Official Journal of the European Union

L 247/1


of 15 July 2014

amending, for the purpose of its adaptation to technical progress, Regulation (EC) No 440/2008 laying down test methods pursuant to Regulation (EC) No 1907/2006 of the European Parliament and of the Council on the Registration, Evaluation, Authorisation and Restriction of Chemicals (REACH)

(Text with EEA relevance)


Having regard to the Treaty on the Functioning of the European Union,

Having regard to Regulation (EC) No 1907/2006 of the European Parliament and of the Council of 18 December 2006 concerning the Registration, Evaluation, Authorisation and Restriction of Chemicals (REACH), establishing a European Chemicals Agency, amending Directive 1999/45/EC and repealing Council Regulation (EEC) No 793/93 and Commission Regulation (EC) No 1488/94 as well as Council Directive 76/769/EEC and Commission Directives 91/155/EEC, 93/67/EEC, 93/105/EC and 2000/21/EC (1), and in particular Article 13(2) thereof,



Commission Regulation (EC) No 440/2008 (2) contains the test methods for the purposes of the determination of the physico-chemical properties, toxicity and eco-toxicity of chemicals to be applied for the purposes of Regulation (EC) No 1907/2006.


It is necessary to update Regulation (EC) No 440/2008 to include with priority new and updated test methods recently adopted by the OECD in order to take into account technical progress, and to ensure the reduction of the number of animals to be used for experimental purposes, in accordance with Directive 2010/63/EU of the European Parliament and of the Council (3). Stakeholders have been consulted on this draft.


This adaptation to technical progress contains six new test methods for the determination of toxicity and other health effects including a developmental neurotoxicity study, an extended one-generation reproductive toxicity study, an transgenic rodent in vivo gene mutation assay, an in vitro test to assess effects on the synthesis of steroid hormones, as well as two in vivo methods to assess oestrogenic and (anti)androgenic effects.


Regulation (EC) No 440/2008 should therefore be amended accordingly.


The measures provided for in this Regulation are in accordance with the opinion of the Committee established under Article 133 of Regulation (EC) No 1907/2006,


Article 1

The Annex to Regulation (EC) No 440/2008 is amended in accordance with the Annex to this Regulation.

Article 2

This Regulation shall enter into force on the third day following that of its publication in the Official Journal of the European Union.

This Regulation shall be binding in its entirety and directly applicable in all Member States.

Done at Brussels, 15 July 2014.

For the Commission

The President

José Manuel BARROSO

(1)   OJ L 396, 30.12.2006, p. 1.

(2)  Commission Regulation (EC) No 440/2008 of 30 May 2008 laying down test methods pursuant to Regulation (EC) No 1907/2006 of the European Parliament and of the Council on the Registration, Evaluation, Authorisation and Restriction of Chemicals (REACH) (OJ L 142, 31.5.2008, p. 1).

(3)  Directive 2010/63/EU of the European Parliament and of the Council of 22 September 2010 on the protection of animals used for scientific purposes (OJ L 276, 20.10.2010, p. 33).


The Annex to Regulation (EC) No 440/2008 is amended as follows:

Chapters B.53, B.54, B.55, B.56, B.57 and B.58 are inserted:



1.   This test method is equivalent to OECD Test Guideline (TG) 426 (2007). In Copenhagen in June 1995, an OECD Working Group on Reproduction and Developmental Toxicity discussed the need to update existing OECD test guidelines for reproduction and developmental toxicity, and the development of new guidelines for endpoints not yet covered (1). The working group recommended that a test guideline for developmental neurotoxicity should be written based on a US EPA guideline, which has since been revised (2). In June 1996, a second consultation meeting was held in Copenhagen to provide the Secretariat with guidance on the outline of a new test guideline on developmental neurotoxicity, including the major elements, e.g. details concerning choice of animal species, dosing period, testing period, endpoints to be assessed, and criteria for evaluating results. A US neurotoxicity risk assessment guideline was published in 1998 (3). An OECD Expert Consultation Meeting and an ILSI Risk Science Institute Workshop were held back-to-back in October 2000 and an expert consultation meeting was held in Tokyo 2005. These meetings were held to discuss the scientific and technical issues related to the current test guideline and the recommendations from the meetings (4)(5)(6)(7) were considered in the development of this test method. Additional information on the conduct, interpretation and terminology used for this test method can be found in OECD Guidance Documents No 43 on “Reproductive Toxicity Testing and Assessment” (8) and No 20 on “Neurotoxicity Testing” (9).


2.   A number of chemicals is known to produce developmental neurotoxic effects in humans and other species (10)(11)(12)(13). Determination of the potential for developmental neurotoxicity may be needed to assess and evaluate the toxic characteristics of a chemical. Developmental neurotoxicity studies are designed to provide data, including dose-response characterisations, on the potential functional and morphological effects on the developing nervous system of the offspring that may arise from exposure in utero and during early life.

3.   A developmental neurotoxicity study can be conducted as a separate study, incorporated into a reproductive toxicity and/or adult neurotoxicity study (e.g. test methods B.34 (14), B.35 (15), B.43 (16)), or added onto a prenatal developmental toxicity study (e.g. test method B.31 (17)). When the developmental neurotoxicity study is incorporated within or attached to another study, it is imperative to preserve the integrity of both study types. All testing should comply with applicable legislation or government and institutional guidelines for the use of laboratory animals in research (e.g. 18).

4.   The testing laboratory should consider all available information on the test chemical prior to conducting the study. Such information will include the identity and structure of the chemical; its physico-chemical properties; the results of any other in vitro or in vivo toxicity tests on the chemical; toxicological data on structurally related chemicals; and the anticipated use(s) of the chemical. This information is necessary to satisfy all concerned that the test is relevant for the protection of human health, and will help in the selection of an appropriate starting dose.


5.   The test chemical is administered to animals during gestation and lactation. Dams are tested to assess effects in pregnant and lactating females and may also provide comparative information (dams versus offspring). Offspring are randomly selected from within litters for neurotoxicity evaluation. The evaluation consists of observations to detect gross neurologic and behavioural abnormalities, including the assessment of physical development, behavioural ontogeny, motor activity, motor and sensory function, and learning and memory; and the evaluation of brain weights and neuropathology during postnatal development and adulthood.

6.   When the test method is conducted as a separate study, additional available animals in each group could be used for specific neurobehavioral, neuropathological, neurochemical or electrophysiological procedures that may supplement the data obtained from the examinations recommended by this test method (16)(19)(20)(21). The supplemental procedures can be particularly useful when empirical observation, anticipated effects, or mechanism/mode-of-action indicate a specific type of neurotoxicity. These supplemental procedures may be used in the dams as well as in the pups. In addition, ex vivo or in vitro procedures may also be used, as long as these procedures do not alter the integrity of the in vivo procedures.


Selection of animal species

7.   The preferred test species is the rat; other species can be used when appropriate. Note, however, the gestational and postnatal days specified in this test method are specific to commonly used strains of rats, and comparable days should be selected if a different species or unusual strain is used. The use of another species should be justified based on toxicological, pharmacokinetic, and/or other data. Justification should include availability of species-specific postnatal neurobehavioral and neuropathological assessments. If there was an earlier test that raised concerns, the species/strain that raised a concern should be considered. Because of the differing performance attributes of different rat strains, there should be evidence that the strain selected for use has adequate fecundity and responsiveness. The reliability and sensitivity of other species to detect developmental neurotoxicity should be documented.

Housing and feeding conditions

8.   The temperature in the experimental animal room should be 22 ± 3 °C. Although the relative humidity should be at least 30 % and preferably not exceed 70 % other than during room cleaning, the aim should be 50-60 %. Lighting should be artificial, the sequence being 12 hours light, 12 hours dark. It is also possible to reverse the light cycle prior to mating and for the duration of the study, in order to perform the assessments of functional and behavioural endpoints during the dark period (under red light), i.e. during the time the animals are normally active (22). Any changes in the light-dark cycle should include adequate acclimation time to allow animals to adapt to the new cycle. For feeding, conventional laboratory diets may be used with an unlimited supply of drinking water. The type of food and water should be reported and both should be analysed for contaminants.

9.   Animals may be housed individually or be caged in small groups of the same sex. Mating procedures should be carried out in cages suitable for the purpose. After evidence of copulation or no later than day 15 of pregnancy, mated animals should be caged separately in delivery or maternity cages. Cages should be arranged in such a way that possible effects due to cage placement are minimised. Mated females should be provided with appropriate and defined nesting materials when parturition is near. It is well known that inappropriate handling or stress during pregnancy can result in adverse outcomes, including prenatal loss and altered foetal and postnatal development. To guard against foetal loss from factors which are not treatment-related, animals should be carefully handled during pregnancy, and stress from outside factors such as excessive outside noise should be avoided.

Preparation of the animals

10.   Healthy animals should be used, which have been acclimated to laboratory conditions and have not been subjected to previous experimental procedures, unless the study is incorporated in another study (see paragraph 3). The test animals should be characterised as to species, strain, source, sex, weight and age. Each animal should be assigned and marked with a unique identification number. The animals of all test groups should, as nearly as practicable, be of uniform weight and age, and should be within the normal range of the species and strain under study. Young adult nulliparous female animals should be used at each dose level. Siblings should not be mated, and care should be taken to ensure this. Gestation Day (GD) 0 is the day on which a vaginal plug and/or sperm are observed. Adequate acclimation time (e.g. 2-3 days) should be allowed when purchasing time-pregnant animals from a supplier. Mated females should be assigned in an unbiased way to the control and treatment groups, and as far as possible, they should be evenly distributed among the groups (e.g. a stratified random procedure is recommended to provide even distribution among all groups, such as that based on body weight). Females inseminated by the same male should be equalised across groups.


Number and sex of animals

11.   Each test and control group should contain a sufficient number of pregnant females to be exposed to the test chemical to ensure that an adequate number of offspring are produced for neurotoxicity evaluation. A total of 20 litters are recommended at each dose level. Replicate and staggered-group dosing designs are allowed if total numbers of litters per group are achieved, and appropriate statistical models are used to account for replicates.

12.   On or before postnatal day (PND) 4 (day of delivery is PND 0), the size of each litter should be adjusted by eliminating extra pups by random selection to yield a uniform litter size for all litters (23). The litter size should not exceed the average litter size for the strain of rodents used (8-12). The litter should have, as nearly as possible, equal numbers of male and female pups. Selective elimination of pups, e.g. based upon body weight, is not appropriate. After standardisation of litters (culling) and prior to further testing of functional endpoints, individual pups that are scheduled for pre-weaning or post-weaning testing should be identified uniquely, using any suitable humane method for pup identification (e.g. 24).

Assignment of animals for functional and behavioural tests, brain weights, and neuropathological evaluations

13.   The test method allows various approaches with respect to the assignment of animals exposed in utero and through lactation to functional and behavioural tests, sexual maturation, brain weight determination, and neuropathological evaluation (25). Other tests of neurobehavioral function (e.g. social behaviour), neurochemistry or neuropathology can be added on a case-by-case basis, as long as the integrity of the original required tests are not compromised.

14.   Pups are selected from each dose group and assigned for endpoint assessments on or after PND 4. Selection of pups should be performed so that to the extent possible both sexes from each litter in each dose group are equally represented in all tests. For motor activity testing the same pair of male and female pups should be tested at all pre-weaning ages (see paragraph 35). For all other tests the same or separate pairs of male and female animals may be assigned to different behavioural tests. Different pups may need to be assigned to weanling versus adult tests of cognitive function in order to avoid confounding the effects of age and prior training on these measurements (26)(27). At weaning (PND 21), pups not selected for testing can be disposed of humanely. Any alterations in pup assignments should be reported. The statistical unit of measure should be the litter (or dam) and not the pup.

15.   There are different ways to assign pups to the pre-weaning and post-weaning examinations, cognitive tests, pathological examinations, etc., (see Figure 1 for general design and Appendix 1 for examples of assignment). Recommended minimum numbers of animals in each dose group for pre-weaning and post-weaning examinations are as follows:

Clinical observations and bodyweight

All animals

Detailed clinical observations

20/sex (1/sex/litter)

Brain weight (post fixation) PND 11-22

10/sex (1/litter)

Brain weight (unfixed) ~ PND 70

10/sex (1/litter)

Neuropathology (immersion or perfusion fixation) PND 11-22

10/sex (1/litter)

Neuropathology (perfusion fixation) PND ~ 70

10/sex (1/litter)

Sexual maturation

20/sex (1/sex/litter)

Other developmental landmarks (optional)

All animals

Behavioural ontogeny

20/sex (1/sex/litter)

Motor activity

20/sex (1/sex/litter)

Motor and sensory function

20/sex (1/sex/litter)

Learning and memory

10/sex (1) (1/litter)


16.   At least three dose levels and a concurrent control should be used. The dose levels should be spaced to produce a gradation of toxic effects. Unless limited by the physico-chemical nature or biological properties of the chemical, the highest dose level should be chosen with the aim to induce some maternal toxicity (e.g. clinical signs, decreased body weight gain (not more than 10 %) and/or evidence of dose-limiting toxicity in a target organ). The high dose may be limited to 1 000 mg/kg/day body weight, with some exceptions. For example, expected human exposure may indicate the need for a higher dose level to be used. Alternatively, pilot studies or preliminary range-finding studies should be performed to determine the highest dosage to be used which should produce a minimal degree of maternal toxicity. If the test chemical has been shown to be developmentally toxic either in a standard developmental toxicity study or in a pilot study, the highest dose level should be the maximum dose which will not induce excessive offspring toxicity, or in utero or neonatal death or malformations, sufficient to preclude a meaningful evaluation of neurotoxicity. The lowest dose level should aim to not produce any evidence of either maternal or developmental toxicity including neurotoxicity. A descending sequence of dose levels should be selected with a view to demonstrating any dose-related response and a No-Observed-Adverse Effect Level (NOAEL), or doses near the limit of detection that would allow the determination of a benchmark dose. Two- to four-fold intervals are frequently optimal for setting the descending dose levels, and the addition of a fourth dose group is often preferable to using very large intervals (e.g. more than a factor of 10) between dosages.

17.   Dose levels should be selected taking into account all existing toxicity data as well as additional information on metabolism and toxicokinetics of the test chemical or related materials. This information may also assist in demonstrating the adequacy of the dosing regimen. Direct dosing of pups should be considered based on exposure and pharmacokinetic information (28)(29). Careful consideration of benefits and disadvantages should be made prior to conducting direct dosing studies (30).

18.   The concurrent control group should be a sham-treated control group or a vehicle-control group if a vehicle is used in administering the test chemical. All animals should normally be administered the same volume of either test chemical or vehicle on a body weight basis. If a vehicle or other additive is used to facilitate dosing, consideration should be given to the following characteristics: effects on the absorption, distribution, metabolism, or retention of the test chemical; effects on the chemical properties of the test chemical which may alter its toxic characteristics; and effects on the food or water consumption or the nutritional status of the animals. The vehicle should not cause effects that could interfere with the interpretation of the study neither should it be neurobehaviourally toxic nor have effects on reproduction or development. For novel vehicles, a sham-treated control group should be included in addition to a vehicle control group. Animals in the control group(s) should be handled in an identical manner to test group animals.

Administration of doses

19.   The test chemical or vehicle should be administered by the route most relevant to potential human exposure, and based on available metabolism and distribution information in the test animals. The route of administration will generally be oral (e.g.gavage, dietary, via drinking water), but other routes (e.g. dermal, inhalation) may be used depending on the characteristics and anticipated or known human exposure routes (further guidance is provided in the Guidance Document 43(8)). Justification should be provided for the route of administration chosen. The test chemical should be administered at approximately the same time every day.

20.   The dose administered to each animal should normally be based on the most recent individual body weight determination. However, caution should be exercised when adjusting the doses during the last third of pregnancy. If excess toxicity is noted in the treated dams, those animals should be humanely killed.

21.   The test chemical or vehicle should, as a minimum, be administered daily to mated females from the time of implantation (GD 6) throughout lactation (PND 21), so that the pups are exposed to the test chemical during pre- and postnatal neurological development. The age at which dosing starts, and the duration and frequency of dosing, may be adjusted if evidence supports an experimental design more relevant to human exposures. Dosing durations should be adjusted for other species to ensure exposure during all early periods of brain development (i.e. equivalent to prenatal and early postnatal human brain growth). Dosing may begin from the initiation of pregnancy (GD 0) although consideration should be given to the potential of the test chemical to cause pre-implantation loss. Administration beginning at GD 6 would avoid this risk, but the developmental stages between GD 0 and 6 would not be treated. When a laboratory purchases time-mated animals, it is impractical to begin dosing at GD 0, and thus GD 6 would be a good starting day. The testing laboratory should set the dosing regimen according to relevant information about the effects of the test chemical, prior experience, and logistical considerations; this may include extension of dosing past weaning. Dosing should not occur on the day of parturition in those animals which have not completely delivered their offspring. In general, it is assumed that exposure of the pups will occur through the maternal milk; however, direct dosing of pups should be considered in those cases where there is a lack of evidence of continued exposure to offspring. Evidence of continuous exposure can be retrieved from e.g. pharmacokinetic information, offspring toxicity or changes in bio-markers (28).


Observations on dams

22.   All dams should be carefully observed at least once daily with respect to their health condition, including morbidity and mortality.

23.   During the treatment and observation periods, more detailed clinical observations should be conducted periodically (at least twice during the gestational dosing period and twice during the lactational dosing period) using at least 10 dams per dose level. The animals should be observed outside the home cage by trained technicians who are unaware of the animals' treatment, using standardised procedures to minimise animal stress and observer bias, and maximise inter-observer reliability. Where possible, it is advisable that the observations in a given study be made by the same technician.

24.   The presence of observed signs should be recorded. Whenever feasible, the magnitude of the observed signs should also be recorded. Clinical observations should include, but not be limited to, changes in skin, fur, eyes, mucous membranes, occurrence of secretions, and autonomic activity (e.g. lacrimation, piloerection, pupil size, unusual respiratory pattern and/or mouth breathing, and any unusual signs of urination or defecation).

25.   Any unusual responses with respect to body position, activity level (e.g. decreased or increased exploration of the standard area) and co-ordination of movement should also be noted. Changes in gait, (e.g. waddling, ataxia), posture (e.g. hunched-back) and reactivity to handling, placing or other environmental stimuli, as well as the presence of clonic or tonic movements, convulsions, tremors, stereotypies (e.g.excessive grooming, unusual head movements, repetitive circling), bizarre behaviour (e.g. biting or excessive licking, self-mutilation, walking backwards, vocalisation), or aggression should be recorded.

26.   Signs of toxicity should be recorded, including the day of onset, time of day, degree, and duration.

27.   Animals should be weighed at the time of dosing at least weekly throughout the study, on or near the day of delivery, and on PND 21 (weaning). For gavage studies dams should be weighed at least twice weekly. Doses should be adjusted at the time of each body weight determination, as appropriate. Food consumption should be measured weekly at a minimum during gestation and lactation. Water consumption should be measured at least weekly if exposure is via the water supply.

Observations on offspring

28.   All offspring should be carefully observed at least daily for signs of toxicity and for morbidity and mortality.

29.   During the treatment and observation periods, more detailed clinical observations of the offspring should be conducted. The offspring (at least one pup/sex/litter) should be observed by trained technicians who are unaware of the animals' treatment, using standardised procedures to minimise bias and maximise inter-observer reliability. Where possible, it is advisable that the observations are made by the same technician. At a minimum, the endpoints described in paragraphs 24 and 25 should be monitored as appropriate for the developmental stage being observed.

30.   All signs of toxicity in the offspring should be recorded, including the day of onset, time of day, degree, and duration.

Physical and developmental landmarks

31.   Changes in pre-weaning landmarks of development (e.g.pinna unfolding, eye opening, incisor eruption) are highly correlated with body weight (30)(31). Body weight may be the best indicator of physical development. Measurement of developmental landmarks is, therefore, recommended only when there is prior evidence that these endpoints will provide additional information. Timing for the assessment of these parameters is indicated in Table 1. Depending on the anticipated effects, and the results of the initial measurements, it may be advisable to add additional time points or to perform the measurements in other developmental stages.

32.   It is advisable to use post-coital age instead of postnatal age when assessing physical development (33). If pups are tested on the day of weaning, it is recommended that this testing be carried out prior to actual weaning to avoid a confounding effect by the stress associated with weaning. In addition, any post-weaning testing of pups should not occur during the two days after weaning.

Table 1

Timing of the assessment of physical and developmental landmarks, and functional/behavioural endpoints  (2)

Age Periods


Pre-weaning (3)

Adolescence (3)

Young adults (3)

Physical and developmental landmarks

Body weight and Clinical Observations

weekly (4)

at least every two weeks

at least every two weeks

Brain weight

PND 22 (5)

at termination


PND 22 (5)

at termination

Sexual maturation

as appropriate

Other developmental landmarks (6)

as appropriate

Functional/behavioural endpoints

Behavioural ontogeny

At least two measures



Motor activity (including habituation)

1–3 times (7)


Motor and sensory function



Learning and memory



33.   Live pups should be counted and sexed e.g. by visual inspection or measurement of anogenital distance (34)(35), and each pup within a litter should be weighed individually at birth or soon thereafter, at least weekly throughout lactation, and at least once every two weeks thereafter. When sexual maturation is evaluated, the age and body weight of the animal when vaginal patency (36) or preputial separation (37) occurs should be determined for at least one male and one female per litter.

Behavioural ontogeny

34.   Ontogeny of selected behaviours should be measured in at least one pup/sex/litter during the appropriate age period, with the same pups being used on all test days for all behaviours assessed. The measurement days should be spaced evenly over that period to define either the normal or treatment-related change in ontogeny of that behaviour (38). The following are some examples of behaviours for which their ontogeny could be assessed: righting reflex, negative geotaxis and motor activity (38)(39)(40).

Motor activity

35.   Motor activity should be monitored (41)(42)(43)(44)(45) during the pre-weaning and adult age periods. For testing at the time of weaning, see paragraph 32. The test session should be long enough to demonstrate intra-session habituation for non-treated controls. Use of motor activity to assess behavioural ontogeny is strongly recommended. If used as a test of behavioural ontogeny, then testing should utilise the same animals for all pre-weaning test sessions. Testing should be frequent enough to assess the ontogeny of intra-session habituation (44). This may require three or more time periods prior to, and including the day of weaning (e.g. PND 13, 17, 21). Testing of the same animals, or littermates, should also occur at an adult age close to study termination (e.g. PND 60-70). Testing on additional days may be done as necessary. Motor activity should be monitored by an automated activity recording apparatus which should be capable of detecting both increases and decreases in activity, (i.e. baseline activity as measured by the device should not be so low as to preclude detection of decreases, nor so high as to preclude detection of increases in activity). Each device should be tested by standard procedures to ensure, to the extent possible, reliability of operation across devices and across days. To the extent possible, treatment groups should be balanced across devices. Each animal should be tested individually. Treatment groups should be counter-balanced across test times to avoid confounding by circadian rhythms of activity. Efforts should be made to ensure that variations in the test conditions are minimal and are not systematically related to treatment. Among the variables that can affect many measures of behaviour, including motor activity, are sound level, size and shape of the test cage, temperature, relative humidity, light conditions, odours, use of home cage or novel test cage and environmental distractions.

Motor and sensory function

36.   Motor and sensory function should be examined in detail at least once for the adolescent period and once during the young adult period (e.g. PND 60-70). For testing at the time of weaning, see paragraph 32. Sufficient testing should be conducted to ensure an adequate quantitative sampling of sensory modalities (e.g. somato-sensory, vestibular) and motor functions (e.g. strength, coordination). A few examples of tests for motor and sensory function are extensor thrust response (46), righting reflex (47)(48), auditory startle habituation (40)(49)(50)(51)(52)(53)(54), and evoked potentials (55).

Learning and memory tests

37.   A test of associative learning and memory should be conducted post-weaning (e.g. 25 ± 2 days) and for young adults (PND 60 and older). For testing at the time of weaning, see paragraph 32. The same or separate test(s) may be used at these two stages of development. Some flexibility is allowed in the choice of test(s) for learning and memory in weanling and adult rats. However, the test(s) should be designed so as to fulfil two criteria. First, learning should be assessed either as a change across several repeated learning trials or sessions, or, in tests involving a single trial, with reference to a condition that controls for non-associative effects of the training experience. Second, the test(s) should include some measure of memory (short-term or long-term) in addition to original learning (acquisition), but this measure of memory cannot be reported in the absence of a measure of acquisition obtained from the same test. If the test(s) of learning and memory reveal(s) an effect of the test chemical, additional tests to rule out alternative interpretations based on alterations in sensory, motivational, and/or motor capacities may be considered. In addition to the above two criteria, it is recommended that the test of learning and memory be chosen on the basis of its demonstrated sensitivity to the class of chemical under investigation, if such information is available in the literature. In the absence of such information, examples of tests that could be made to meet the above criteria include: passive avoidance (43)(56)(57), delayed-matching-to-position for the adult rat (58) and for the infant rat (59), olfactory conditioning (43)(60), Morris water maze (61)(62)(63), Biel or Cincinnati maze (64)(65), radial arm maze (66), T-maze (43), and acquisition and retention of schedule-controlled behaviour (26)(67)(68). Additional tests are described in the literature for weanling (26)(27) and adult rats (19)(20).

Post-mortem examination

38.   Maternal animals can be euthanised after weaning of the offspring.

39.   Neuropathological evaluation of the offspring will be conducted using tissues from animals humanely killed at PND 22 or at an earlier time point between PND 11 and PND 22, as well as at study termination. For offspring killed through PND 22, brain tissues should be evaluated; for animals killed at termination, both central nervous system (CNS) tissues and peripheral nervous system (PNS) tissues should be evaluated. Animals killed on PND 22 or earlier may be fixed either by immersion or perfusion. Animals killed at study termination should be fixed by perfusion. All aspects of the preparation of tissue samples, from the perfusion of animals, through the dissection of tissue samples, tissue processing, and staining of slides should employ a counterbalanced design such that each batch contains representative samples from each dose group. Additional guidance on neuropathology can be found in OECD Guidance Document No 20(9), see also (103).

Processing of tissue samples

40.   All gross abnormalities apparent at the time of necropsy should be noted. Tissue samples taken should represent all major regions of the nervous system. The tissue samples should be retained in an appropriate fixative and processed according to standardised published histological protocols (69)(70)(71)(103). Paraffin embedding is acceptable for tissues of the CNS and PNS, but the use of osmium in post-fixation, together with epoxy embedding, may be appropriate when a higher degree of resolution is required (e.g. for peripheral nerves when a peripheral neuropathy is suspected and/or for morphometric analysis of peripheral nerves). Brain tissue collected for morphometric analysis should be embedded in appropriate media at all dose levels at the same time in order to avoid shrinkage artefacts that may be associated with prolonged storage in fixative (6).

Neuropathological examination

41.   The purposes of the qualitative examination are:


to identify regions within the nervous system exhibiting evidence of neuropathological alterations;


to identify types of neuropathological alterations resulting from exposure to the test chemical; and


to determine the range of severity of the neuropathological alterations.

Representative histological sections from the tissue samples should be examined microscopically by an appropriately trained pathologist for evidence of neuropathological alterations. All neuropathologic alterations should be assigned a subjective grade indicating severity. A hematoxylin and eosin stain may be sufficient for evaluating brain sections from animals humanely killed at PND 22, or earlier. However, a myelin stain (e.g. luxol fast blue/cresyl violet) and a silver stain (e.g. Bielschowsky's or Bodians stains) are recommended for sections of CNS and PNS tissues from animals killed at study termination. Subject to the professional judgement of the pathologist and the kind of alterations observed, other stains may be considered appropriate to identify and characterise particular types of alterations (e.g. glial fibrillary acidic protein (GFAP) or lectin histochemistry to assess glial and microglial alterations (72), fluoro-jade to detect necrosis (73)(74), or silver stains specific for neural degeneration (75)).

42.   Morphometric (quantitative) evaluation should be performed as these data may assist in the detection of a treatment-related effect and are valuable in the interpretation of treatment-related differences in brain weight or morphology (76)(77). Nervous tissue should be sampled and prepared to enable morphometric evaluation. Morphometric evaluations may include e.g. linear or areal measurements of specific brain regions (78). Linear or areal measurements require the use of homologous sections carefully selected based on reliable microscopic landmarks (6). Stereology may be used to identify treatment-related effects on parameters such as volume or cell number for specific neuroanatomic regions (79)(80)(81)(82)(83)(84).

43.   The brains should be examined for any evidence of treatment-related neuropathological alterations and adequate samples should be taken from all major brain regions (e.g. olfactory bulbs, cerebral cortex, hippocampus, basal ganglia, thalamus, hypothalamus, midbrain (tectum, tegmentum, and cerebral peduncles), pons, medulla oblongata, cerebellum) to ensure a thorough examination. It is important that sections for all animals are taken in the same plane. In adults humanely killed at study termination, representative sections of the spinal cord and the PNS should be sampled. The areas examined should include the eye with optic nerve and retina, the spinal cord at the cervical and lumbar swellings, the dorsal and ventral root fibres, the proximal sciatic nerve, the proximal tibial nerve (at the knee), and the tibial nerve calf muscle branches. The spinal cord and peripheral nerve sections should include both cross or transverse and longitudinal sections.

44.   Neuropathological evaluation should include an examination for indications of developmental damage to the nervous system (6)(85)(86)(87)(88)(89), in addition to the cellular alterations (e.g. neuronal vacuolation, degeneration, necrosis) and tissue changes (e.g. gliosis, leukocytic infiltration, cystic formation). In this regard, it is important that treatment-related effects be distinguished from normal developmental events known to occur at a developmental stage corresponding to the time of sacrifice (90). Examples of significant alterations indicative of developmental insult include, but are not restricted to:

alterations in the gross size or shape of the olfactory bulbs, cerebrum or cerebellum;

alterations in the relative size of various brain regions, including decreases or increases in the size of regions resulting from the loss or persistence of normally transient populations of cells or axonal projections (e.g. external germinal layer of cerebellum, corpus callosum);

alterations in proliferation, migration, and differentiation, as indicated by areas of excessive apoptosis or necrosis, clusters or dispersed populations of ectopic, disoriented or malformed neurons or alterations in the relative size of various layers of cortical structures;

alterations in patterns of myelination, including an overall size reduction or altered staining of myelinated structures;

evidence of hydrocephalus, in particular enlargement of the ventricles, stenosis of the cerebral aqueduct and thinning of the cerebral hemispheres.

Analysis of the dose-response relationship of neuropathological alterations

45.   The following stepwise procedure is recommended for the qualitative and quantitative neuropathological analyses. First, sections from the high dose group are compared with those of the control group. If no evidence of neuropathological alterations is found in animals of the high dose group, no further analysis is required. If evidence of neuropathological alterations is found in the high dose group, then animals from the intermediate and low dose groups are examined. If the high dose group is terminated due to death or other confounding toxicity, the high and intermediate dose groups should be analysed for neuropathological alterations. If there is any indication of neurotoxicity in lower dose groups, neuropathological analysis should be performed in those groups. If any treatment-related neuropathological alterations are found in the qualitative or quantitative examination, the dose-dependence of the incidence, frequency and severity grade of the lesions or of the morphometric alterations should be determined, based on an evaluation of all animals from all dose groups. All regions of the brain that exhibit any evidence of neuropathologic alteration should be included in this evaluation. For each type of lesion, the characteristics used to define each severity grade should be described, indicating the features used to differentiate each grade. The frequency of each type of lesion and its severity grade should be recorded and a statistical analysis should be performed to evaluate the nature of a dose-response relationships. The use of coded slides is recommended (91).



46.   Data should be reported individually and summarised in tabular form, showing for each test group the types of change and the number of dams, offspring by sex, and litters displaying each type of change. If direct postnatal exposure of the offspring has been performed, the route, duration and period of exposure should be reported.

Evaluation and interpretation of results

47.   A developmental neurotoxicity study will provide information on the effects of repeated exposure to a chemical during in utero and early postnatal development. Since emphasis is placed on both general toxicity and developmental neurotoxicity endpoints, the results of the study will allow for the discrimination between neurodevelopmental effects occurring in the absence of general maternal toxicity, and those which are only expressed at levels that are also toxic to the maternal animal. Due to the complex interrelationships among study design, statistical analysis, and biological significance of the data, adequate interpretation of developmental neurotoxicity data will involve expert judgment (107)(109). The interpretation of test results should use a weight-of-evidence-approach (20)(92)(93)(94). Patterns of behavioural or morphological findings, if present, as well as evidence of dose-response should be discussed. Data from all studies relevant to the evaluation of developmental neurotoxicity, including human epidemiological studies or case reports, and experimental animal studies (e.g. toxicokinetic data, structure-activity information, data from other toxicity studies) should be included in this characterisation. This includes the relationship between the doses of the test chemical and the presence or absence, incidence, and extent of any neurotoxic effect for each sex (20)(95).

48.   Evaluation of data should include a discussion of both the biological and statistical significance. Statistical analysis should be viewed as a tool that guides rather than determines the interpretation of data. Lack of statistical significance should not be the sole rationale for concluding a lack of treatment related effect, just as statistical significance should not be the sole justification for concluding a treatment-related effect. To guard against possible false-negative findings and the inherent difficulties in “proving a negative,” available positive and historical control data should be discussed, especially when there are no treatment-related effects (102)(106). The probability of false positives should be discussed in light of the total statistical evaluation of the data (96). The evaluation should include the relationship, if any, between observed neuropathological and behavioural alterations.

49.   All results should be analysed using statistical models appropriate to the experimental design (108). The choice of a parametric or a nonparametric analysis should be justified by considering factors such as the nature of the data (transformed or not) and their distribution, as well as the relative robustness of the statistical analysis selected. The purpose and design of the study should guide the choice of statistical analyses to minimise Type I (false positive) and Type II (false negative) errors (96)(97)(104)(105). Developmental studies using multiparous species where multiple pups per litter are tested should include the litter in the statistical model to guard against an inflated Type I error rates (98)(99)(100)(101). The statistical unit of measure should be the litter and not the pup. Experiments should be designed such that littermates are not treated as independent observations. Any endpoint repeatedly measured in the same subject should be analysed using statistical models that account for the non-independence of those measures.

Test report

50.   The test report should include the following information:

Test chemical:

physical nature and, where relevant, physiochemical properties;

identification data, including source;

purity of the preparation, and known and/or anticipated impurities.

Vehicle (if appropriate):

justification for choice of vehicle, if other than water or physiological saline solution.

Test animals:

species and strain used, and a justification if other than the rat;

supplier of test animals;

number, age at start, and sex of animals;

source, housing conditions, diet, water, etc.;

individual weights of animals at the start of the test.

Test conditions:

rationale for dose level selection;

rationale for dosing route and time period;

specifications of the doses administered, including details of the vehicle, volume and physical form of the material administered;

details of test chemical formulation/diet preparation, achieved concentration, stability and homogeneity of the preparation;

method used for unique identification of dams and offspring;

a detailed description of the randomisation procedure(s) used to assign dams to treatment groups, to select pups for culling, and to assign pups to test groups;

details of the administration of the test chemical;

conversion from diet/drinking water or inhalation test chemical concentration (ppm) to the actual dose (mg/kg body weight/day), if applicable;

environmental conditions;

details of food and water (e.g. tap, distilled) quality;

dates of study start and end.

Observations and test procedures:

a detailed description of the procedures used to standardise observations and procedures as well as operational definitions for scoring observations;

a list of all test procedures used, and justification for their use;

details of the behavioural/functional, pathological, neurochemical or electrophysiological procedures used, including information and details on automated devices;

procedures for calibrating and ensuring the equivalence of devices and the balancing of treatment groups in testing procedures;

a short justification explaining any decisions involving professional judgement.

Results (individual and summary, including mean and variance when appropriate):

the number of animals at the start of the study and the number at the end of the study;

the number of animals and litters used for each test method;

identification number of each animal and the litter from which it came;

litter size and mean weight at birth by sex;

body weight and body weight change data, including terminal body weight for dams and offspring;

food consumption data, and water consumption data if appropriate (e.g. if test chemical is administered via water);

toxic response data by sex and dose level, including signs of toxicity or mortality, including time and cause of death, if appropriate;

nature, severity, duration, day of onset, time of day, and subsequent course of the detailed clinical observations;

score on each developmental landmark (weight, sexual maturation and behavioural ontogeny) at each observation time;

a detailed description of all behavioural, functional, neuropathological, neurochemical, electrophysiological findings by sex, including both increases and decreases from controls;

necropsy findings;

brain weights;

any diagnoses derived from neurological signs and lesions, including naturally-occurring diseases or conditions;

images of exemplar findings;

low-power images to assess homology of sections used for morphometry;

absorption and metabolism data, including complementary data from a separate toxicokinetic study, if available;

statistical treatment of results, including statistical models used to analyse the data, and the results, regardless of whether they were significant or not;

list of study personnel, including professional training.

Discussion of results:

dose response information, by sex and group;

relationship of any other toxic effects to a conclusion about the neurotoxic potential of the test chemical, by sex and group;

impact of any toxicokinetic information on the conclusions;

similarities of effects to any known neurotoxicants;

data supporting the reliability and sensitivity of the test method (i.e. positive and historical control data);

relationships, if any, between neuropathological and functional effects;

NOAEL or benchmark dose for dams and offspring, by sex and group.


a discussion of the overall interpretation of the data based on the results, including a conclusion of whether or not the test chemical caused developmental neurotoxicity and the NOAEL.



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Chapter B.34 of this Annex, One-generation reproduction toxicity study.


Chapter B.35 of this Annex, Two-generation reproduction toxicity study.


Chapter B.43 of this Annex, Neurotoxicity Study in Rodents.


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Tamura, R.N., Buelke-Sam, J. (1992) The use of repeated measures analysis in developmental toxicology studies. Neurotoxicol. Teratol., 14(3):205-210.


Tukey, J.W., Ciminera, J.L., Heyse, J.F. (1985) Testing the statistical certainty of a response to increasing doses of a drug. Biometrics, 41:295-301.


Crofton, K.M., Foss, J.A., Haas, U., Jensen, K., Levin, E.D., and Parker, S.P. (2008) Undertaking positive control studies as part of developmental neurotoxicity testing: report from the ILSI Research Foundation/Risk Science Institute expert working group on neurodevelopmental endpoints. Neurotoxicology and Teratology, 30(4):266-287.


Raffaele, K.C., Fisher, E., Hancock, S., Hazelden, K., and Sobrian, S.K. (2008) Determining normal variability in a developmental neurotoxicity test: report from the ILSI Research Foundation/Risk Science Institute expert working group on neurodevelopmental endpoints. Neurotoxicology and Teratology, 30(4):288-325.


Holson, R.R., Freshwater, L., Maurissen, J.P.J., Moser, V.C., and Phang, W. (2008) Statistical issues and techniques appropriate for developmental neurotoxicity testing: a report from the ILSI Research Foundation/Risk Science Institute expert working group on neurodevelopmental endpoints. Neurotoxicology and Teratology, 30(4):326-348.


Tyl, R.W., Crofton, K.M., Moretto, A., Moser, V.C., Sheets, L.P., and Sobotka, T.J. (2008) Identification and interpretation of developmental neurotoxicity effects: a report from the ILSI Research Foundation/Risk Science Institute expert working group on neurodevelopmental endpoints Neurotoxicology and Teratology, 30(4):349-381.

Figure 1

General testing scheme for functional/behavioural tests, neuropathology evaluation, and brain weights. This diagram is based on the description in paragraphs 13-15 (PND=postnatal day). Examples of animal assignment are given in Appendix 1.

Image 1

Approximately 20 litters/group

Offspring: Approximately 80/sex/group:

Selected on or before PND 4 for pre- and post-weaning investigations:

Clinical observations and body weight (all animals)

Detailed clinical observation (20/sex/group)

Behavioural ontogeny (20/sex/group)

Motor activity (20/sex/group)

Sexual maturation (20/sex/group)

Motor and sensory function (20/sex/group)

Learning and memory (10-20/sex/group)

Neuropathology: PND 11-22


Immersion or perfusion fixation of brains for neuropathology evaluation. Brain weight (fixed).

Option: Additional testing


Brain weight (unfixed).

Neuropathology: PND 70 (study termination)


Perfusion fixation of brains for neuropathology evaluation.


Brain weight (unfixed).

Neuropathology not required.


Option: Additional testing

Appendix 1


Examples of possible assignments are described and tabulated below. These examples are provided to illustrate that assignment of study animals to various testing paradigms can be accomplished in a number of different ways.

Example 1


One set of 20 pups/sex/dose level (i.e. 1 male and 1 female per litter) is used for pre-weaning testing of behavioural ontogeny. Out of these animals, 10 pups/sex/dose level (i.e. 1 male or 1 female per litter) are humanely killed at PND 22. The brains are removed, weighed and processed for histopathologic evaluation. In addition, brain weight data are collected using unfixed brains from the remaining 10 males and 10 females per dose level.


Another set of 20 animals/sex/dose level (i.e. 1 male and 1 female per litter) is used for post-weaning functional/behavioral tests (detailed clinical observations, motor activity, auditory startle and cognitive function testing in adolescents) and assessing age of sexual maturation. Of these animals, 10 animals/sex/dose level (i.e. 1 male or 1 female per litter), are anesthetised and fixed via perfusion at study termination (approximately PND 70). After additional fixation in situ, the brain is removed and processed for neuropathological evaluation.


For cognitive function testing in young adults (e.g. PND 60-70), a third set of 20 pups/sex/dose level is used (i.e. 1 male and 1 female per litter). Of these animals, 10 animals/sex/group (1 male or 1 female per litter) are killed at study termination and the brain is removed and weighed.


The remaining 20 animals/sex/group are reserved for possible additional tests.

Table 1

Pup No (8)

No of pups assigned to test








20 m + 20 f

Behavioural ontogeny



10 m + 10 f

PND 22 brain weight/neuropathology/morphometry



10 m + 10 f

PND 22 brain weight







20 m + 20 f

Detailed clinical observations



20 m + 20 f

Motor activity



20 m + 20 f

Sexual maturation



20 m + 20 f

Motor and sensory function



20 m + 20 f

Learning and memory (PND 25)



10 m + 10 f

Young adult brain weight/neuropathology/morphometry ~ PND 70







20 m + 20 f

Learning and memory (young adults)



10 m + 10 f

Young adult brain weight ~ PND 70



Reserve animals for replacements or additional tests

Example 2


One set of 20 pups/sex/dose level (i.e. 1 male and 1 female per litter) is used for pre-weaning testing of behavioural ontogeny. Out of these animals, 10 pups/sex/dose level (1 male or 1 female per litter), are humanely killed at PND 11. The brains are removed, weighed and processed for histopathologic evaluation.


Another set of 20 animals/sex/dose level (1 male and1 female per litter) is used for post-weaning examinations (detailed clinical observations, motor activity, assessing age of sexual maturation and motor and sensory function). Of these animals, 10 animals/sex/dose level (i.e.1 male or 1 female per litter) are anesthetised and fixed via perfusion at study termination (approximately PND 70). After additional fixation in situ, the brain is removed, weighed and processed for neuropathological evaluation.


For cognitive function testing in adolescents and young adults, 10 pups/sex/dose level are used(i.e. 1 male or 1 female per litter). Different animals are used for testing for cognitive function tests at PND 23 and young adults. At termination, the 10 animals/sex/group tested as adults are killed, the brain is removed and weighed.


The remaining 20 animals/sex/group not selected for testing are killed and discarded at weaning.

Table 2

Pup No (9)

No of pups assigned to test








20 m + 20 f

Behavioural ontogeny



10 m + 10 f

PND 11 brain weight/neuropathology/morphometry



20 m + 20 f

Detailed clinical observations



20 m + 20 f

Motor activity



20 m + 20 f

Sexual maturation



20 m + 20 f

Motor and sensory function



10 m + 10 f

Young adult brain weight/neuropathology/morphometry ~ PND 70







10 m + 10 f (10)

Learning and memory (PND 23)



10 m + 10 f (10)

Learning and memory (young adults)




Young adult brain weight



Animals killed and discarded PND 21.

Example 3


One set 20 pups/sex/dose level (i.e. 1 male and 1 female per litter) is used for brain weight and neuropathology assessment at PND 11. Out of these animals, 10 pups/sex/dose level (i.e. 1 male or 1 female per litter) are humanely killed at PND 11 and brains are removed, weighed and processed for histopathologic evaluation. In addition, brain weight data are collected using unfixed brains from the remaining 10 males and 10 females per dose level.


Another set of of 20 animals/sex/dose level (i.e. 1 male and 1 female per litter) are used for behavioural ontogeny (motor activity), post-weaning examinations (motor activity and assessing age of sexual maturation), and cognitive function testing in adolescents.


Another set of 20 animals/sex/dose level (i.e. 1 male and 1 female per litter) is used for motor and sensory function tests (auditory startle) and detailed clinical observations. Of these animals, 10 animals/sex/dose level (i.e. 1 male or 1 female per litter) are anesthetised and fixed via perfusion at study termination (approximately PND 70). After additional fixation in situ, the brain is removed, weighed and processed for neuropathological evaluation.


Another set of 20 pups/sex/dose level are usedfor cognitive function testing in young adults (i.e. 1 male and 1 female per litter). Of these, 10 animals/sex/group (i.e. 1 male or 1 female per litter) are killed at termination, the brain removed and weighed.

Table 3

Pup No (11)

No of pups assigned to test








10 m + 10 f

PND 11 brain weight/neuropathology/morphometry



10 m + 10 f

PND 11 brain weight



20 m + 20 f

Behavioural ontogeny (motor activity)



20 m + 20 f

Motor activity



20 m + 20 f

Sexual maturation



20 m + 20 f

Learning and memory (PND 27)







20 m + 20 f

Auditory startle (adolescents and young adults)



20 m + 20 f

Detailed clinical observations



10 m + 10 f

Young adult brain weight/neuropathology/morphometry ~ PND 70



20 m + 20 f

Learning and memory (young adults)



10 m + 10 f

Young adult brain weight





Appendix 2


Chemical: A substance or a mixture

Test chemical: Any substance or mixture tested using this test method



1.   This test method is equivalent to OECD Test Guideline (TG) 440 (2007). The OECD initiated a high-priority activity in 1998 to revise existing guidelines and to develop new guidelines for the screening and testing of potential endocrine disrupters (1). One element of the activity was to develop a test guideline for the rodent Uterotrophic Bioassay. The rodent Uterotrophic Bioassay then underwent an extensive validation programme including the compilation of a detailed background document (2)(3) and the conduct of extensive intra- and interlaboratory studies to show the relevance and reproducibility of the bioassay with a potent reference oestrogen, weak oestrogen receptor agonists, a strong oestrogen receptor antagonist, and a negative reference chemical (4)(5)(6)(7)(8)(9). This test method B.54 is the outcome of the experience gained during the validation test programme and the results obtained thereby with oestrogenic agonists.

2.   The Uterotrophic Bioassay is a short-term screening test that originated in the 1930s (27)(28) and was first standardised for screening by an expert committee in 1962 (32)(35). It is based on the increase in uterine weight or uterotrophic response (for review, see 29). It evaluates the ability of a chemical to elicit biological activities consistent with agonists or antagonists of natural oestrogens (e.g. 17ß-estradiol), however, its use for antagonist detection is much less common than for agonists. The uterus responds to oestrogens in two ways. An initial response is an increase in weight due to water imbibition. This response is followed by a weight gain due to tissue growth (30). The uterus responses in rats and mice qualitatively are comparable.

3.   This bioassay serves as an in vivo screening assay and its application should be seen in the context of the “OECD Conceptual Framework for the Testing and Assessment of Endocrine Disrupting Chemicals” (Appendix 2). In this Conceptual Framework the Uterotrophic Bioassay is contained in Level 3 as an in vivo assay providing data about a single endocrine mechanism, i.e. oestrogenicity.

4.   The Uterotrophic Bioassay is intended to be included in a battery of in vitro and in vivo tests to identify chemicals with potential to interact with the endocrine system, ultimately leading to risk assessments for human health or the environment. The OECD validation programme used both strong and weak oestrogen agonists to evaluate the performance of the assay to identify oestrogenic chemicals (4)(5)(6)(7)(8). Thereby the sensitivity of the test procedure for oestrogen agonists was well demonstrated besides a good intra- and interlaboratory reproducibility.

5.   With regard to negative chemicals, only one “negative” reference chemical already reported negative by uterotrophic assay as well as in vitro receptor binding and receptor assays was included in the validation programme, but additional test data, not related to the OECD validation programme, have been evaluated, giving further support to the specificity of the Uterotrophic Bioassay for the screening of oestrogen agonists (16).


6.   Oestrogen agonists and antagonists act as ligands for oestrogen receptors a and b and may activate or inhibit, respectively, the transcriptional action of the receptors. This may have the potential to lead to adverse health hazards, including reproductive and developmental effects. Therefore, the need exists to rapidly assess and evaluate a chemical as a possible oestrogen agonist or antagonist. While informative, the affinity of a ligand for an oestrogen receptor or transcriptional activation of reporter genes in vitro is only one of several determinants of possible hazard. Other determinants can include metabolic activation and deactivation upon entering the body, distribution to target tissues, and clearance from the body, depending at least in part on the route of administration and the chemical being tested. This leads to the need to screen the possible activity of a chemical in vivo under relevant conditions, unless the chemical's characteristics regarding Absorption — Distribution — Metabolism — Elimination (ADME) already provide appropriate information. Uterine tissues respond with rapid and vigorous growth to stimulation by oestrogens, particularly in laboratory rodents, where the oestrous cycle lasts approximately 4 days. Rodent species, particularly the rat, are also widely used in toxicity studies for hazard characterisation. Therefore, the rodent uterus is an appropriate target organ for the in vivo screening of oestrogen agonists and antagonists.

7.   This test method is based on those protocols employed in the OECD validation study which have been shown to be reliable and repeatable in intra- and interlaboratory studies (5)(7). Currently two methods, namely the ovariectomised adult female method (ovx-adult method) and the immature non-ovariectomised method (immature method) are available. It was shown in the OECD validation test programme that both methods have comparable sensitivity and reproducibility. However, the immature, as it has an intact hypothalamic-pituitary-gonadal (HPG) axis, is somewhat less specific but covers a larger scope of investigation than the ovariectomised animal because it can respond to chemicals that interact with the HPG axis rather than just the oestrogen receptor. The HGP axis of the rat is functional at about 15 days of age. Prior to that, puberty cannot be accelerated with treatments like GnRH. As the females begin to reach puberty, prior to vaginal opening, the female will have several silent cycles that do not result in vaginal opening or ovulation, but there are some hormonal fluctuations. If a chemical stimulates the HPG axis directly or indirectly, precocious puberty, early ovulation and accelerated vaginal opening result. Not only chemicals that act on the HPG axis do this but some diets with higher metabolisable energy levels than others will stimulate growth and accelerate vaginal opening without being oestrogenic. Such chemicals would not induce an uterotrophic response in OVX adult animals as their HPG axis does not work.

8.   For animal welfare reasons preference should be given to the method using immature rats, avoiding surgical pre-treatment of the animals and avoiding also a possible non-use of those animals which indicate any evidence entering oestrous (see paragraph 30).

9.   The uterotrophic response is not entirely of oestrogenic origin, i.e. chemicals other than agonists or antagonists of oestrogens may also provide a response. For example, relatively high doses of progesterone, testosterone, or various synthetic progestins may all lead to a stimulative response (30). Any response may be analysed histologically for keratinisation and cornification of the vagina (30). Irrespective of the possible origin of the response, a positive outcome of an Uterotrophic Bioassay should normally initiate actions for further clarification. Additional evidence of oestrogenicity could come from in vitro assays, such as the ER binding assays and transcriptional activation assays, or from other in vivo assays such as the female pubertal assay.

10.   Taking into account that the Uterotrophic Bioassay serves as an in vivo screening assay, the validation approach taken served both animal welfare considerations and a tiered testing strategy. To this end, effort was directed at rigorously validating reproducibility and sensitivity for oestrogenicity — the main concern for many chemicals-, while little effort was directed at the antioestrogenicity component of the assay. Only one antioestrogen with strong activity was tested since the number of chemicals with a clear antioestrogenic profile (not obscured by some oestrogenic activity) is very limited. Thus this test method is dedicated to the oestrogenic protocol, while the protocol describing the antagonist mode of the assay is included in a Guidance Document (37). The reproducibility and sensitivity of the assay for chemicals with purely anti-oestrogenic activity will be more clearly defined later on, after the test procedure has been in routine use for some time and more chemicals with this modality of action are identified.

11.   It is acknowledged that all animal based procedures will conform to local standards of animal care; the descriptions of care and treatment set forth below are minimal performance standards, and will be superseded by local regulations such as Directive 2010/63/EU of the European Parliament and of the Council of 22 September 2010 on the protection of animals used for scientific purposes (38). Further guidance of the humane treatment of animals is given by the OECD (25).

12.   As with all assays using live animals, it is essential to ensure that the data are truly necessary prior to the start of the assay. For example, two conditions where the data may be required are:

high exposure potential (Level 1 of the Conceptual Framework, Appendix 2) or indications for oestrogenicity (Level 2) to investigate whether such effects may occur in vivo;

effects indicating oestrogenicity in Level 4 or 5 in vivo tests to substantiate that the effects were related to an oestrogenic mechanism that cannot be elucidated using an in vitro test.

13.   Definitions used in this test method are given in Appendix 1.


14.   The Uterotrophic Bioassay relies for its sensitivity on an animal test system in which the hypothalamic-pituitary-ovarian axis is not functional, leading to low endogenous levels of circulating oestrogen. This will ensure a low baseline uterine weight and a maximum range of response to administered oestrogens. Two oestrogen sensitive states in the female rodent meet this requirement:


immature females after weaning and prior to puberty; and


young adult females after ovariectomy with adequate time for uterine tissues to regress.

15.   The test chemical is administered daily by oral gavage or subcutaneous injection. Graduated test chemical doses are administered to a minimum of two treatment groups (see paragraph 33 for guidance) of experimental animals using one dose level per group and an administration period of three consecutive days for immature method and a minimum administration period of three consecutive days for ovx-adult method. The animals are necropsied approximately 24 hours after the last dose. For oestrogen agonists, the mean uterine weight of the treated animal groups relative to the vehicle group is assessed for a statistically significant increase. A statistically significant increase in the mean uterine weight of a test group indicates a positive response in this bioassay.


Selection of animal species

16.   Commonly used laboratory rodent strains may be used. As an example, Sprague-Dawley and Wistar strains of rats were used during the validation. Strains with uteri known or suspected to be less responsive should not be used. The laboratory should demonstrate the sensitivity of the strain used as described in paragraphs 26 and 27.

17.   The rat and mouse have been routinely used in the Uterotrophic Bioassay since the 1930s. The OECD validation studies were only performed with rats based on an understanding that both species are expected to be equivalent and therefore one species should be enough for the world-wide validation in order to save resources and animals. The rat is the species of choice in most reproductive and developmental toxicity studies. Taking into consideration that a vast historical database exists for mice and thus to broaden the scope of the Uterotrophic Bioassay test method in rodents to the use of mice as test species, a limited follow-up validation study was carried out in mice (16). A bridging approach with a limited number of test chemicals, participating laboratories and without coded sample testing has been selected in keeping with the original intent to save resources and animals. This bridging validation study shows for the Uterotrophic Bioassay in young adult ovariectomised mice that, qualitatively and quantitatively, the data obtained in rats and mice correspond well with each other. Where the Uterotrophic Bioassay result may be preliminary to a long-term study, this allows animals from the same strain and source to be used in both studies. The bridging approach was limited to the OVX mice and the report does not provide a robust data set to validate the immature model, thus the immature model for mice is not considered under the scope of the current test method.

18.   Thus, in some cases mice may be used instead of rats. A rationale should be given for this species, based on toxicological, pharmacokinetic, and/or other criteria. Modifications of the protocol may be necessary for mice. For example, the food consumption of mice on a body weight basis is higher than that of rats and therefore the phyto-oestrogen content in food should be lower for mice than for rats (9)(20)(22).

Housing and feeding conditions

19.   All procedures should conform with local standards of laboratory animal care. These descriptions of care and treatment are minimum standards and will be superseded by local regulations such as Directive 2010/63/EU of the European Parliament and of the Council of 22 September 2010 on the protection of animals used for scientific purposes (38). The temperature in the experimental animal room should be 22 °C (with an approximate range ± 3 °C). The relative humidity should be a minimum of 30 % and preferably should not exceed a maximum 70 %, other than during room cleaning. The aim should be relative humidity of 50-60 %. Lighting should be artificial. The daily lighting sequence should be 12 hours light, 12 hours dark.

20.   Laboratory diet and drinking water should be provided ad libitum. Young adult animals may be housed individually or be caged in groups of up to three animals. Due to the young age of the immature animals, social group housing is recommended.

21.   High levels of phyto-oestrogens in laboratory diets have been known to increase uterine weights in rodents to a degree enough as to interfere with the Uterotrophic Bioassay (13)(14)(15). High levels of phyto-oestrogens and of metabolisable energy in laboratory diets may also result in early puberty, if immature animals are used. The presence of phyto-oestrogens results primarily from the inclusion of soy and alfalfa products in the laboratory diets and concentrations of phyto-oestrogens have been shown to vary from batch-to-batch of standard laboratory diets (23). Body weight is an important variable, as the quantity of food consumed is related to body weight. Therefore, the actual phyto-oestrogen dose consumed from the same diet may vary among species and by age (9). For immature female rats, food consumption on a body weight basis may be approximately double that of ovariectomised young adult females. For young adult mice, food consumption on a body weight basis may be approximately quadruple that of ovariectomised young adult female rats.

22.   Uterotrophic Bioassay results (9)(17)(18)(19), however, show that limited quantities of dietary phyto-oestrogens are acceptable and do not reduce the sensitivity of the bioassay. As a guide, dietary levels of phyto-oestrogens should not exceed 350 μg of genistein equivalents/gram of laboratory diet for immature female Sprague Dawley and Wistar rats (6)(9). Such diets should also be appropriate when testing in young adult ovariectomised rats because food consumption on a body weight basis is less in young adult as compared to immature animals. If adult ovariectomised mice or more phyto-oestrogen-sensitive rats are to be used, proportional reduction in dietary phyto-oestrogen levels must be considered (20). In addition, the differences in available metabolic energy from different diets may lead to time shifts for the onset of puberty (21)(22).

23.   Prior to the study, careful selection is required of a diet without an elevated level of phyto-oestrogens (for guidance see (6)(9)) or metabolisable energy, that can confound the results (15)(17)(19)(22)(36). Ensuring the proper performance of the test system used by the laboratory as specified in paragraphs 26 and 27 is an important check on both of these factors. As a safeguard consistent with good laboratory practice (GLP) representative sampling of each batch of diet administered during the study should be conducted for possible analysis of phyto-oestrogen content (e.g. in the case of high uterine control weight relative to historic controls or an inadequate response to the reference oestrogen, 17 alpha ethinyl estradiol). Aliquots should be analysed as part of the study or frozen at – 20 °C or in such a way as to prevent the sample from decomposing prior to analysis.

24.   Some bedding materials may contain naturally occurring oestrogenic or antioestrogenic chemicals (e.g. corn cob is known to affects the cyclicity of rats and appears to be antioestrogenic). The selected bedding material should contain a minimum level of phyto-oestrogens.

Preparation of animals

25.   Experimental animals without evidence of any disease or physical abnormalities are randomly assigned to the control and treatment groups. Cages should be arranged in such a way that possible effects due to cage placement are minimised. The animals should be identified uniquely. Preferably, immature animals should be caged with dams or foster dams until weaning during acclimatisation. The acclimatisation period prior to the start of the study should be about 5 days for young adult animals and for the immature animals delivered with dams or foster dams. If immature animals are obtained as weanlings without dams a shorter duration of the acclimatisation period may become necessary as dosing should start immediately after weaning (see paragraph 29).


Verification of Laboratory Proficiency

26.   Two different options can be used to verify laboratory proficiency:

Periodic verification, relying on an initial baseline positive control study (see paragraph 27). At least every 6 months and each time there is a change that may influence the performance of the assay (e.g. a new formulation of diet, change in personnel performing dissections, change in animal strain or supplier, etc.), the responsiveness of the test system (animal model) should be verified using an appropriate dose (based on the baseline positive control study described in paragraph 27) of a reference oestrogen: 17a-ethinyl estradiol (CAS No 57-63-6) (EE).

Use of concurrent controls, by including a group administered with an appropriate dose of reference oestrogen in each assay.

If the system does not respond as expected, the experimental conditions should be examined and modified accordingly. It is recommended that the dose of reference oestrogen to be used in either approach be approximately the ED70 to 80.

27.    Baseline Positive Control Study — Before a laboratory conducts a study under this test method for the first time, laboratory proficiency should be demonstrated by testing the responsiveness of the animal model, by establishing the dose response of a reference oestrogen: 17a-ethinyl estradiol (CAS No 57-63-6) (EE) with a minimum of four doses. The uterine weight response will be compared to established historical data (see reference (5)). If this baseline positive control study does not yield the anticipated results the experimental conditions should be examined and modified.

Number and condition of animals

28.   Each treated and control group should include at least 6 animals (for both immature and ovx-adult method protocols).

Age of immature animals

29.   For the Uterotrophic Bioassay with immature animals the day of birth must be specified. Dosing should begin early enough to ensure that, at the end of test chemical administration, the physiological rise of endogenous oestrogens associated with puberty has not yet taken place. On the other hand, there is evidence that very young animals may be less sensitive. For defining the optimal age each laboratory should take its own background data on maturation into consideration.

As a general guide, dosing in rats may begin immediately after early weaning on postnatal day 18 (with the day of birth being postnatal day 0). Dosing in rats preferably should be completed on postnatal day 21 but in any case prior to postnatal day 25, because, after this age, the hypothalamic-pituitary-ovarian axis becomes functional and endogenous oestrogen levels may begin to rise with a concomitant increase in baseline uterine weight means and an increase in the group standard deviations (2)(3)(10)(11)(12).

Procedure for ovariectomy

30.   For the ovariectomised female rat and mouse (treatment and control groups), ovariectomy should occur between 6 and 8 weeks of age. For rats, a minimum of 14 days should elapse between ovariectomy and the first day of administration in order to allow the uterus to regress to a minimum, stable baseline. For mice, at least 7 days should elapse between ovariectomy and the first day of administration. As small amounts of ovarian tissue are sufficient to produce significant circulating levels of oestrogens (3), the animals should be tested prior to use by observing epithelial cells swabbed from the vagina on at least five consecutive days (e.g. days 10-14 after ovariectomy for rats). If the animals indicate any evidence entering oestrous, the animals should not be used. Further, at necropsy, the ovarian stubs should be examined for any evidence that ovarian tissue is present. If so, the animal should not be used in the calculations (3).

31.   The ovariectomy procedure begins with the animal in ventral recumbency after the animal has been properly anesthetised. The incision opening the dorso-lateral abdominal wall should be approximately 1 cm lengthways at the mid-point between the costal inferior border and the iliac crest, and a few millimetres lateral to the lateral margin of the lumbar muscle. The ovary should be removed from the abdominal cavity onto an aseptic field. The ovary should be disconnected at the junction of the oviduct and the uterine body. After confirming that no massive bleeding is occurring, the abdominal wall should be closed by a suture and the skin closed by autoclips or appropriate suture. The ligation points are shown schematically in Figure 1. Appropriate post-operative analgesia should be used as recommended by a veterinarian experienced in rodent care.

Body weight

32.   In the ovx-adult method, body weight and uterine weight are not correlated because uterine weight is affected by hormones like oestrogens but not by the growth factors that regulate body size. On the contrary, body weight is related to uterine weight in the immature model, while it is maturing (34). Thus, at the commencement of the study the weight variation of animals used, in the immature model, should be minimal and not exceed ± 20 % of the mean weight. This means that the litter size should be standardised by the breeder, to ensure that offspring of different mother animals will be fed approximately the same. Animals should be assigned to groups (both control and treatment) by randomised weight distribution, so that mean body weight of each group is not statistically different from any other group. Consideration should be given to avoid assignment of littermates to the same treatment group as far as practicable without increasing the number of litters to be used for the investigation.


33.   In order to establish whether a test chemical can have oestrogenic action in vivo, two dose groups and a control are normally sufficient and this design is therefore preferred for animal welfare reasons. If the purpose is either to obtain a dose-response curve or to extrapolate to lower doses, at least 3 dose groups are needed. If information beyond identification of oestrogenic activity (such as an estimate of potency) is required, a different dosing regimen should be considered. Except for treatment with the test chemical, animals in the control group should be handled in an identical manner to the test group subjects. If a vehicle is used in administering the test chemical, the control group should receive the same amount of vehicle used with the treated groups (or highest volume used with the test groups if different among groups).

34.   The objective in the case of the Uterotrophic Bioassay is to select doses that ensure animal survival and that are without significant toxicity or distress to the animals after three consecutive days of chemical administration up to a maximum dose of 1 000 mg/kg/d. All dose levels should be proposed and selected taking into account any existing toxicity and (toxico-) kinetic data available for the test chemical or related materials. The highest dose level should first take into consideration the LD50 and/or acute toxicity information in order to avoid death, severe suffering or distress in the animals (24)(25)(26). The highest dose should represent the maximum tolerated dose (MTD); a study conducted at a dose level that induced a positive uterotrophic response would be accepted too. As a screen, large intervals (e.g. one half log units corresponding to a dose progression of 3,2 or even up to one log units) between dosages are generally acceptable. If there are no suitable data available, a range finding study may be performed to aid the determination of the doses to be used.

35.   Alternatively, if the oestrogenic potency of an agonist can be estimated by in vitro (or in silico) data, these may be taken into consideration for dose selection. For example, the amount of the test chemical that would produce uterotrophic responses equivalent to the reference agonist (ethinyl estradiol) is estimated by its relative in vitro potencies to ethinyl estradiol. The highest test dose would be given by multiplying this equivalent dose by an appropriate factor e.g. 10 or 100.

Considerations for range finding

36.   If necessary, a preliminary range finding study can be carried out with few animals. In this respect, OECD Guidance Document No 19(25) may be used defining clinical signs indicative of toxicity or distress to the animals. If feasible within this range finding study after three days of administration, the uteri may be excised and weighed approximately 24-hours after the last dose. These data could then be used to assist the main study design (select an acceptable maximum and lower doses and recommend the number of dose groups).

Administration of doses

37.   The test chemical is administered by oral gavage or subcutaneous injection. Animal welfare considerations as well as toxicological aspects like the relevance to the human route of exposure to the chemical (e.g. oral gavage to model ingestion, subcutaneous injection to model inhalation or dermal adsorption), the physical/chemical properties of the test material and especially existing toxicological information and data on metabolism and kinetics (e.g. need to avoid first pass metabolism, better efficiency via a particular route) have to be taken into account when choosing the route of administration.

38.   It is recommended that, wherever possible, the use of an aqueous solution/suspension be considered first. But as most oestrogen ligands or their metabolic precursors tend to be hydrophobic, the most common approach is to use a solution/suspension in oil (e.g. corn, peanut, sesame or olive oil). However, these oils have different caloric and fat content, thus the vehicle might affect total metabolisable energy (ME) intake, thereby potentially altering measured endpoints such as the uterine weight especially in the immature method (33). Thus, prior to the study, any vehicle to be used should be tested against controls without vehicles. Test chemicals can be dissolved in a minimal amount of 95 % ethanol or other appropriate solvents and diluted to final working concentrations in the test vehicle. The toxic characteristics of the solvent must be known, and should be tested in a separate solvent-only control group. If the test chemical is considered stable, gentle heating and vigorous mechanical action can be used to assist in dissolving the test chemical. The stability of the test chemical in the vehicle should be determined. If the test chemical is stable for the duration of the study, then one starting aliquot of the test chemical may be prepared, and the specified dosage dilutions prepared daily.

39.   Dosage timing will depend of the model used (refer to paragraph 29 for the immature model and to paragraph 30 for ovx-adult model). Immature female rats are dosed with the test chemical daily for three consecutive days. A three-day treatment is also recommended for ovariectomised female rats but longer exposures are acceptable and may improve the detection of weakly active chemicals. With ovariectomised female mice, an application duration of 3 days should be sufficient without a significant advantage by an extension of up to seven days for strong oestrogen agonists, however, this relation was not demonstrated for weak oestrogens in the validation study (16) thus dosage should be extended up to 7 consecutive days in ovx-adult mice.The dose should be given at similar times each day. They should be adjusted as necessary to maintain a constant dose level in terms of animal body weight (e.g. mg of test chemical per kg of body weight per day). Regarding the test volume, its variability, on a body weight basis, should be minimised by adjusting the concentration of the dosing solution to ensure a constant volume on a body weight basis at all dose levels and for any route of administration.

40.   When the test chemical is administered by gavage, this should be done in a single daily dose to the animals using a stomach tube or a suitable intubation cannula. The maximum volume of liquid that can be administered at one time depends on the size of the test animal. Local animal care guidelines should be followed, but the volume should not exceed 5 ml/kg body weight, except in the case of aqueous solutions where 10 ml/kg body weight may be used.

41.   When the test chemical is administered by subcutaneous injection, this should be done in a single daily dose. Doses should be administered to the dorsoscapular or lumbar regions via sterile needle (e.g. 23- or 25-gauge) and a tuberculin syringe. Shaving the injection site is optional. Any losses, leakage at the injection site or incomplete dosing should be recorded. The total volume injected per rat per day should not exceed 5 ml/kg body weight, divided into 2 injection sites, except in the case of aqueous solutions where 10 ml/kg body weight may be used.


General and clinical observations

42.   General clinical observations should be made at least once a day and more frequently when signs of toxicity are observed. Observations should be carried out preferably at the same time(s) each day and considering the period of anticipated peak effects after dosing. All animals are to be observed for mortality, morbidity and general clinical signs such as changes in behaviour, skin, fur, eyes, mucous membranes, occurrence of secretions and excretions and autonomic activity (e.g. lacrimation, piloerection, pupil size, unusual respiratory pattern).

Body weight and food consumption

43.   All animals should be weighed daily to the nearest 0,1 g, starting just prior to initiation of treatment i.e. when the animals are allocated into groups. As an optional measurement, the amount of food consumed during the treatment period may be measured per cage by weighing the feeders. The food consumption results should be expressed in grams per rat per day.

Dissection and measurement of uterus weight

44.   Twenty-four hours after the last treatment, the rats will be humanely killed. Ideally, the necropsy order will be randomised across groups to avoid progression directly up or down dose groups that could subtly affect the data. The bioassay objective is to measure both the wet and blotted uterus weights. The wet weight includes the uterus and the luminal fluid contents. The blotted weight is measured after the luminal contents of the uterus have been expressed and removed.

45.   Before dissection the vagina will be examined for opening status in immature animals. The dissection procedure begins by opening the abdominal wall starting at the pubic symphysis. Then, uterine horn and ovaries, if present, are detached from the dorsal abdominal wall. The urinary bladder and ureters are removed from the ventral and lateral side of uterus and vagina. Fibrous adhesion between the rectum and the vagina is detached until the junction of vaginal orifice and perineal skin can be identified. The uterus and vagina are detached from the body by incising the vaginal wall just above the junction between perineal skin as shown in Figure 2. The uterus should be detached from the body wall by gently cutting the uterine mesentery at the point of its attachment along the full length of the dorsolateral aspect of each uterine horn. Once removed from the body, uterine handling should be sufficiently rapid to avoid desiccation of the tissues. Loss of weight due to desiccation becomes more important with small tissues such as the uterus (23). If ovaries are present, the ovaries are removed at the oviduct avoiding loss of luminal fluid from the uterine horn. If the animal has been ovariectomised, the stubs should be examined for the presence of any ovarian tissue. Excess fat and connective tissue should be trimmed away. The vagina is removed from the uterus just below the cervix so that the cervix remains with the uterine body as shown in Figure 2.

46.   Each uterus should be transferred to a uniquely marked and weighed container (e.g. a petri-dish or plastic weight boat) with continuing care to avoid desiccation before weighing (e.g. filter paper slightly dampened with saline may be placed in the container). The uterus with luminal fluid will be weighed to the nearest 0,1 mg (wet uterine weight).

47.   Each uterus will then be individually processed to remove the luminal fluid. Both uterine horns will be pierced or cut longitudinally. The uterus will be placed on lightly moistened filter paper (e.g. Whatman No 3) and gently pressed with a second piece of lightly moistened filter paper to completely remove the luminal fluid. The uterus without the luminal contents will be weighed to the nearest 0,1 mg (blotted uterine weight).

48.   The uterus weight at termination can be used to ensure that the appropriate age in the immature intact rat was not exceeded, however, the historical data of the rat strain used by the laboratory are decisive in this respect (see paragraph 56 for interpretation of the results).

Optional investigations

49.   After weighing, the uterus may be fixed in 10 % neutral buffered formalin to be examined histopathologically after Haematoxylin & Eosin (HE)-staining. The vagina may be investigated accordingly (see paragraph 9). In addition, morphometric measurement of endometrial epithelium may be done for quantitative comparison.



50.   Study data should include:

the number of animals at the start of the assay,

the number and identity of animals found dead during the assay or killed for humane reasons and the date and time of any death or humane kill,

the number and identity of animals showing signs of toxicity, and a description of the signs of toxicity observed, including time of onset, duration, and severity of any toxic effects, and

the number and identity of animals showing any lesions and a description of the type of lesions.

51.   Individual animal data should be recorded for the body weights, the wet uterine weight, and the blotted uterine weight. One-tailed statistical analyses for agonists should be used to determine whether the administration of a test chemical resulted in a statistically significant (p < 0,05) increase in the uterine weight. Appropriate statistical analyses should be carried out to test for treatment related changes in blotted and wet uterine weight. For example, the data may be evaluated by an analysis of covariance (ANCOVA) approach with body weight at necropsy as the co-variable. A variance-stabilising logarithmic transformation may be carried out on the uterine data prior to the data analysis. Dunnett and Hsu's test are appropriate for making pair wise comparisons of each dosed group to vehicle controls and to calculate the confidence intervals. Studentised residual plots can be used to detect possible outliers and to assess homogeneity of variances. These procedures were applied in the OECD validation programme using the PROC GLM in the Statistical Analysis System (SAS Institute, Cary, NC), version 8 (6)(7).

52.   A final report shall include:

Testing facility:

Responsible personnel and their study responsibilities

Data from the Baseline Positive Control Test and periodic positive control data (see paragraphs 26 and 27)

Test chemical:

Characterisation of test chemicals

Physical nature and where relevant physicochemical properties

Method and frequency of preparation of dilutions

Any data generated on stability

Any analyses of dosing solutions


Characterisation of test vehicle (nature, supplier and lot)

Justification of choice of vehicle (if other than water)

Test animals:

Species and strain and justification for their choice

Supplier and specific supplier facility

Age on supply with birth date

If immature animals, whether or not supplied with dam or foster dam and date of weaning

Details of animal acclimatisation procedure

Number of animals per cage

Detail and method of individual animal and group identification

Assay Conditions:

Details of randomisation process (i.e. method used)

Rationale for dose selection

Details of test chemical formulation, its achieved concentrations, stability and homogeneity

Details of test chemical administration and rationale for the choice of exposure route

Diet (name, type, supplier, content, and, if known, phyto-oestrogen levels)

Water source (e.g. tap water or filtered water) and supply (by tubing from a large container, in bottles, etc.)

Bedding (name, type, supplier, content)

Record of caging conditions, lighting interval, room temperature and humidity, room cleaning

Detailed description of necropsy and uterine weighing procedures

Description of statistical procedures


For individual animals:

All daily individual body weights (from allocation into groups through necropsy) (to the nearest 0,1 g)

Age of each animal (in days counting day of birth as day 0) when administration of test chemical begins

Date and time of each dose administration

Calculated volume and dosage administered and observations of any dosage losses during or after administration

Daily record of status of animal, including relevant symptoms and observations

Suspected cause of death (if found during study in moribund state or dead)

Date and time of humane killing with time interval to last dosing

Wet uterine weight (to the nearest 0,1 mg) and any observations of luminal fluid losses during dissection and preparation for weighing

Blotted uterine weight (to the nearest 0,1 mg)

For each group of animals:

Mean daily body weights (to the nearest 0,1 g) and standard deviations (from allocation into groups through necropsy)

Mean wet uterine weights and mean blotted uterine weights (to the nearest 0,1 mg) and standard deviations

If measured, daily food consumption (calculated as grams of food consumed per animal)

The results of statistical analyses comparing both the wet and blotted uterine weights of treated groups relative to the same measures in the vehicle control groups.

The results of statistical analysis comparing the total body weight and the body weight gain of treated groups relative to the same measures in the vehicle control groups.

53.   Summary of the important guidance facts of the test method






Commonly used laboratory rodent strain

Number of animals

A minimum of 6 animals per dose group

Number of groups

A minimum of 2 test groups (see paragraph 33 for guidance) and a negative control group

For guidance on positive control groups see paragraphs 26 and 27

Housing and feeding conditions

T° in animal room

22 °C ± 3 °C

Relative humidity

50-60 % and not below 30 % or above 70 %

Daily lighting sequence

12 hours light, 12 hours dark

Diet and drinking water

Ad libitum


Individually or in groups of up to three animals (social group housing is recommended for immature animals)

Diet and bedding

Low level of phyto-oestrogens recommended in diet and bedding



Immature non-ovariectomised method (the preferred one).

Ovariectomised adult female method

Ovariectomised adult female method

Age of dosing for immature animals

PND 18 at the earliest. Dosing should be completed prior to PND 25

Not relevant under the scope of the current test method.

Age of ovariectomy

Between 6 and 8 weeks of age.

Age of dosing for ovariectomised animals

A minimum of 14 days should elapse between ovariectomy and the 1st day of administration.

A minimum of 7 days should elapse between ovariectomy and the 1st day of administration.

Body weight

Body weight variation should be minimal and not exceed ± 20 % of the mean weight.


Route of administration

Oral gavage or subcutaneous injection

Frequency of administration

Single daily dose

Volume amount for gavage and injection

≤ 5 ml/kg body weight (or up to 10 ml/kg body weight in case of aqueous solutions) (in 2 injection sites for subcutaneous route)

Duration of administration

3 consecutive days for immature model

Minimum of 3 consecutive days for the OVX model

7 consecutive days for the OVX model

Time of necropsy

Approximately 24 hours after the last dose


Positive response

Statistically significant increase of the mean uterus weight (wet and/or blotted)

Reference oestrogen

17α-ethinyl estradiol


54.   In general, a test for oestrogenicity should be considered positive if there is a statistically significant increase in uterine weight (p < 0,05) at least at the high dose level as compared to the solvent control group. A positive result is further supported by the demonstration of a biologically plausible relationship between the dose and the magnitude of the response, bearing in mind that overlapping oestrogenic and antioestrogenic activities of the test chemical may affect the shape of the dose-response curve.

55.   Care must be taken in order not to exceed the maximum tolerated dose to allow a meaningful interpretation of the data. Reduction of body weight, clinical signs and other findings should be thoroughly assessed in this respect.

56.   An important consideration for the acceptance of the data from the Uterotrophic Bioassay is the uterine weights of the vehicle control group. High control values may compromise the responsiveness of the bioassay and the ability to detect very weak oestrogen agonists. Literature reviews and the data generated during the validation of the Uterotrophic Bioassay suggest that instances of high control means do occur spontaneously, particularly in immature animals (2)(3)(6)(9). As the uterine weight of immature rats depends on many variables like strain or body weight, no definitive upper limit for the uterine weight can be given. As a guide, if blotted uterine weights in immature control rats are comprised between 40 and 45 mg, results should be considered as suspicious and uterine weights above 45 mg may lead to rerun the test. However, this needs to be considered on a case by case basis (3)(6)(8). When testing in adult rats incomplete ovariectomy will leave ovarian tissue that can produce endogenous oestrogen and retard the regression of the uterine weight.

57.   Blotted vehicle control uterine weights less than 0,09 % of body weight for immature female rats and less than 0,04 % for ovariectomised young adult females appear to yield acceptable results (see Table 31 (2)). If the control uterine weights are greater than these numbers, various factors should be scrutinised including the age of the animals, proper ovariectomy, dietary phyto-oestrogens, and so on, and a negative assay result (no indication for oestrogenic activity) should be used with caution.

58.   Historical data for vehicle control groups should be maintained in the laboratory. Historical data for responses to positive reference oestrogens, such as 17a-ethinyl estradiol, should also be maintained in the laboratory. Laboratories may also test the response to known weak oestrogen agonists. All these data can be compared to available data (2)(3)(4)(5)(6)(7)(8) to ensure that the laboratory's methods yield sufficient sensitivity.

59.   The blotted uterine weights showed less variability in the course of the OECD validation study than the wet uterine weights (6)(7). However, a significant response in either measure would indicate that the test chemical is positive for oestrogenic activity.

60.   The uterotrophic response is not entirely of oestrogenic origin, however, a positive result of the Uterotrophic Bioassay should generally be interpreted as evidence for oestrogenic potential in vivo, and should normally initiate actions for further clarification (see paragraph 9 and the “OECD Conceptual Framework for the Testing and Assessment of Endocrine Disrupting Chemicals”, Annex 2).

Figure 1

Schematic diagram showing the surgical removal of the ovaries

Image 2



Cut here



Mesometrium, vasculature and fat pad not shown

The procedure begins by opening dorso-lateral abdominal wall at the mid-point between the costal inferior border and the iliac crest, and a few millimetres lateral to the lateral margin of the lumbar muscle. Within the abdominal cavity, the ovaries should be located. On an aseptic field, the ovaries are then physically removed from the abdominal cavity, a ligature placed between the ovary and uterus to control bleeding, and the ovary detached by incision above the ligature at the junction of the oviduct and each uterine horn. After confirming that no significant bleeding persists, the abdominal wall should be closed by suture, and the skin closed, e.g. by autoclips or suture. The animals should be allowed to recover and the uterus weight to regress for a minimum of 14 days before use.

Figure 2

The removal and preparation of the uterine tissues for weight measurement.

Image 3


Disconnection line at necropsy

The procedure begins by opening the abdominal wall at the pubic symphysis. Then, each ovary, if present and uterine horn is detached from the dorsal abdominal wall. Urinary bladder and ureters are removed from the ventral and lateral side of uterus and vagina. Fibrous adhesion between the rectum and the vagina are detached until the junction of vaginal orifice and perineal skin can be identified. The uterus and vagina are detached from the body by incising the vaginal wall just above the junction between perineal skin as shown in the figure. The uterus should be detached from the body wall by gently cutting the uterine mesentery at the point of its attachment along the full length of the dorsolateral aspect of each uterine horn. After removal from the body, the excess fat and connective tissue is trimmed away. If ovaries are present, the ovaries are removed at the oviduct avoiding loss of luminal fluid from the uterine horn. If the animal has been ovarectomised, the stubs should be examined for the presence of any ovarian tissue. The vagina is removed from the uterus just below the cervix so that the cervix remains with the uterine body as shown in the figure. The uterus can then be weighed.

Appendix 1


Antioestrogenicity is the capability of a chemical to suppress the action of estradiol 17ß in a mammalian organism.

Chemical means a substance or a mixture.

Date of birth is postnatal day 0.

Dosage is a general term comprising of dose, its frequency and the duration of dosing.

Dose is the amount of test chemical administered. For the Uterotrophic Bioassay, the dose is expressed as weight of test chemical per unit body weight of test animal per day (e.g. mg/kg body weight/day).

Maximum Tolerable Dose (MTD) is the highest amount of a chemical that, when introduced into the body does not kill test animals (denoted by LD0) (IUPAC, 1993)

Oestrogenicity is the capability of a chemical to act like estradiol 17ß in a mammalian organism.

Postnatal day X is the Xth day of life after the day of birth.

Sensitivity is the proportion of all positive/active chemicals that are correctly classified by the test. It is a measure of accuracy for a test method that produces categorical results, and is an important consideration in assessing the relevance of a test method.

Specificity is the proportion of all negative/inactive chemicals that are correctly classified by the test. It is a measure of accuracy for a test method that produces categorical results and is an important consideration in assessing the relevance of a test method.

Test chemical means any substance or mixture tested using this test method.

Uterotrophic is a term used to describe a positive influence on the growth of uterine tissues.

Validation is a scientific process designed to characterise the operational requirements and limitations of a test method and to demonstrate its reliability and relevance for a particular purpose.

Appendix 2