos that are developing normally after 18 or 21 days
of incubation for bobwhite and mallards. This is determined by candling
the eggs. Values are expressed both as number of 18-day old quail or
21-day old duckling embryos per pen per test and as a percentage of viable
embryos (fertile eggs).
Viable embryos (fertility). Eggs in which fertilization has occurred
and embryonic development has begun. This is determined by candling
the eggs 11 days after incubation has begun. It is difficult to distinguish
between the absence of fertilization and early embryonic death. The dis-
tinction can be made by breaking out eggs that appear infertile and exam-
ining further. This distinction is especially important when a test
compound induces early embryo mortality. Values are expressed as a per-
centage of eggs set.
(d) Test procedures—(1) Summary of the test, (i) The birds should
be observed for health and acclimated for at least 2 weeks after arrival.
(ii) Test birds should be randomly assigned to control and various
treatment groups.
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(iii) The test substance should be thoroughly and evenly mixed into
the diet. All treatment levels should be analyzed for test substance con-
centrations at the beginning and midway through the test.
(iv) Birds should be weighed at the beginning of the test, at 14-day
intervals until the onset of laying, and at termination of the test.
(v) Photoperiod should be carefully controlled on a shortday basis
during the initial exposure phase, then increased to 16 to 17 h to induce
egg laying.
(vi) Birds should be observed regularly for abnormal behavior or mor-
tality throughout the test.
(vii) Eggs should be removed daily and stored until there is a suffi-
cient quantity for incubation. All eggs should be candled at day 0 for
cracks and cracked eggs removed. Once every 2 weeks, all eggs produced
that day should be analyzed for eggshell thickness. Incubated eggs should
be candled on day 11 and day 18 for bobwhite and day 14 and day 21
for mallards. Hatching should be completed by day 24.
(viii) Hatchlings should be maintained until they are 14 days old. Ab-
normal behavior or death should be reported. Hatchlings should be
weighed on day 14.
(ix) A statistical analysis should be performed, preferably by analysis
of variance or regression analysis.
(x) The report should include all conditions, procedures, and results.
Data should be sufficiently detailed for an independent statistical analysis.
(xi) All treated birds should be sacrificed and disposed of properly.
Control birds may be kept as breeding stock, but should not be used in
any other tests. Control offspring may be reared and used in another test
as adults.
(2) Definitive test—(i) Test substance. (A) Concentrations for the
test substance should be based on measured or calculated residues expected
in the diet, unless otherwise specified. The concentrations should include
an actual or expected field residue exposure level and a multiple level
such as 5. Generally, three treatment groups and a control group will be
used. The highest nonlethal level may be estimated from the avian dietary
LC50.
(B) The material to be tested should be analytically pure and the de-
gree of purity should be reported along with the percentage of each impu-
rity. The technical grade should be tested.
(ii) Controls. A concurrent control is required during every test. The
control birds should be from the same hatch as the test groups. Control
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and test birds should be kept under the same experimental conditions. The
test procedures should be the same for control and treated birds, except
that no test substance should be added to the diets of control birds. If
a carrier or diluent is used in preparation of the test diets, the same carrier
should be added to the diets of control birds in the highest concentration
used for test diets.
(iii) Test groups and numbers of birds. (A) For bobwhite, each
of the three treatment groups and the control group should consist of a
minimum of 12 replicate pens. Each pen should contain one male and
one female, or alternatively one male and two females. For mallards each
of the three treatment groups and the control group shall consist of a mini-
mum of 12 replicate pens, with each pen containing one male and one
female or with each pen containing one male and three females and with
a minimum of 8 replicates. The use of 20 replicate pens in the control
group may yield a test with greater statistical power. Either arrangement
is acceptable if productivity reaches the definitive values given in para-
graph (d)(2)(xi)(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 ex-
periment first without test substances in order to determine the feasibility
of obtaining acceptable productivity levels.
(B) All control and treatment birds should be randomly distributed
to pens from the same population.
(iv) Duration of test. (A) The test consists of three phases following
acclimation to test facilities. The initial phase begins with exposure of
treatment groups to diets containing the test substance and is typically 6
to 8 weeks long. After the initial phase, the photoperiod is manipulated
according to paragraph (d)(4)(v) of this guideline to bring the hens into
laying condition. This second phase ends with the onset of egg-laying and
is typically 2 to 4 weeks long. 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 phase if reduced reproduction is
observed. The withdrawal period, if used, need not exceed 3 weeks.
(B) Exposure of adult birds to the test substance should be continuous
throughout the test. Unless otherwise specified, test birds should be ex-
posed for at least 10 weeks prior to the onset of egg laying.
(v) Preparation for reproduction (photoperiod). (A) Lighting re-
gimes (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 defini-
tive values given in paragraph (d)(4)(xii) 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
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be controlled carefully. It is important during the initial phase to not inter-
rupt the dark period unless absolutely necessary.
(B) A suggested photoperiod regime would consist of maintaining
birds under a photoperiod for 7 or 8 h of light during the initial phase.
At the end of the initial phase, the photoperiod may be increased to 16
to 17 h of light per day. The photoperiod may be maintained at this level
for the remainder of the study, although an increase each week of 15 min/
day is acceptable.
(vi) Observations of record on adult birds. (A) Body weights
should be recorded for each adult bird at the beginning of the treatment
period, at 14-day intervals until the onset of egg laying, and at termination
of treatment. 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.
(B) Observations on adult birds should be made at least once a day.
Any mortality or other signs of toxicity should be described and recorded
by date or day of test. Gross pathological examinations should be con-
ducted on all birds that die during the test period, and for all survivors
at the end of the test. Analysis of two or more tissues (e.g. muscle, fat)
for test substance residues is encouraged, but not required unless the test
substance is persistent, bioaccumulates, or if the log P (w-octanol/water)
is higher than 3.
(vii) Egg collection, storage, and incubation. All eggs should be
collected daily, marked according to the pen from which collected, and
should be stored at 16 °C and 55 to 80 percent relative humidity. Storage
in plastic bags may improve uniformity of hatching. Stored eggs should
be turned daily. At weekly or biweekly intervals, eggs should be removed
from storage and be candled to detect eggshell cracks. Except for eggs
with cracked shells and those eggs removed for eggshell thickness meas-
urements, all eggs should be set after candling for incubation in a commer-
cial incubator. If incubators are not equipped to turn eggs automatically,
they should be turned daily by hand. During the incubation period, eggs
should be maintained at 37.5 °C and approximately 70 percent relative
humidity. Eggs should be candled again on day 11 of incubation to deter-
mine fertility and early death of embryos. A final candling should be done
on day 18 to measure embryo survival. On day 21, eggs should be re-
moved to a separate incubator or hatcher. Hatching will normally be com-
plete by the end of day 24. During hatching the temperature and relative
humidity should be 37.5 °C and 70 percent, respectively. Forced draft in-
cubators and hatchers should be used. If still-air, gravity-vented incubators
and hatchers are used, temperatures should be 1.5-2 °C higher, and the
relative humidity should be increased by about 10 percent. At higher ele-
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vations, higher relative humidity is necessary. Temperatures in brooder
pens should be measured at 2.5 to 4 cm above the pen floor.
(viii) Chick or duckling maintenance. By day 24 or 27 of incuba-
tion, 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 individ-
ually marked (such as by leg bands) as to parental group and housed to-
gether. Chicks or ducklings should be maintained in commercial brooder
pens or pens of similar construction. Pens should be constructed of galva-
nized metal or stainless steel. Temperature in the pens should be con-
trolled, preferably by a thermostatically controlled device. A temperature
gradient in the pen from approximately 35 °C to approximately 22 °C
will allow young birds to seek a proper temperature. Temperature require-
ments for young birds typically decline over this range from birth through
the first several weeks of life. Chicks or ducklings should be provided
a standard commercial game bird starter ration, or its nutritional equiva-
lent. No test substance may be added to the diet of chicks or ducklings.
Chicks or ducklings should be maintained until they are 14 days old.
Lighting should be on a diurnal basis (e.g. 14 h of light, 10 h of dark,
with a 15-30 min transition at dawn and dusk, but other lighting regimes
are acceptable.
(ix) Observations of record on chicks or ducklings. The number
of embryos that mature, embryos that pip shell, and embryos that liberate
themselves, hatchability, percentage of normal hatchlings, percentage of
14-day old survivors, and number of 14-day old survivors per hen should
be recorded and reported. 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. Average body
weights should be recorded for chicks or ducklings at day 14.
(x) Eggshell thickness. 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 h. The thickness of the shell plus the dried membrane should be meas-
ured at a minimum of 3 points around the girth using a micrometer cali-
brated at least to 0.01 mm units.
(xi) Typical observed values. The values reported here represent
those observed from a few testing facilities under their conditions. These
values are not necessarily representative of those from all facilities, but
if a reproduction test does not meet or at least approach these values for
control birds, then there is likely to be a problem with test procedures
or conditions that should be investigated and corrected.
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(A) Eggs laid. Normal values for both bobwhite and mallards—28
to 38 eggs per hen per season.
(B) Eggs cracked. Normal values for bobwhite—0.6 to 2.0 percent
of eggs laid. Normal values for mallards—0.6 to 6 percent of eggs laid.
(C) Viable embryos (fertility). Normal fertility values for bob-
white—75 to 90 percent of eggs set. Normal fertility value for mallards—
85 to 98 percent of eggs set.
(D) Live 18-day embryos. Normal values for bobwhite and mal-
lards—97 to 99 percent of viable embryos.
(E) Hatchability. Normal values for bobwhite and mallards—50 to
90 percent of viable embryos (fertile eggs).
(F) 14-day-old survivors. Normal values for bobwhite—75 to 90
percent of eggs hatched. Normal value for mallards—94 to 99 percent of
eggs hatched.
(G) Eggshell thickness. Normal average values for bobwhite—0.19
to 0.24 mm. Normal values for mallards—0.34 to 0.39 mm.
(xii) Definitive test criteria. (A) A test is unacceptable if bobwhite
chick or mallard duckling productivity in control groups does not average
12 or 10, respectively, 14-day old survivors per pen over a 10-week pe-
riod.
(B) A test is unacceptable if the average eggshell thickness in control
groups is less than 0.19 mm for bobwhite and 0.34 mm for mallards.
(C) A test is unacceptable if more than 10 percent of the adult control
birds die during the test.
(3) Analytical measurements—(i) Statistical analysis. Experimental
groups should be individually compared to the control group by analysis
of variance. Other accepted statistical methods may be used as long as
they are documented. In particular, regression analysis is highly desirable
if the data and number of dose levels allow the use of this technique.
Sample units are the individual pens within each treatment level or control.
Analysis should include:
(A) Body weight of adults.
(B) Food consumption of adults.
(C) Percentage of hens laying eggs. This should always be determined
when pens contain a single pair; if feasible, it should be determined when
pens contain groups.
(D) Number of eggs laid per pen.
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(E) Percent and number of cracked eggs.
(F) Percent viable embryos and number of eggs set.
(G) Percent live 18-day embryos of viable embryos.
(H) Percent and number hatching of viable embryos.
(I) Percent and number of hatchlings that are normal.
(J) Percent 14-day-old survivors of normal hatchlings.
(K) Number of 14-day-old survivors per hen.
(L) Body weights of 14-day-old survivors.
(M) Eggshell thickness.
(ii) Test substance concentrations. (A) Samples of treated diets
should be analyzed to confirm proper dietary concentrations of the test
substance. If samples cannot be analyzed immediately, they should be
stored appropriately (e.g. frozen at a temperature of • 15 °C or lower)
until analysis can be performed. Analyses should be conducted on all test
substance concentrations at the beginning of the test period and again 10
to 12 weeks later. If not otherwise available, data should be generated
to indicate whether or not the test substance degrades or volatilizes. If
the test substance is known or found to be volatile or labile to the extent
that 20 percent or more loss occurs within 1 week, then test substance
diets should be prepared (freshly or from frozen concentrate) at a fre-
quency that will prevent more than 20 percent loss of test substance.
(B) The assay method used to determine actual concentrations should
be reported according to paragraph (f)(l)(vi) of this guideline.
(iii) Analysis of basal diet. A nutrient analysis of the basal diet
should be included with the test report. For commercially prepared basal
diets, the list of ingredients supplied by the manufacturer is normally suffi-
cient if it is detailed. The composition of any vitamin or other supplements
should also be reported.
(e) Test conditions—(1) Test species—(i) Selection. (A) The bob-
white, Colinus virginianus (L.), or the mallard, Anas platyrhynchos (L.),
is the test species. Test birds should be pen-reared. They may be reared
in the laboratory or purchased from commercial breeders. Rearing stock
and/or tests birds should be obtained only from sources that have met the
requirements for "U.S. Pullorum-Typhoid Clean" classification under
paragraph (g)(l) of this guideline. Birds should be obtained only from
sources whose colonies have known breeding histories. If possible, a his-
tory of rearing practices for test birds should be obtained and made avail-
able upon request. This history should include lighting practices during
rearing, disease record, drug and any other medication administered, and
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exact age. Test birds should be phenotypically indistinguishable from wild
stock. Conscientious breeders of such birds will periodically outbreed their
flocks with genetically wild stock in order to maintain a genetic composi-
tion that approximates the heterogeneity of naturally occurring birds.
(B) All control and experimental birds used in a test should be from
the same source and strain. If shipped, all birds should be examined fol-
lowing shipment for possible physical injury that may have occurred in
transit. All birds should have a health observation period of at least 2
weeks prior to selection for treatment. Birds should be in apparent good
health. Deformed, abnormal, sick, or injured birds should not be used. A
population of birds should not be used if more than 3 percent of either
sex become debilitated during the health observation period. Birds should
not have been selected in any way for resistance to toxic substances. Birds
should not have been used in a previous test, either in a control or treat-
ment group. Offspring of birds used in a treatment group in a previous
test should not be used, but offspring of birds used as a control in a pre-
vious test are acceptable.
(C) Tests birds should be approaching their first breeding season and
should be at least 7 months old. All test birds should be the same age
within 1 month. The age of test birds should be reported.
(D) Bobwhite 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. Birds should be randomly assigned to
treatment and control pens. However, when birds in a pen are incompat-
ible, they may be rearranged within a control or treatment group at any
time prior to initiating treatment.
(E) During holding, acclimation, and testing, birds should be shielded
from excessive noise, activity, or other disturbance. Birds should be han-
dled only as much as is necessary to conform to test procedures.
(ii) Diet—(A) Adult birds. (7) A standard commercial game bird
breeder ration, or its nutritional equivalent, should be used for diet prepara-
tion. This ration or basal diet should be used for both control and treatment
birds and should be constant throughout the duration of the study. Anti-
biotics or other medication should not be used in the diet or water of
breeding birds. It may not be possible to obtain food that is completely
free of pesticides, heavy metals, and other contaminants. However, diets
should be analyzed periodically for these substances and should be selected
to be as free from contaminants as possible. A nutrient analysis (quan-
titative list of ingredients) of the diet should be included with the test
report.
(2) The test substance should be mixed into the diet in a manner
that will ensure even distribution of the test substance throughout the diet.
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If possible, the test substance should be added to the diet without the use
of a carrier or diluent. If a diluent is needed, the preferred diluent is dis-
tilled 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 com-
pletely evaporated prior to feeding. Other acceptable diluents may be used,
if necessary, and include table grade corn oil, propylene glycol, and gum
arabic (acacia). If a diluent is used, it should comprise no more than 2
percent by weight of the treated diet, and an equivalent amount of diluent
should be added to control diets.
(3) Diets may be mixed by commercial or mechanical food mixers.
Other means are acceptable as long as they result in even distribution of
the test substance throughout the diet. Screening of the basal diet before
mixing is suggested to remove large particles. For many tests 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 con-
centrate. 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 other-
wise specified or determined by degradation or volatility studies, it is rec-
ommended that final diets be prepared weekly, either fresh or from a con-
centrate. 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 percent. Analysis of diets for test sub-
stance concentration is required as specified in paragraph (e)(l)(ii) of this
guideline.
(4) Clean water should be available ad libitum. Water bottles or auto-
matic watering devices are recommended. If water pans or bowls are used,
water should be changed daily or more often.
(B) Young birds. Young birds produced during the test should be
fed a commercial game bird starter ration, or its nutritional equivalent.
No test substance should be added to the diets of young birds. No anti-
biotics or medication may be used in the diet. Bacitracin, or one of its
forms, may be added to the drinking water of young birds, if necessary.
(2) Facilities, (i) Bobwhite should be housed in breeding pens or
cages of adequate size conforming to good husbandry practices. Space re-
quirements for bobwhite have not been well defined, but it is rec-
ommended that there be at least 5,000 cm2 (approximately 2.3 ft2) of floor
space per bird. Space requirements for mallards have not been well de-
fined, but it is recommended that there be at least 10,000 cm2 (approxi-
mately 5.4 ft2) of floor space per bird. Documentation that reproductive
10
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parameters and health of birds are not adversely affected should be pro-
vided for cages much smaller than this area. The preferred construction
materials are stainless steel, galvanized sheeting, and wire mesh. For en-
closed cages, floors and external walls may be wire mesh; ceilings and
common walls should be solid sheeting. Wire mesh for floors should be
fine enough so as to not interfere with normal movement of the birds.
Open-topped pens may be constructed of the same materials for the side
walls with open tops and wire mesh or concrete floors. Concrete floors
should be covered with litter such as straw, wood shavings, or sawdust.
Other construction materials, except wood, are acceptable if they can be
kept clean. Wood may be used as vertical framing posts for the support
of wire mesh or for horizontal framing along the top of the pen. Wood
should not be used for floors or lower sides of pens unless it has been
coated with a nonadsorbent material such as perfluorocarbon plastic (e.g.
Teflon), or unless the wood is replaced between tests.
(ii) Pens should be disassembled (if feasible) and should be cleaned
thoroughly between tests. Steam cleaning of enclosed cages is rec-
ommended. Enclosed cages may be brushed thoroughly, as an alternative
method. For open-topped pens, the sides and vertical supports should be
thoroughly brushed. Any used floor litter should be discarded. The floor
composition will dictate methods used to clean the floor. If litter is used
on the floor, it should be fresh and clean when birds are placed in the
pen. The use of detergents or bleach is acceptable, but other chemical
disinfectants (such as quaternary ammonium compounds) should not be
used. When necessary to control disease vectors, hot or cold sterilization
techniques are recommended, as long as such techniques will not leave
chemical residues on the cages. For cold sterilization, ethylene oxide is
recommended.
(iii) Pens should be kept indoors in order to better control lighting,
temperature, humidity, and other factors. Outdoor pens may be used only
during the normal breeding season. The photoperiod should be carefully
controlled, preferably by automatic timers. A 15 to 30 min transition pe-
riod is desirable. The photoperiod regime is described under test proce-
dures under paragraph (d)(2)(v) of this guideline. Lights should emit a
spectrum simulating that of daylight. The use of shorter wave-length
"cool-white" fluorescent lights that do not emit the daylight spectrum
should be avoided. Illumination intensity should be about 6 fc at the level
of the birds.
(iv) Temperature and humidity should be controlled during the study.
Recommended levels are 21 °C and 55 percent relative humidity. Tem-
perature should be recorded at least weekly at the same time of day and
should be reported. For tests conducted without temperature control, tem-
perature 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 cm above the floor of the cage. Recording
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of approximate humidity levels is also desirable. Good ventilation should
be maintained. Suggested ventilation rates are 4 changes per hour in winter
and 15 changes per hour in summer.
(v) If facilities are being used for the first time, it may be desirable
to allow birds to breed in the facility prior to testing in order to ensure
that controls will have acceptable productivity according to the require-
ments given in paragraph (d)(2)(xi) and (xii) of this guideline.
(f) Reporting. (1) The test report should include the following infor-
mation:
(i) Name of test, sponsor, test laboratory and location, principal
investigator(s), and actual dates of beginning and end of test.
(ii) Name of species tested (including scientific name), age of birds
(in months) at the beginning of the test, source of birds, and body weights
for adult birds throughout the test.
(iii) Description of housing conditions, including type, size, and mate-
rial of pen, temperature, humidity, photoperiod and lighting intensity, and
any changes during the test.
(iv) Detailed description of the basal diet, including source, composi-
tion, diluents (if used), and supplements (if used). A nutrient analysis of
the basal diet should be included.
(v) Detailed description of the test substance including its chemical
name(s), source, lot number, composition (identity of major ingredients
and impurities), and known physical and chemical properties pertinent to
the test (e.g. solubility, volatility, degradation rate, etc.).
(vi) The number of concentrations used, nominal and measured con-
centrations of test substance in each level, assay method used to determine
actual concentrations, storage conditions and stability of treated diets, num-
ber of birds per pen and number of replicate pens per concentration and
for controls.
(vii) Acclimation procedures and methods of assigning birds to test
pens, including method of randomization.
(viii) Frequency, duration, and methods of observation.
(ix) Description of any signs of intoxication or any other abnormal
behavior, including time of onset, duration, severity (including death), and
numbers affected (including accidental deaths or injuries), and any remis-
sions.
(x) Food consumption per pen and any observations of repellancy
or food palatability.
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(xi) Method of marking all birds and eggs.
(xii) Details of necropsies.
(xiii) Egg and hatching data in summary and by pen per week in
sufficient detail to allow an independent statistical analysis. Data should
be presented for all of the parameters listed in paragraph (d)(3)(i) of this
guideline. The number of eggs set should also be reported.
(xiv) Egg storage, incubation, and hatching temperatures, relative hu-
midities, and turning frequencies.
(xv) Observations of health and weights of young at 14 days of age;
food consumption of young—first and second week after hatching.
(xvi) Location of all raw data storage.
(xvii) Methods of statistical analysis, interpretation of results (includ-
ing NOEC and any statistically significant effect levels), and sufficient
data to verify calculation of statistical values.
(xviii) Results of residue analysis (if performed).
(xix) Test parameters are to be reported using the metric system.
(xx) Names of toxicants (if any) used as reference substances and
method of preparation of test concentration.
(xxi) Anything unusual about the test, any deviation from these proce-
dures, and any other relevant information.
(2) In addition, the following information should be available upon
request:
(i) A general description of the support facilities.
(ii) A description of the quality control/quality assurance program,
including the average quality level (AQL) for the program element per-
forming the test, procedures used, and documentations that these levels
have been achieved.
(iii) The names, qualifications, and experience of personnel working
in the program element performing the test, including the study director,
principal investigator, quality assurance officer, as well as other personnel
involved in the study.
(iv) Standard operating procedures for all phases of the test and equip-
ment involved in the test.
(v) Sources of all supplies and equipment involved in the test.
(vi) Diagram of the test layout.
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(vii) Originals or exact copies of all raw data generated in performing
the test.
(viii) A detailed description, with references, of all statistical methods.
(g) References. For additional background information on this test
guideline the following references should be consulted:
(1) U.S. Department of Agriculture. National Poultry Improvement
Plan, Report No. 2., in Directory of Participants Handling Waterfowl, Ex-
hibition Poultry, and Game Birds. USDA, Science and Education Admin-
istration, Beltsville, MD 20705 (1979).
(2) [Reserved]
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