Council 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) (Text with EEA relevance)

• 43.The correlation coefficient (R²) for calibration curves should be greater than 0,99. However, a high correlation is not sufficient to guarantee adequate prediction of concentration in all ranges. In addition to having a sufficiently high correlation for the calibration curve, the concentration of each standard, as calculated from the calibration curve, should all fall between 70 and 120 % of its nominal concentration. If the nominal concentrations trend away from the calibration regression line (e.g. at lower concentrations), it may be necessary to split the calibration curve into low and high ranges or to use a nonlinear model to adequately fit the absorbance data. If the curve is split, both line segments should have R² > 0,99.

• 44.The limit of detection (LOD) is defined as the concentration of the lowest analytical standard, and limit of quantitation (LOQ) is defined as the concentration of the lowest analytical standard multiplied by the lowest dilution factor.

• 45.On each day that vitellogenin assays are performed, a fortification sample made using an inter-assay reference standard will be analysed (Appendix 7). The ratio of the expected concentration to the measured concentration will be reported along with the results from each set of assays performed on that day.

DATA AND REPORTING

Evaluation of Biomarker Responses by Analysis of Variance (ANOVA)

• 46.To identify potential endocrine activity of a chemical, responses are compared between treatments and control groups using analysis of variance (ANOVA). Where a solvent control is used, an appropriate statistical test should be performed between the dilution water and solvent controls for each endpoint. Guidance on how to handle dilution water and solvent control data in the subsequent statistical analysis can be found in OECD, 2006c (27). All biological response data should be analysed and reported separately by sex. If the required assumptions for parametric methods are not met — non-normal distribution (e.g. Shapiro-Wilk's test) or heterogeneous variance (Bartlett's test or Levene's test), consideration should be given to transforming the data to homogenise variances prior to performing the ANOVA, or to carrying out a weighted ANOVA. Dunnett's test (parametric) on multiple pair-wise comparisons or a Mann-Whitney with Bonferroni adjustment (non-parametric) may be used for non-monotonous dose-response. Other statistical tests may be used (e.g. Jonckheere-Terpstra test or Williams test) if the dose-response is approximately monotone. A statistical flowchart is provided in Appendix 8 to help in the decision on the most appropriate statistical test to be used. Additional information can also be obtained from the OECD Document on Current Approaches to Statistical Analysis of Ecotoxicity Data (27).

Reporting of test results

• 47.Study data should include:

Testing facility:

• — Responsible personnel and their study responsibilities

• — Each laboratory should have demonstrated proficiency using a range of representative chemicals

Test chemical:

• — Characterisation of test chemical

• — Physical nature and relevant physicochemical properties

• — Method and frequency of preparation of test concentrations

• — Information on stability and biodegradability

Solvent:

• — Characterization of solvent (nature, concentration used)

• — Justification of choice of solvent (if other than water)

Test animals:

• — Species and strain

• — Supplier and specific supplier facility

• — Age of the fish at the start of the test and reproductive/spawning status

• — Details of animal acclimation procedure

• — Body weight of the fish at the start of the exposure (from a sub-sample of the fish stock)

Test Conditions:

• — Test procedure used (test-type, loading rate, stocking density, etc.);

• — Method of preparation of stock solutions and flow-rate;

• — The nominal test concentrations, weekly measured concentrations of the test solutions and analytical method used, means of the measured values and standard deviations in the test vessels and evidence that the measurements refer to the concentrations of the test chemical in true solution;

• — Dilution water characteristics (including pH, hardness, alkalinity, temperature, dissolved oxygen concentration, residual chlorine levels, total organic carbon, suspended solids and any other measurements made)

• — Water quality within test vessels: pH, hardness, temperature and dissolved oxygen concentration;

• — Detailed information on feeding (e.g. type of food(s), source, amount given and frequency and analyses for relevant contaminants if available (e.g. PCBs, PAHs and organochlorine pesticides).

Results

• — Evidence that the controls met the acceptance criteria of the test;

• — Data on mortalities occurring in any of the test concentrations and control;

• — Statistical analytical techniques used, treatment of data and justification of techniques used;

• — Data on biological observations of gross morphology, including secondary sex characteristics and vitellogenin;

• — Results of the data analyses preferably in tabular and graphical form;

• — Incidence of any unusual reactions by the fish and any visible effects produced by the test chemical

GUIDANCE FOR THE INTERPRETATION AND ACCEPTANCE OF THE TEST RESULTS

• 48.This section contains a few considerations to be taken into account in the interpretation of test results for the various endpoints measured. The results should be interpreted with caution where the test chemical appears to cause overt toxicity or to impact on the general condition of the test animal.

• 49.In setting the range of test concentrations, care should be taken not to exceed the maximum tolerated concentration to allow a meaningful interpretation of the data. It is important to have at least one treatment where there are no signs of toxic effects. Signs of disease and signs of toxic effects should be thoroughly assessed and reported. For example, it is possible that production of VTG in females can also be affected by general toxicity and non-endocrine toxic modes of action, e.g. hepatotoxicity. However, interpretation of effects may be strengthened by other treatment levels that are not confounded by systemic toxicity.

• 50.There are a few aspects to consider for the acceptance of test results. As a guide, the VTG levels in control groups of males and females should be distinct and separated by about three orders of magnitude in fathead minnow and zebrafish, and about one order of magnitude for medaka. Examples of the range of values encountered in control and treatment groups are available in the validation reports (1, 2, 3, 4). High VTG values in control males could compromise the responsiveness of the assay and its ability to detect weak oestrogen agonists. Low VTG values in control females could compromise the responsiveness of the assay and its ability to detect aromatase inhibitors and oestrogen antagonists. The validation studies were used to build that guidance.

• 51.If a laboratory has not performed the assay before or substantial changes (e.g. change of fish strain or supplier) have been made it is advisable that a technical proficiency study is conducted. It is recommended that chemicals covering a range of modes of action or impacts on a number of the test endpoints are used. In practice, each laboratory is encouraged to build its own historical control data for males and females and to perform a positive control chemical for estrogenic activity (e.g. 17β-estradiol at 100 ng/l, or a known weak agonist) resulting in increased VTG in male fish, a positive control chemical for aromatase inhibition (e.g. fadrozole or prochloraz at 300 μg/l) resulting in decreased VTG in female fish, and a positive control chemical for androgenic activity (e.g. 17β-trenbolone at 5 μg/l) resulting in induction of secondary sex characteristics in female fathead minnow and medaka. All these data can be compared to available data from the validation studies (1, 2, 3) to ensure laboratory proficiency.

• 52.In general, vitellogenin measurements should be considered positive if there is a statistically significant increase in VTG in males (p < 0,05), or a statistically significant decrease in females (p < 0,05) at least at the highest dose tested compared to the control group, and in the absence of signs of general toxicity. A positive result is further supported by the demonstration of a biologically plausible relationship between the dose and the response curve. As mentioned earlier, the vitellogenin decrease may not entirely be of endocrine origin; however a positive result should generally be interpreted as evidence of endocrine activity in vivo, and should normally initiate actions for further clarification.

LITERATURE

• (1)OECD (2006a). Report of the Initial Work Towards the Validation of the 21-Day Fish Screening Assay for the Detection of Endocrine active Substances (Phase 1A). OECD Environmental Health and Safety Publications Series on Testing and Assessment No.60, ENV/JM/MONO(2006)27.

• (2)OECD (2006b). Report of the Initial Work Towards the Validation of the 21-Day Fish Screening Assay for the Detection of Endocrine active Substances (Phase 1B). OECD Environmental Health and Safety Publications Series on Testing and Assessment No.61, ENV/JM/MONO(2006)29.

• (3)OECD (2007). Final report of the Validation of the 21-day Fish Screening Assay for the Detection of Endocrine Active Substances. Phase 2: Testing Negative Substances. OECD Environmental Health and Safety Publications Series on Testing and Assessment No.78, ENV/JM/MONO(2007)25.

• (4)Owens JW (2007). Phase 3 report of the validation of the OECD Fish Screening Assay. CEFIC LRI Project, Endocrine. http://www.cefic-lri.org/index.php?page=projects (accessed 18/09/08).

• (5)US EPA 2007. Validation of the Fish Short-Term Reproduction Assay: Integrated Summary Report. Unpublished report dated 15 December 2007. US Environmental Protection Agency, Washington, DC. 104 pp.

• (6)OECD, 2008. Report of the Validation Peer Review for the 21-Day Fish Endocrine Screening Assay and Agreement of the Working Group of the National Coordinators of the Test Guidelines Programme on the Follow-up of this Report. OECD Environmental Health and Safety Publications Series on Testing and Assessment No.94, ENV/JM/MONO(2008)21.

• (7)Sumpter and Jobling (1995). Vitellogenesis as a biomarker for estrogenic contamination of the aquatic environment. Environmental Health Perspectives;103 Suppl 7:173-8 Review.

• (8)Pawlowski S, Sauer A, Shears JA, Tyler CR, Braunbeck T (2004). Androgenic and estrogenic effects of the synthetic androgen 17alpha-methyltestosterone on sexual development and reproductive performance in the fathead minnow (Pimephales promelas) determined using the gonadal recrudescence assay. Aquatic Toxicology; 68(3):277-91.

• (9)Andersen L, Goto-Kazato R, Trant JM, Nash JP, Korsgaard B, Bjerregaard P (2006). Short-term exposure to low concentrations of the synthetic androgen methyltestosterone affects vitellogenin and steroid levels in adult male zebrafish (Danio rerio). Aquatic Toxicology; 76(3-4):343-52.

• (10)Ankley GT, Kahl MD, Jensen KM, Hornung MW, Korte JJ, Makynen EA, Leino RL (2002). Evaluation of the aromatase inhibitor fadrozole in a short-term reproduction assay with the fathead minnow (Pimephales promelas). Toxicological Sciences;67(1):121-30.

• (11)Panter GH, Hutchinson TH, Hurd KS, Sherren A, Stanley RD, Tyler CR (2004). Successful detection of (anti-)androgenic and aromatase inhibitors in pre-spawning adult fathead minnows (Pimephales promelas) using easily measured endpoints of sexual development. Aquatic Toxicology; 70(1):11-21.

• (12)Parks LG, Cheek AO, Denslow ND, Heppell SA, McLachlan JA, LeBlanc GA, Sullivan CV (1999). Fathead minnow (Pimephales promelas) vitellogenin: purification, characterization and quantitative immunoassay for the detection of estrogenic compounds. Comparative Biochemistry and Physiology. Part C Pharmacology, toxicology and endocrinology; 123(2):113-25.

• (13)Panter GH, Tyler CR, Maddix S, Campbell PM, Hutchinson TH, Länge R, Lye C, Sumpter JP, 1999. Application of an ELISA to quantify vitellogenin concentrations in fathead minnows (Pimephales promelas) exposed to endocrine disrupting chemicals. CEFIC-EMSG research report reference AQ001. CEFIC, Brussels, Belgium.

• (14)Fenske M., van Aerle, R.B., Brack, S.C., Tyler, C.R., Segner, H., (2001). Development and validation of a homologous zebrafish (Danio rerio Hamilton- Buchanan) vitellogenin enzyme-linked immunosorbent assay (ELISA) and its application for studies on estrogenic chemicals. Comp. Biochem. Phys. C 129 (3): 217-232.

• (15)Holbech H, Andersen L, Petersen GI, Korsgaard B, Pedersen KL, Bjerregaard P. (2001). Development of an ELISA for vitellogenin in whole body homogenate of zebrafish (Danio rerio). Comparative Biochemistry and Physiology. Part C Pharmacology, toxicology and endocrinology; 130: 119-131

• (16)Rose J, Holbech H, Lindholst C, Noerum U, Povlsen A, Korsgaard B, Bjerregaard P. 2002. Vitellogenin induction by 17β-estradiol and 17β-ethinylestradiol in male zebrafish (Danio rerio). Comp. Biochem. Physiol. C. 131: 531-539.

• (17)Brion F, Nilsen BM, Eidem JK, Goksoyr A, Porcher JM, Development and validation of an enzyme-linked immunosorbent assay to measure vitellogenin in the zebrafish (Danio rerio). Environmental Toxicology and Chemistry; vol 21: 1699-1708.

• (18)Yokota H, Morita H, Nakano N, Kang IJ, Tadokoro H, Oshima Y, Honjo T, Kobayashi K. 2001. Development of an ELISA for determination of the hepatic vitellogenin in Medaka (Oryzias latipes). Jpn J Environ Toxicol 4:87–98.

• (19)Tatarazako N, Koshio M, Hori H, Morita M and Iguchi T., 2004. Validation of an enzyme-linked immunosorbent assay method for vitellogenin in the Medaka. Journal of Health Science 50:301-308.

• (20)Ankley GT, Jensen KM, Makynen EA, Kahl MD, Korte JJ, Homung MW, Henry TR, Denny JS, Leino RL, Wilson VS, Cardon MC, Hartig PC, Gray LE (2003). Effects of the androgenic growth promoter 17-beta-trenbolone on fecundity and reproductive endocrinology of the fathead minnow. Environmental Toxicology and Chemistry; 22(6): 1350-60.

• (21)Seki M, Yokota H, Matsubara H, Maeda M, Tadokoro H, Kobayashi K (2004). Fish full life-cycle testing for androgen methyltestosterone on medaka (Oryzias latipes). Environmental Toxicology and Chemistry; 23(3):774-81.

• (22)OECD (2000) Guidance Document on Aquatic Toxicity Testing of Difficult Substances and Mixtures. Environmental Health and Safety Publications. Series on Testing and Assessment. No. 23. Paris

• (23)Hutchinson TH, Shillabeer N, Winter MJ, Pickford DB, 2006a. Acute and chronic effects of carrier solvents in aquatic organisms: A critical review. Review. Aquatic Toxicology, 76; pp.69–92.

• (24)Hutchinson TH, Ankley GT, Segner H, Tyler CR, 2006b. Screening and testing for endocrine disruption in fish-biomarkers as ‘signposts’, not ‘traffic lights’, in risk assessment. Environmental Health Perspectives;114 Suppl 1:106-14.

• (25)Miles-Richardson, SR, Kramer VJ, Fitzgerald SD, Render JA, Yamini B, Barbee SJ, Giesy JP. 1999. Effects of waterborne exposure to 17β-estradiol on secondary sex characteristics and gonads of the fathead minnow (Pimephales promelas). Aquat. Toxicol. 47, 129-145.

• (26)Martinovic, D., L.S. Blake, E.J. Durhan, K.J. Greene, M.D. Kahl, K.M., Jensen, E.A. Makynen, D.L. Villeneuve and G.T. Ankley. 2008. Characterization of reproductive toxicity of vinclozolin in the fathead minnow and co-treatment with an androgen to confirm an anti-androgenic mode of action. Environ. Toxicol. Chem. 27, 478-488.

• (27)OECD (2006c). Current Approaches in the Statistical Analysis of Ecotoxicity Data: A Guidance to Application. OECD environmental Health and Safety Publications Series on Testing and Assessment No.54. ENV/JM/MONO(2006)18

• (28)OECD (2012) OECD Conceptual Framework for Testing and Assessment of Endocrine Disrupters (revised). Annex I to Draft Guidance Document on Standardised Test Guidelines for Evaluating Chemicals for Endocrine Disruption. Series on Testing and Assessment No 150. ENV/JM/MONO(2012)22

Appendix 1

Abbreviations & definitions

Chemical : A substance or a mixture

CV : Coefficient of variation.

ELISA : Enzyme-Linked Immunosorbent Assay.

Loading rate : Wet weight of fish per volume of water.

Stocking density : Number of fish per volume of water.

VTG (Vitellogenin) : Phospholipoglycoprotein precursor to egg yolk protein that normally occurs in sexually active females of all oviparous species.

HPG axis : Hypothalamic-pituitary-gonadal axis.

MTC : Maximum Tolerated Concentration, representing about 10 % of the LC50.

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

Appendix 2

Experimental conditions for the fish endocrine screening assay

1. Recommended species	Fathead minnow (Pimephales promelas)	Medaka (Oryzias latipes)	Zebrafish (Danio rerio)
2. Test type	Flow-through	Flow-through	Flow-through
3. Water temperature	25 ± 2 °C	25 ± 2 °C	26 ± 2 °C
4. Illumination quality	Fluorescent bulbs (wide spectrum)	Fluorescent bulbs (wide spectrum)	Fluorescent bulbs (wide spectrum)
5. Light intensity	10-20 μE/m2/s, 540-1 000 lux, or 50-100 ft-c (ambient laboratory levels)	10-20 μE/m2/s, 540-1 000 lux, or 50-100 ft-c (ambient laboratory levels)	10-20 μE/m2/s, 540-1 000 lux, or 50-100 ft-c (ambient laboratory levels)
6. Photoperiod (dawn/dusk transitions are optional, however not considered necessary)	16 h light, 8 h dark	12-16 h light, 12-8 h dark	12-16 h light, 12-8 h dark
7. Loading rate	< 5 g per l	< 5 g per l	< 5 g per l
8. Test chamber size	10 l (minimum)	2 l (minimum)	5 l (minimum)
9. Test solution volume	8 l (minimum)	1.5 l (minimum)	4 l (minimum)
10. Volume exchanges of test solutions	Minimum of 6 daily	Minimum of 5 daily	Minimum of 5 daily
11. Age of test organisms	See paragraph 20	See paragraph 20	See paragraph 20
12. Approximate wet weight of adult fish (g)	Females: 1,5 ± 20 % Males: 2,5 ± 20 %	Females: 0,35 ± 20 % Males: 0,35 ± 20 %	Females: 0,65 ± 20 % Males: 0,4 ± 20 %
13. No. of fish per test vessel	6 (2 males and 4 females)	10 (5 males and 5 females)	10 (5 males and 5 females)
14. No. of treatments	= 3 (plus appropriate controls)	= 3 (plus appropriate controls)	= 3 (plus appropriate controls)
15. No. vessels per treatment	4 minimum	2 minimum	2 minimum
16. No. of fish per test concentration	16 adult females and 8 males (4 females and 2 males in each replicate vessel)	10 adult females and 10 males (5 females and 5 males in each replicate vessel)	10 adult females and 10 males (5 females and 5 males in each replicate vessel)
17. Feeding regime	Live or frozen adult or nauplii brine shrimp two or three times daily (ad libitum), commercially available food or a combination of the above	Brine shrimp nauplii two or three times daily (ad libitum), commercially available food or a combination of the above	Brine shrimp nauplii two or three times daily (ad libitum), commercially available food or a combination of the above
18. Aeration	None unless DO concentration falls below 60 % air saturation	None unless DO concentration falls below 60 % air saturation	None unless DO concentration falls below 60 % air saturation
19. Dilution water	Clean surface, well or reconstituted water or dechlorinated tap water	Clean surface, well or reconstituted water or dechlorinated tap water	Clean surface, well or reconstituted water or dechlorinated tap water
20. Pre-exposure period	7 days recommended	7 days recommended	7 days recommended
21. Chemical exposure duration	21 d	21 d	21 d
22. Biological endpoints	survival behaviour 2y sex characteristics VTG	survival behaviour 2y sex characteristics VTG	survival behaviour VTG
23. Test acceptability	Dissolved oxygen > 60 % of saturation; mean temperature of 25 ± 2 °C; 90 % survival of fish in the controls; measured test concentrations within 20 % of mean measured values per treatment level.	Dissolved oxygen > 60 % of saturation; mean temperature of 24 ± 2 °C; 90 % survival of fish in the controls; measured test concentrations within 20 % of mean measured values per treatment level.	Dissolved oxygen > 60 % of saturation; mean temperature of 26 ± 2 °C; 90 % survival of fish in the controls; measured test concentrations within 20 % of mean measured values per treatment level.

Appendix 3

Some chemical characteristics of acceptable dilution water

Component	Concentrations
Particulate matter	< 20 mg/l
Total organic carbon	< 2 mg/l
Unionised ammonia	< 1 μg/l
Residual chlorine	< 10 μg/l
Total organophosphorus pesticides	< 50 ng/l
Total organochlorine pesticides plus polychlorinated biphenyls	< 50 ng/l
Total organic chlorine	< 25 ng/l

Appendix 4A

Spawning substrate for zebrafish

Spawning tray : all glass instrument dish, for example 22 × 15 × 5,5 cm (l × w × d), covered with a removable stainless steel wire lattice (mesh width 2 mm). The lattice should cover the opening of the instrument dish at a level below the brim.

Image omitted; see official source

On the lattice, spawning substrate should be fixed. It should provide structure for the fish to move into. For example, artificial aquaria plants made of green plastic material are suitable (NB: possible adsorption of the test chemical to the plastic material should be considered). The plastic material should be leached out in sufficient volume of warm water for sufficient time to ensure that no chemicals may be disposed to the test water. When using glass materials it should be ensured that the fish are neither injured nor cramped during their vigorous actions.

The distance between the tray and the glass panes should be at least 3 cm to ensure that the spawning is not performed outside the tray. The eggs spawned onto the tray fall through the lattice and can be sampled 45-60 min after the start of illumination. The transparent eggs are non-adhesive and can easily be counted by using transversal light. When using five females per vessel, egg numbers up to 20 at a day can be regarded as low, up to 100 as medium and more than 100 as high numbers. The spawning tray should be removed, the eggs collected and the spawning tray re-introduced in the test vessel, either as late as possible in the evening or very early in the morning. The time until re-introduction should not exceed one hour since otherwise the cue of the spawning substrate may induce individual mating and spawning at an unusual time. If a situation needs a later introduction of the spawning tray, this should be done at least 9 hours after start of the illumination. At this late time of the day, spawning is not induced any longer.

Appendix 4B

Spawning substrate for fathead minnow

Two or three combined plastic/ceramic/glass or stainless steel spawning tiles and trays are placed in each of the test chamber (e.g., 80 mm length of grey semi-circular guttering sitting on a lipped tray of 130mm length) (see picture). Properly seasoned PVC or ceramic tiles have demonstrated to be appropriate for a spawning substrate (Thorpe et al, 2007).

It is recommended that the tiles are abraded to improve adhesion. The tray should also be screened to prevent fish from access to the fallen eggs unless the egg adhesion efficiency has been demonstrated for the spawning substrate used.

Image omitted; see official source

The base is designed to contain any eggs that do not adhere to the tile surface and would therefore fall to the bottom of the tank (or those eggs laid directly onto the flat plastic base). All spawning substrates should be leached for a minimum of 12 hours, in dilution water, before use.

REFERENCES

Thorpe KL, Benstead R, Hutchinson TH, Tyler CR, 2007. An optimised experimental test procedure for measuring chemical effects on reproduction in the fathead minnow, Pimephales promelas. Aquatic Toxicology, 81, 90–98.

Appendix 5A

Assessment of secondary sex characteristics in fathead minnow for the detection of certain endocrine active chemicals

Overview

Potentially important characteristics of physical appearance in adult fathead minnows in endocrine disrupter testing include body colour (i.e. light/dark), coloration patterns (i.e. presence or absence of vertical bands), body shape (i.e. shape of head and pectoral region, distension of abdomen), and specialized secondary sex characteristics (i.e. number and size of nuptial tubercles, size of dorsal pad and ovipositor).

Nuptial tubercles are located on the head (dorsal pad) of reproductively-active male fathead minnows, and are usually arranged in a bilaterally-symmetric pattern (Jensen et al. 2001). Control females and juvenile males and females exhibit no tubercle development (Jensen et al. 2001). There can be up to eight individual tubercles around the eyes and between the nares of the males. The greatest numbers and largest tubercles are located in two parallel lines immediately below the nares and above the mouth. In many fish there are groups of tubercles below the lower jaw; those closest to the mouth generally occur as a single pair, while the more ventral set can be comprised of up to four tubercles. The actual numbers of tubercles is seldom more than 30 (range, 18-28; Jensen et al. 2001). The predominant tubercles (in terms of numbers) are present as a single, relatively round structure, with the height approximately equivalent to the radius. Most reproductively-active males also have, at least some, tubercles which are enlarged and pronounced such that they are indistinguishable as individual structures.

Some types of endocrine-disrupting chemicals can cause the abnormal occurrence of certain secondary sex characteristics in the opposite sex; for example, androgen receptor agonists, such as 17β-methyltestosterone or 17β-trenbolone, can cause female fathead minnows to develop nuptial tubercles (Smith 1974; Ankley et al. 2001; 2003), while oestrogen receptor agonists may decrease number or size of nuptial tubercles in males (Miles-Richardson et al. 1999; Harries et al. 2000).

Below is a description of the characterization of nuptial tubercles in fathead minnows based on procedures used at the U.S. Environmental Protection Agency lab in Duluth, MN. Specific products and/or equipment can be substituted with comparable materials available.

Viewing is best accomplished using an illuminated magnifying glass or 3X illuminated dissection scope. View fish dorsally and anterior forward (head toward viewer).

• a) Place fish in small Petri dish (e.g., 100 mm in diameter), anterior forward, and ventral down. Focus viewfinder to allow identification of tubercles. Gently and slowly roll fish from side to side to identify tubercle areas. Count and score tubercles.

• b) Repeat the observation on the ventral head surface by placing the fish dorsal anterior forward in the Petri dish.

• c) Observations should be completed within 2 min for each fish.

Tubercle Counting and Rating

Six specific areas have been identified for assessment of tubercle presence and development in adult fathead minnows. A template was developed to map the location and quantity of tubercles present (see end of this Appendix). The number of tubercles is recorded and their size can be quantitatively ranked as: 0- absence, 1-present, 2-enlarged and 3-pronounced for each organism (Fig. 1).

Rate 0- absence of any tubercle. Rating 1-present, is identified as any tubercle having a single point whose height is nearly equivalent to its radius (diameter). Rating 2- enlarged, is identified by tissue resembling an asterisk in appearance, usually having a large radial base with grooves or furrows emerging from the centre. Tubercle height is often more jagged but can be somewhat rounded at times. Rating 3- pronounced, is usually quite large and rounded with less definition in structure. At times these tubercles will run together forming a single mass along an individual or combination of areas (B, C and D, described below). Coloration and design are similar to rating 2 but at times are fairly indiscriminate. Using this rating system generally will result in overall tubercle scores of < 50 in a normal control male possessing a tubercle count of 18 to 20 (Jensen et al. 2001).

Figure 1

Image omitted; see official source

The actual number of tubercles in some fish may be greater than the template boxes (Appendix A) for a particular rating area. If this happens, additional rating numbers may be marked within, to the right or to the left of the box. The template therefore does not need to display symmetry. An additional technique for mapping tubercles which are paired or joined vertically along the horizontal plane of the mouth could be done by double-marking two tubercle rating points in a single box.

Mapping regions:

A — Tubercles located around eye. Mapped dorsal to ventral around anterior rim of eye. Commonly multiple in mature control males, not present in control females, generally paired (one near each eye) or single in females exposed to androgens.

B — Tubercles located between nares, (sensory canal pores). Normally in pairs for control males at more elevated levels (2- enlarged or 3- pronounced) of development. Not present in control females with some occurrence and development in females exposed to androgens.

C — Tubercles located immediately anterior to nares, parallel to mouth. Generally enlarged or pronounced in mature control males. Present or enlarged in less developed males or androgen-treated females.

D — Tubercles located parallel along mouth line. Generally rated developed in control males. Absent in control females but present in androgen-exposed females.

E — Tubercles located on lower jaw, close to mouth, usually small and commonly in pairs. Varying in control or treated males, and treated females.

F — Tubercles located ventral to E. Commonly small and paired. Present in control males and androgen-exposed females.

REFERENCES

• (1)Ankley GT, Jensen KM, Kahl MD, Korte JJ, Makynen ME. 2001. Description and evaluation of a short-term reproduction test with the fathead minnow (Pimephales promelas). Environ Toxicol Chem 20:1276-1290.

• (2)Ankley GT, Jensen KM, Makynen EA, Kahl MD, Korte JJ, Hornung MW, Henry TR, Denny JS, Leino RL, Wilson VS, Cardon MC, Hartig PC, Gray EL. 2003. Effects of the androgenic growth promoter 17-β trenbolone on fecundity and reproductive endocrinology of the fathead minnow. Environ Toxicol Chem 22:1350-1360.

• (3)Harries JE, Runnalls T, Hill E, Harris CA,Maddix S, Sumpter JP, Tyler CR. 2000. Development of a reproductive performance test for endocrine disrupting chemicals using pair-breeding fathead minnows (Pimephales promelas). Environ Sci Technol 34:3003-3011.

• (4)Jensen KM, Korte JJ, Kahl MD, Pasha MS, Ankley GT. 2001. Aspects of basic reproductive biology and endocrinology in the fathead minnow (Pimephales promelas). Comp Biochem Physiol C 128:127-141.

• (5)Kahl MD, Jensen KM, Korte JJ, Ankley GT. 2001. Effects of handling on endocrinology and reproductive performance of the fathead minnow. J Fish Biol 59:515-523.

• (6)Miles-Richardson SR, Kramer VJ, Fitzgerald SD, Render JA, Yamini B, Barbee SJ, Giesy JP. 1999. Effects of waterborne exposure of 17-estradiol on secondary sex characteristics and gonads of fathead minnows (Pimephales promelas). Aquat Toxicol 47:129-145.

• (7)Smith RJF. 1974. Effects of 17-methyltestosterone on the dorsal pad and tubercles of fathead minnows (Pimephales promelas). Can J Zool 52:1031-1038.

Tubercle Template	Numerical Rating
ID	1-present
Date	2-enlarged
Total Score	3-pronounced
---	---
	A
---	---
	B
---	---
	C
	D
---	---
	F

Appendix 5B

Assessment of secondary sex characteristics in medaka for the detection of certain endocrine active chemicals

Below is a description of the measurement of papillary processes ([^E0199]), which are the secondary sex characteristics in medaka (Oryzias latipes).

• (1)After the excision of the liver (Appendix 6), the carcass is placed into a conical tube containing about 10 ml of 10 % neutral buffered formalin (upside: head, downside: tail). If the gonad is fixed in a solution other than 10 % neutral buffered formalin, make a transverse cut across the carcass between anterior region of anal fin and anus using razor, taking care not to harm the gonopore and gonad itself (Fig. 3). Place the cranial side of the fish body into the fixative solution to preserve the gonad, and the tail side of the fish body into the 10 % neutral buffered formalin as described above.

• (2)After placing the fish body into 10 % neutral buffered formalin, grasp the anterior region of the anal fin with tweezers and fold it for about 30 seconds to keep the anal fin open. When grasping the anal fin with tweezers, grasp a few fin rays in the anterior region with care not to scratch the papillary processes.

• (3)After keeping the anal fin open for about 30 seconds, store the fish body in 10 % neutral buffered formalin at room temperature until the measurement of the papillary processes (measurement should be conducted after fixing for at least 24 hours).

Measurement

• (1)After fixing the fish body in the 10 % neutral buffered formalin for at least 24 hours, pick up the fish carcass from the conical tube and wipe the formalin on the filter paper (or paper towel).

• (2)Place the fish abdomen side up. Then cut the anal fin using small dissection scissors carefully (it is preferable to cut the anal fin with small amount of pterygiophore).

• (3)Grasp the anterior region of the severed anal fin with tweezers and put it on a glass slide with a several drops of water. Then cover the anal fin with a cover glass. Be careful not to scratch the papillary processes when grasping the anal fin with tweezers.

• (4)Count the number of the joint plate with papillary processes using the counter under a biological microscope (upright microscope or inverted microscope). The papillary processes are recognized when a small formation of processes is visible on the posterior margin of joint plate. Write the number of joint plate with papillary processes in each fin ray to the worksheet (e.g. first fin ray: 0, second fin ray: 10, third fin ray: 12, etc.) and enter the sum of this number on the Excel spreadsheet by individual fish. If necessary, take a photograph of the anal fin and count the number of joint plate with papillary processes on the photograph.

• (5)After the measurement, put the anal fin into the conical tube described in (1) and store it.

Fig.1.

Diagram showing sexual difference in shape and size of the anal fin. A, male; B, female. Oka, T. B., 1931. On the processes on the fin rays of the male of Oryzias latipes and other sex characters of this fish. J. Fac. Sci., Tokyo Univ., IV, 2: 209-218.

Image omitted; see official source

Fig.2.A.

Processes on joint plates of anal fin-ray. J.P., joint plate; A.S., axial space; P., process. B, Distal extremity of fin-ray. Actinotrichia (Act.) are on the tip. Oka, T. B., 1931. On the processes on the fin rays of the male of Oryzias latipes and other sex characters of this fish. J. Fac. Sci., Tokyo Univ., IV, 2: 209-218.

Image omitted; see official source

Fig.3.

Photograph of fish body showing the cut site when the gonad is fixed in the fixing solution other than 10 % neutral buffered formalin. In that case, the remaining body will be cut off between anterior region of anal fin and anal using razor (red bar), and the head side of fish body will be put into the fixing solution for gonad and the tail side of the fish body will be put into the 10 % neutral buffered formalin.

Image omitted; see official source

Appendix 6

Recommended procedures for sample collection for vitellogenin analysis

Care should be taken to avoid cross-contamination between VTG samples of males and females.

Procedure 1A: Fathead Minnow, Blood Collection from the Caudal Vein/Artery

After anaesthetisation, the caudal peduncle is partially severed with a scalpel blade and blood is collected from the caudal vein/artery with a heparinised microhematocrit capillary tube. After the blood has been collected, the plasma is quickly isolated by centrifugation for 3 min at 15 000 g (or alternatively for 10 min. at 15 000 g at 4 °C). If desired, percent hematocrit can be determined following centrifugation. The plasma portion is then removed from the microhematocrit tube and stored in a centrifuge tube with 0,13 units of aprotinin (a protease inhibitor) at – 80 °C until determination of vitellogenin can be made. Depending on the size of the fathead minnow (which is sex-dependent), collectable plasma volumes generally range from 5 to 60 microlitres per fish (Jensen et al. 2001).

Procedure 1B: Fathead Minnow, Blood Collection from Heart

Alternatively, blood may also be collected by cardiac puncture using a heparinized syringe (1 000 units of heparin per ml). The blood is transferred into Eppendorf tubes (held on ice) and then centrifuged (5 min, 7 000 g, room temperature). The plasma should be transferred into clean Eppendorf tubes (in aliquots if the volume of plasma makes this feasible) and promptly frozen at – 80 °C, until analyzed (Panter et al., 1998).

Procedure 2A: Japanese Medaka, Excision of the Liver in Medaka

Removal of the test fish from the test chamber

• (1)Test fish should be removed from the test chamber using the small spoon-net. Be careful not to drop the test fish into other test chambers.

• (2)In principle, the test fish should be removed in the following order: control, solvent control (where appropriate), lowest concentration, middle concentration, highest concentration and positive control. In addition, all males should be removed from one test chamber before the remaining females are removed.

• (3)The sex of each test fish is identified on the basis of external secondary sex characteristics (e.g. the shape of the anal fin).

• (4)Place the test fish in a container for transport and carry it to the workstation for excision of the liver. Check the labels of the test chamber and the transport container for accuracy and to confirm that the number of fish that have been removed from the test chamber and that the number of fish remaining in the test chamber are consistent with expectation.

• (5)If the sex cannot be identified by the fish's external appearance, remove all fish from the test chamber. In this case, the sex should be identified by observing the gonad or secondary sex characteristics under a stereoscopic microscope.

Excision of the liver

• (1)Transfer the test fish from the container for transport to the anaesthetic solution using the small spoon-net.

• (2)After the test fish is anaesthetised, transfer the test fish on the filter paper (or a paper towel) using tweezers (commodity type). When grasping the test fish, apply the tweezers to the sides of the head to prevent breaking the tail.

• (3)Wipe the water on the surface of the test fish on the filter paper (or the paper towel).

• (4)Place the fish abdomen side up. Then make a small transverse incision partway between the ventral neck region and the mid-abdominal region using dissection scissors.

• (5)Insert the dissection scissors into the small incision, and incise the abdomen from a point caudal to the branchial mantle to the cranial side of the anus along the midline of the abdomen. Be careful not to insert the dissection scissors too deeply so as to avoid damaging the liver and gonad.

• (6)Conduct the following operations under the stereoscopic microscope.

• (7)Place the test fish abdomen side up on the paper towel (glass Petri dish or slide glass are also available).

• (8)Extend the walls of the abdominal cavity with precision tweezers and exteriorise the internal organs. It is also acceptable to exteriorise the internal organs by removing one side of the wall of the abdominal cavity if necessary.

• (9)Expose the connected portion of the liver and gallbladder using another pair of precision tweezers. Then grasp the bile duct and cut off the gallbladder. Be careful not to break the gallbladder.

• (10)Grasp the oesophagus and excise the gastrointestinal tract from the liver in the same way. Be careful not to leak the contents of the gastrointestinal tract. Excise the caudal gastrointestinal tract from the anus and remove the tract from the abdominal cavity.

• (11)Trim the mass of fat and other tissues from the periphery of the liver. Be careful not to scratch the liver.

• (12)Grasp the hepatic portal area using the precision tweezers and remove the liver from the abdominal cavity.

• (13)Place the liver on the slide glass. Using the precision tweezers, remove any additional fat and extraneous tissue (e.g., abdominal lining), if needed, from the surface of the liver.

• (14)Measure the liver weight with 1.5 ml microtube as a tare using an electronic analytical balance. Record the value on the worksheet (read: 0,1 mg). Confirm the identification information on the microtube label.

• (15)Close the cap of the microtube containing the liver. Store it in a cooling rack (or ice rack).

• (16)Following the excision of one liver, clean the dissection instruments or replace them with clean ones.

• (17)Remove livers from all of the fish in the transport container as described above.

• (18)After the livers have been excised from all of the fish in the transport container (i.e., all males or females in a test chamber), place all liver specimens in a tube rack with a label for identification and store it in a freezer. When the livers are donated for pre-treatment shortly after the excision, the specimens are carried to the next workstation in a cooling rack (or ice rack).

Following liver excision, the fish carcass is available for measurement of secondary sex characteristics.

Specimen

Store the liver specimens taken from the test fish at ≤ – 70 °C if they are not used for the pre-treatment shortly after the excision.

Figure 1

A cut is made just anterior to pectoral fins with scissors.

Image omitted; see official source

Figure 2

The midline of abdomen is incised with scissors to a point approximately 2 mm cranial to the anus.

Image omitted; see official source

Figure 3

The abdominal walls are spread with forceps for exposure of the liver and other internal organs. (Alternatively, the abdominal walls may be pinned laterally).

Image omitted; see official source

Figure 4

The liver is bluntly dissected and excised using forceps.

Image omitted; see official source

Figure 5

The intestines are gently retracted using forceps.

Image omitted; see official source

Testis 6

Both ends of the intestines and any mesenteric attachments are severed using scissors.

Image omitted; see official source

Testis 7 (female)

The procedure is identical for the female.

Image omitted; see official source

Testis 8

The completed procedure.

Image omitted; see official source

Procedure 2 B: Japanese Medaka (Oryzias latipes), Liver Pre-treatment for Vitellogenin Analysis

Take the bottle of homogenate buffer from the ELISA kit and cool it with crushed ice (temperature of the solution: ≤ 4 °C). If homogenate buffer from EnBio ELISA system is used, thaw the solution at room temperature, and then cool the bottle with crushed ice.

Calculate the volume of homogenate buffer for the liver on the basis of its weight (add 50 μl of homogenate buffer per mg liver weight for homogenate). For example, if the weight of the liver is 4,5 mg, the volume of homogenate buffer for the liver is 225 μl. Prepare a list of the volume of homogenate buffer for all livers.

Preparation of the liver for pre-treatment

• (1)Take the 1,5 ml microtube containing the liver from the freezer just before the pre-treatment.

• (2)Pre-treatment of the liver from males should be performed before females to prevent vitellogenin contamination. In addition, the pre-treatment for test groups should be conducted in the following order: control, solvent control (where appropriate), lowest concentration, middle concentration, highest concentration and positive control.

• (3)The number of 1,5 ml microtubes containing liver samples taken from the freezer at a given time should not exceed the number that can be centrifuged at that time.

• (4)Arrange the 1,5 ml microtubes containing liver samples in the order of specimen number on the ice rack (no need to thaw the liver).

Operation of the pre-treatment

• (1)Check the list for the volume of the homogenate buffer to be used for a particular sample of liver and adjust the micropipette (volume range: 100-1 000 μl) to the appropriate volume. Attach a clean tip to the micropipette.

• (2)Take the homogenate buffer from the reagent bottle and add the buffer to the 1,5 ml microtube containing the liver.

• (3)Add the homogenate buffer to all of 1,5 ml microtubes containing the liver according to the procedure described above. There is no need to change the micropipette tip to a new one. However, if the tip is contaminated or suspected to be contaminated, the tip should be changed.

• (1)Attach a new pestle for homogenisation to the microtube homogeniser.

• (2)Insert the pestle into the 1,5 ml microtube. Hold the microtube homogeniser to press the liver between the surface of the pestle and the inner wall of the 1,5 ml microtube.

• (3)Operate the microtube homogeniser for 10 to 20 seconds. Cool the 1,5 ml microtube with crushed ice during the operation.

• (4)Lift up the pestle from the 1,5 ml microtube and leave it at rest for about 10 seconds. Then conduct a visual check of the state of the suspension.

• (5)If pieces of liver are observed in the suspension, repeat the operations (3) and (4) to prepare satisfactory liver homogenate.

• (6)Cool the suspended liver homogenate on the ice rack until centrifugation.

• (7)Change the pestle to the new one for each homogenate.

• (8)Homogenise all livers with homogenate buffer according to the procedure described above.

• (1)Confirm the temperature of the refrigerated centrifuge chamber at ≤ 5 °C.

• (2)Insert the 1,5 ml microtubes containing the suspended liver homogenate in refrigerated centrifuge (adjust the balance if necessary).

• (3)Centrifuge the suspended liver homogenate at 13 000 g for 10 min at ≤ 5 °C. However, if the supernatants are adequately separated, centrifugal force and time may be adjusted as needed.

• (4)Following centrifugation, check that the supernatants are adequately separated (surface: lipid, intermediate: supernatant, bottom layer: liver tissue). If the separation is not adequate, centrifuge the suspension again under the same conditions.

• (5)Remove all specimens from the refrigerated centrifuge and arrange them in the order of specimen number on the ice rack. Be careful not to resuspend each separated layer after the centrifugation.

• (1)Place four 0,5 ml microtubes for storage of the supernatant into the tube rack.

• (2)Collect 30 μl of each supernatant (separated as the intermediate layer) with the micropipette and dispense it to one 0,5 ml microtube. Be careful not to collect the lipid on the surface or the liver tissue in the bottom layer.

• (3)Collect the supernatant and dispense it to other two 0,5 ml microtubes in the same manner as described above.

• (4)Collect the rest of the supernatant with the micropipette (if feasible: ≥ 100 μl). Then dispense the supernatant to the remaining 0,5 ml microtube. Be careful not to collect the lipid on the surface or the liver tissue in the bottom layer.

• (5)Close the cap of the 0,5 ml microtube and write the volume of the supernatant on the label. Then immediately cool the microtubes on the ice rack.

• (6)Change the tip of the micropipette to the new one for each supernatant. If a large amount of lipid becomes attached to the tip, change it to the new one immediately to avoid contamination of the liver extract with fat.

• (7)Dispense all of the centrifuged supernatant to four 0,5 ml microtubes according to the procedure described above.

• (8)After dispensing the supernatant to the 0,5 ml microtubes, place all of them in the tube rack with the identification label, and then freeze them in the freezer immediately. If the VTG concentrations are measured immediately after the pre-treatment, keep one 0,5 ml microtube (containing 30 μl of supernatant) cool in the tube rack and transfer it to the workstation where the ELISA assay is conducted. In such case, place the remaining microtubes in the tube racks and freeze them in the freezer.

• (9)After the collection of the supernatant, discard the residue adequately.

Storage of the specimen

Store the 0,5 ml microtubes containing the supernatant of the liver homogenate at ≤ – 70 °C until they are used for the ELISA.

Procedure 3A: Zebrafish, Blood Collection from the Caudal Vein / Artery

Immediately following anaesthesia, the caudal peduncle is severed transversely, and the blood is removed from the caudal artery/vein with a heparinised microhematocrit capillary tube. Blood volumes range from 5 to 15 μl depending on fish size. An equal volume of aprotinin buffer (6 μgml in PBS) is added to the microcapillary tube, and plasma is separated from the blood via centrifugation (5 minutes at 600 g). Plasma is collected in the test tubes and stored at – 20 °C until analyzed for vitellogenin or other proteins of interest.

Procedure 3B: Zebrafish, Blood Collection by Cardiac Puncture

To avoid coagulation of blood and degradation of protein the samples are collected within Phosphate-buffered saline (PBS) buffer containing heparin (1 000 units/ml) and the protease inhibitor aprotinin (2 TIU/ml). As ingredients for the buffer, heparin ammonium salt and lyophilised aprotinin are recommended. For blood sampling, a syringe (1 ml) with a fixed thin needle (e.g. Braun Omnikan-F) is recommended. The syringe should be prefilled with buffer (approximately 100 μl) to completely elute the small blood volumes from each fish. The blood samples are taken by cardiac puncture. At first the fish should be anesthetized with MS-222 (100 mg/l). The proper plane of anaesthesia allows the user to distinguish the heartbeat of the zebrafish. While puncturing the heart, keep the syringe piston under weak tension. Collectable blood volumes range between 20 - 40 microliters. After cardiac puncture, the blood/buffer-mixture should be filled into the test tube. Plasma is separated from the blood via centrifugation (20 min; 5 000 g) and should be stored at – 80 °C until required for analysis.

Procedure 3C: SOP: Zebrafish, homogenisation of head & tail

• (1)The fish are anaesthetised and euthanised in accordance with the test description.

• (2)The head and tail are cut of the fish in accordance with Figure 1.

Important: All dissection instruments, and the cutting board should be rinsed and cleaned properly (e.g. with 96 % ethanol) between handling of each single fish to prevent ‘vitellogenin pollution’ from females or induced males to uninduced males.

Figure 1

Image omitted; see official source

Text of image

Cut behind

Cut behind dorsal fin

For Vtg analysis

For Vtg analysis

For gonad histology

• (3)The weight of the pooled head and tail from each fish is measured to the nearest mg.

• (4)After being weighed, the parts are placed in appropriate tubes (e.g. 1,5 ml eppendorf) and frozen at – 80 °C until homogenisation or directly homogenised on ice with two plastic pistils. (Other methods can be used if they are performed on ice and the result is a homogenous mass). Important: The tubes should be numbered properly so that the head and tail from the fish can be related to their respective body-section used for gonad histology.

• (5)When a homogenous mass is achieved, 4 x the tissue weight of ice-cold homogenisation buffer ([^E0200]) is added. Keep working with the pistils until the mixture is homogeneous. Important note: New pistils are used for each fish.

• (6)The samples are placed on ice until centrifugation at 4 °C at 50 000 × g for 30 min.

• (7)Use a pipette to dispense portions of 20 μl supernatant into at least two tubes by dipping the tip of the pipette below the fat layer on the surface and carefully sucking up the supernatant without fat- or pellet fractions.

• (8)The tubes are stored at – 80 °C until use.

Appendix 7

Vitellogenin fortification samples and inter-assay reference standard

On each day that vitellogenin assays are performed, a fortification sample made using an inter-assay reference standard will be analysed. The vitellogenin used to make the inter-assay reference standard will be from a batch different from the one used to prepare calibration standards for the assay being performed.

The fortification sample will be made by adding a known quantity of the inter-assay standard to a sample of control male plasma. The sample will be fortified to achieve a vitellogenin concentration between 10 and 100 times the expected vitellogenin concentration of control male fish. The sample of control male plasma that is fortified may be from an individual fish or may be a composite from several fish.

A subsample of the unfortified control male plasma will be analysed in at least two duplicate wells. The fortified sample also will be analysed in at least two duplicate wells. The mean quantity of vitellogenin in the two unfortified control male plasma samples will be added to the calculated quantity of vitellogenin added to fortification the samples to determine an expected concentration. The ratio of this expected concentration to the measured concentration will be reported along with the results from each set of assays performed on that day.

Appendix 8

Decision flowchart for the statistical analysis

Image omitted; see official source

Text of image

Dunnett test on nested ANOVA

No

Yes

Dunn or Mann-Whitney test on rep means

variance stabilising transform

=4 reps per conc

<=3 reps per conc

Dunn test

Yes

No

Normalising transform?

Tamhane-Dunnett test

=4 reps per conc

<=3 reps per conc

Nested ANOVA not normal

No

Yes

No

Yes

variance stabilising transformation

Dunnett test

Variances unequal

Variances equal

Not monotone

Stept-down Jonckheere or Williams' test

Step-down trendtest on replicate means

Monotone

Tamhane-Dunnett test on nested ANOVA

Dunn test on rep means

Dunn or Mann-Whitney test

Nested ANOVA normal

Rep means normally distributed

Rep means normally distributed

Step-down Jonckheere test

Rep means normal or not homogenous

Rep means normal & homogenous

Determine whether Dose-Response is monotone

C.38.   THE AMPHIBIAN METAMORPHOSIS ASSAY

INTRODUCTION

• 1.This test method is equivalent to OECD test guideline (TG) 231 (2009). The need to develop and validate an assay capable of detecting chemicals active in the thyroid system of vertebrate species originates from concerns that environmental levels of chemicals may cause adverse effects in both humans and wildlife. In 1998, the OECD initiated a high-priority activity to revise existing TGs and to develop new TGs for the screening and testing of potential endocrine disrupters. One element of the activity was to develop a TG for the screening of chemicals active on the thyroid system of vertebrate species. Both an enhancement of the Repeated dose 28-day oral toxicity study in rodents (Chapter B.7 of this Annex) and the Amphibian Metamorphosis Assay (AMA) were proposed. The enhanced test method B.7 underwent validation and a revised test method has been issued. The Amphibian Metamorphosis Assay (AMA) underwent an extensive validation programme which included intra- and inter-laboratory studies demonstrating the relevance and reliability of the assay (1, 2). Subsequently, the validation of the assay was subject to peer-review by a panel of independent experts (3). This test method is the outcome of the experience gained during the validation studies for the detection of thyroid active chemicals, and of work conducted elsewhere in OECD member countries.

PRINCIPLE OF THE TEST

• 2.The Amphibian Metamorphosis Assay (AMA) is a screening assay intended to empirically identify chemicals which may interfere with the normal function of the hypothalamic-pituitary-thyroid (HPT) axis. The AMA represents a generalised vertebrate model to the extent that it is based on the conserved structures and functions of the HPT axis. It is an important assay because amphibian metamorphosis provides a well-studied, thyroid-dependent process which responds to chemicals active within the HPT axis, and it is the only existing assay that detects thyroid activity in an animal undergoing morphological development.

• 3.The general experimental design entails exposing stage 51 Xenopus laevis tadpoles to a minimum of three different concentrations of a test chemical and a dilution water control for 21 days. There are four replicates of each test treatment. Larval density at test initiation is 20 tadpoles per test tank for all treatment groups. The observational endpoints are hind limb length, snout to vent length (SVL), developmental stage, wet weight, thyroid histology, and daily observations of mortality.

DESCRIPTION OF THE METHOD

Test Species

• 1. Xenopus laevis is routinely cultured in laboratories worldwide and is easily obtainable through commercial suppliers. Reproduction can be easily induced in this species throughout the year using human chorionic gonadotropin (hCG) injections and the resultant larvae can be routinely reared to selected developmental stages in large numbers to permit the use of stage-specific test protocols. It is preferred that larvae used in the assay are derived from in-house adults. As an alternative although this is not the preferred procedure, eggs or embryos may be shipped to the laboratory performing the test and allowed to acclimate; the shipping of larval stages for use in the test is unacceptable.

Equipment and Supplies

• 5.The following equipment and supplies are needed for the conduct of this assay:

• (a) Exposure system (see description below);

• (b) Glass or stainless steel aquaria (see description below);

• (c) Breeding tanks;

• (d) Temperature controlling apparatus (e.g., heaters or coolers (adjustable to 22° ± 1 °C));

• (e) Thermometer;

• (f) Binocular dissection microscope;

• (g) Digital camera with at least 4 megapixel resolution and micro function;

• (h) Image digitising software;

• (i) Petri dish (e.g. 100 × 15 mm) or transparent plastic chamber of comparable size;

• (j) Analytical balance capable of measuring to 3 decimal places (mg);

• (k) Dissolved oxygen meter;

• (l) pH meter;

• (m) Light intensity meter capable of measuring in lux units;

• (n) Miscellaneous laboratory glassware and tools;

• (o) Adjustable pipettes (10 to 5 000 μl) or assorted pipettes of equivalent sizes;

• (p) Test chemical in sufficient quantities to conduct the study, preferably of one lot;

• (q) Analytical instrumentation appropriate for the chemical on test or contracted analytical services.

Chemical Testability

• 6.The AMA is based upon an aqueous exposure protocol whereby test chemical is introduced into the test chambers via a flow-through system. Flow-through methods however, introduce constraints on the types of chemicals that can be tested, as determined by the physicochemical properties of the chemical. Therefore, prior to using this protocol, baseline information about the chemical should be obtained that is relevant to determining the testability, and the OECD Guidance Document on Aquatic Toxicity Testing of Difficult Substances and Mixtures (4) should be consulted. Characteristics which indicate that the chemical may be difficult to test in aquatic systems include: high octanol water partitioning coefficients (log Kow), high volatility, susceptibility to hydrolysis, and susceptibility to photolysis under ambient laboratory lighting conditions. Other factors may also be relevant to determining testability and should be determined on a case by case basis. If a successful test is not possible for the chemical using a flow-through test system, a static renewal system may be employed. If neither system is capable of accommodating the test chemical, then the default is to not test it using this protocol.

Exposure System

• 7.A flow-through diluter system is preferred, when possible, over a static renewal system. If physical and/or chemical properties of any of the test chemicals are not amenable to a flow-through diluter system, then an alternative exposure system (e.g., static-renewal) can be employed. The system components should have water-contact components of glass, stainless steel, and/or Polytetrafluoroethylene. However, suitable plastics can be utilised if they do not compromise the study. Exposure tanks should be glass or stainless steel aquaria, equipped with standpipes that result in an approximate tank volume between 4,0 and 10,0 l and minimum water depth of 10 to 15 cm. The system should be capable of supporting all exposure concentrations and a control, with four replicates per treatment. The flow rate to each tank should be constant in consideration of both the maintenance of biological conditions and chemical exposure (e.g. 25 ml/min). The treatment tanks should be randomly assigned to a position in the exposure system in order to reduce potential positional effects, including slight variations in temperature, light intensity, etc. Fluorescent lighting should be used to provide a photoperiod of 12 hr light: 12 hr dark at an intensity that ranges from 600 to 2 000 lux (lumen/m²) at the water surface. Water temperature should be maintained at 22° ± 1 °C, pH maintained between 6,5 to 8,5, and the dissolved oxygen (DO) concentration > 3,5 mg/l (> 40 % of the air saturation) in each test tank. As a minimum water temperature, pH and dissolved oxygen should be measured weekly; temperature should preferably be measured continuously in at least one test vessel. Appendix 1 outlines the experimental conditions under which the protocol should be executed. For further information on setting up flow-through exposure systems and/or static renewal systems, please refer to the ASTM Standard Guide for Conducting Acute Toxicity Tests on Test Materials with Fishes, Macroinvertebrates, and Amphibians (5) and general aquatic toxicology tests.

Water quality

• 8.Any water that is locally available (e.g. springwater or charcoal-filtered tap water) and permits normal growth and development of X. laevis tadpoles could be used. Because local water quality can differ substantially from one area to another, analysis of water quality should be undertaken, particularly, if historical data on the utility of the water for raising Xenopus is not available. Special attention should be given that the water is free of copper, chlorine and chloramines, all of which are toxic to frogs and tadpoles. It is further recommended to analyse the water concerning background levels of fluoride, perchlorate and chlorate (by-products of drinking water disinfection) as all of these anions are substrates of the iodine transporter of the thyroid gland and elevated levels of each of these anions may confound the study outcome. Analysis should be performed before testing begins and the testing water should normally be free from these anions.

Iodide Concentration in Test Water

• 9.In order for the thyroid gland to synthesise TH, sufficient iodide needs to be available to the larvae through a combination of aqueous and dietary sources. Currently, there are no empirically derived guidelines for minimal iodide concentrations. However, iodide availability may affect the responsiveness of the thyroid system to thyroid active agents and is known to modulate the basal activity of the thyroid gland, an aspect that deserves attention when interpreting the results from thyroid histopathology. Therefore, measured aqueous iodide concentrations from the test water should be reported. Based on the available data from the validation studies, the protocol has been demonstrated to work well when test water iodide (I^-) concentrations ranged between 0,5 and 10 μg/l. Ideally, the minimum iodide concentration in the test water should be 0,5 μg/l. If the test water is reconstituted from deionised water, iodine should be added at a minimum concentration of 0,5 μg/l. Any additional supplementation of the test water with iodine or other salts should be noted in the report.

Holding of animals

Adult Care and Breeding

• 10.Adult care and breeding is conducted in accordance with standard guidelines and the reader is directed to the standard guide for performing the Frog Embryo Teratogenesis Assay (FETAX) (6) for more detailed information. Such standard guidelines provide an example of appropriate care and breeding methods, but strict adherence is not required. To induce breeding, pairs (3-5) of adult females and males are injected with human chorionic gonadotropin (hCG). Female and male specimens are injected with approximately 800 IU-1 000 IU and 600 IU-800 IU, respectively, of hCG dissolved in 0,6-0,9 % saline solution. Breeding pairs are held in large tanks, undisturbed and under static conditions in order to promote amplexus. The bottom of each breeding tank should have a false bottom of stainless steel or plastic mesh which permits the egg masses to fall to the bottom of the tank. Frogs injected in the late afternoon will usually deposit most of their eggs by mid morning of the next day. After a sufficient quantity of eggs are released and fertilised, adults should be removed from the breeding tanks.

Larval Care and Selection

• 11.After the adults are removed from the breeding tanks, the eggs are collected and evaluated for viability using a representative sub-set of the embryos from all breeding tanks. The best individual spawn(s) (2-3 recommended to evaluate the quality of the spawns) should be retained based upon embryo viability and the presence of an adequate number (minimum of 1 500 ) of embryos. All the organisms used in a study should originate from a single spawning event (i.e., the spawns should not be co-mixed). The embryos are transferred into a large flat pan or dish and all obvious dead or abnormal eggs (see definition in (5)) are removed using a pipette or eyedropper. The sound embryos from each of the three spawns are transferred into three separate hatching tanks. Four days after being placed in the hatching tanks, the best spawn, based on viability and hatching success, is selected and the larvae are transferred into an appropriate number of rearing tanks at 22° ± 1 °C. In addition, some additional larvae are moved into extra tanks for use as replacements in the event that mortalities occur in the rearing tanks during the first week. This procedure maintains consistent organism density and thereby reduces developmental divergence within the cohort of a single spawn. All rearing tanks should be siphoned clean daily. As a precaution, vinyl or nitrile gloves are preferred to latex gloves. Mortalities should be removed daily and replacement larvae should be added back to maintain the organism density during the first week. Feeding should occur at least twice per day.