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)

| --- | --- | --- | --- | | 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 21 | See paragraph 21 | See paragraph 21 | | 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) | 6 (3 males and 3 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 | 4 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) | 12 adult females and 12 males (3 females and 3 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 | 7-14 days recommended | 7-14 days recommended | 7-14 days recommended | | 21. Chemical exposure duration | 21-d | 21-d | 21-d | | 22. Biological endpoints | — survival — behaviour — fecundity — 2y sex characteristics — VTG — optionally gonadal histopathology | — survival — behaviour — fecundity — 2y sex characteristics — VTG — optionally gonadal histopathology | — survival — behaviour — fecundity — VTG — optionally gonadal histopathology | | 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 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 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 2mm). 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.

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 specialised 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 17α-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 characterisation 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).

• — 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.

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

- — 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. 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 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 17α-methyltestosterone on the dorsal pad and tubercles of fathead minnows (Pimephales promelas). Can J Zool 52:1031-1038.

Image omitted; see official source

Text of image

Tubercle Template:

ID

Numerical Rating

Date

1-present

Total Score

2-enlarged

3-pronounced

A

X1

X1

X1

X1

B

X1

X1

X1

X1

C

X1

X1

X1

X1

X1

X1

X1

X1

X1

X1

D

X1

X1

X1

X1

X1

X1

X1

X1

X1

X1

E

X1

X1

F

X1

X1

X1

X1

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 ([^E0255]), 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 recognised 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 haematocrit 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 VTG can be made. Depending on the size of the fathead minnow (which is sex-dependent), collectable plasma volumes generally range from 5 to 60 microliters 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 heparinised 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 analysed (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 anesthetised, 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 gonad histology and 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).

Arrow shows liver

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

Figure 6

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

Image omitted; see official source

Figure 7 (female)

The procedure is identical for the female

Image omitted; see official source

Figure 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 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) Addition of the homogenate buffer

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.

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

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.

(2) Homogenisation of the liver

• — Attach a new pestle for homogenisation to the microtube homogeniser.

• — 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.

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

• — 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.

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

• — Cool the suspended liver homogenate on the ice rack until centrifugation.

• — Change the pestle to the new one for each homogenate.

• — Homogenise all livers with homogenate buffer according to the procedure described above.

(3) Centrifugation of the suspended liver homogenate

• — Confirm the temperature of the refrigerated centrifuge chamber at ≤ 5 °C.

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

• — 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.

• — 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.

• — 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.

(4) Collection of the supernatant

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

• — 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.

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

• — 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.

• — 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.

• — 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.

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

• — 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.

• — 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 microliters depending on fish size. An equal volume of aprotinin buffer (6 micrograms/ml 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 analysed for VTG 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 (1ml) with a fixed thin needle (e.g. Braun Omnikan-F) is recommended. The syringe should be prefilled with buffer (approximately 100 microliter) 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

For gonad histology

Cut behind

Cut behind dorsal fin

For Vtg analysis

For Vtg analysis

• 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 ([^E0256]) 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 VTG assays are performed, a fortification sample made using an inter-assay reference standard will be analysed. The VTG 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 VTG 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 VTG 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

Rep means normally distributed

Stept-down Jonckheere or Williams' test

Normalising transform?

variance stabilising transform?

Tamhane-Dunnett test on nested ANOVA

Dunnett test on nested ANOVA

No

Yes

Yes

No

=4 reps per conc

<=3 reps per conc

Dunn or Mann-Whitney test on rep means

Dunn test on rep means

Dunn or Mann-Whitney test

Dunn test

<=3 reps per conc

=4 reps per conc

Tamhane-Dunnett test

Normalising transform?

Nested ANOVA normal

Nested ANOVA not normal

No

Yes

Yes

No

variance stabilising transformation

Variances unequal

Dunnett test

Variances equal

Step-down Jonckheere test

Rep means normal or not homogenous

Rep means normal & homogenous

Rep means normally distributed

Step-down trendtest on replicate means

Not monotone

Monotone

Determine whether Dose-Response is monotone

C.49.   FISH EMBRYO ACUTE TOXICITY (FET) TEST

INTRODUCTION

• 1.This test method (TM) is equivalent to OECD test guideline (TG) 236 (2013). It describes a Fish Embryo Acute Toxicity (FET) test with the zebrafish (Danio rerio). This test is designed to determine acute toxicity of chemicals on embryonic stages of fish. The FET-test is based on studies and validation activities performed on zebrafish (1)(2)(3)(4)(5)(6)(7)(8)(9)(10)(11)(12)(13)(14). The FET-test has been successfully applied to a wide range of chemicals exhibiting diverse modes of action, solubilities, volatilities, and hydrophobicities (reviewed in 15 and 16).

- 2.Definitions used in this test method are given in Appendix 1.

PRINCIPLE OF THE TEST

- 3.Newly fertilised zebrafish eggs are exposed to the test chemical for a period of 96 hrs. Every 24 hrs, up to four apical observations are recorded as indicators of lethality (6): (i) coagulation of fertilised eggs, (ii) lack of somite formation, (iii) lack of detachment of the tail-bud from the yolk sac, and (iv) lack of heartbeat. At the end of the exposure period, acute toxicity is determined based on a positive outcome in any of the four apical observations recorded, and the LC50 is calculated.

INITIAL CONSIDERATIONS

• 4.Useful information about substance-specific properties include the structural formula, molecular weight, purity, stability in water and light, pKa and Kow, water solubility and vapour pressure as well as results of a test for ready biodegradability (TM C.4 (17) or TM C.29 (18)). Solubility and vapour pressure can be used to calculate Henry's law constant, which will indicate whether losses due to evaporation of the test chemical may occur. A reliable analytical method for the quantification of the substance in the test solutions with known and reported accuracy and limit of detection should be available.

• 5.If the test method is used for the testing of a mixture, its composition should, as far as possible, be characterised, e.g. by the chemical identity of its constituents, their quantitative occurrence and their substance-specific properties (see paragraph 4). Before use of the test method for regulatory testing of a mixture, it should be considered whether it will provide acceptable results for the intended regulatory purpose.

• 6.Concerning substances that may be activated via metabolism, there is evidence that zebrafish embryos do have biotransformation capacities (19)(20)(21)(22). However, the metabolic capacity of embryonic fish is not always similar to that of juvenile or adult fish. For instance, the protoxicant allyl alcohol (9) has been missed in the FET. Therefore, if there are any indications that metabolites or other transformation products of relevance may be more toxic than the parent compound, it is also recommended to perform the test with these metabolites/transformation products and to also use these results when concluding on the toxicity of the test chemical, or alternatively perform another test which takes metabolism into further account.

- 7.For substances with a molecular weight ≥ 3kDa, a very bulky molecular structure, and substances causing delayed hatch which might preclude or reduce the post-hatch exposure, embryos are not expected to be sensitive because of limited bioavailability of the substance, and other toxicity tests might be more appropriate.

VALIDITY OF THE TEST

• 8.For the test results to be valid, the following criteria apply:

• (a) The overall fertilisation rate of all eggs collected should be ≥ 70 % in the batch tested.

• (b) The water temperature should be maintained at 26 ± 1 °C in test chambers at any time during the test.

• (c) Overall survival of embryos in the negative (dilution-water) control, and, where relevant, in the solvent control should be ≥ 90 % until the end of the 96 hrs exposure.

• (d) Exposure to the positive control (e.g. 4,0 mg/l 3,4-dichloroaniline for zebrafish) should result in a minimum mortality of 30 % at the end of the 96 hrs exposure.

• (e) Hatching rate in the negative control (and solvent control if appropriate) should be ≥ 80 % at the end of 96 hrs exposure.

- (f) At the end of the 96 hrs exposure, the dissolved oxygen concentration in the negative control and highest test concentration should be ≥ 80 % of saturation.

DESCRIPTION OF THE METHOD

- 9.An overview of recommended maintenance and test conditions is available in Appendix 2.

Apparatus

• 10.The following equipment is needed:

• (a) Fish tanks made of chemically inert material (e.g. glass) and of a suitable capacity in relation to the recommended loading (see ‘Maintenance of brood fish’, paragraph 14);

• (b) Inverted microscope and/or binocular with a capacity of at least 80-fold magnification. If the room used for recording observations cannot be adjusted to 26 ± 1 °C, a temperature-controlled cross movement stage or other methods to maintain temperature are necessary;

• (c) Test chambers; e.g., standard 24-well plates with a depth of approx. 20 mm. (see ‘Test chambers’, paragraph 11);

• (d) e.g., self-adhesive foil to cover the 24-well plates;

• (e) Incubator or air-conditioned room with controlled temperature, allowing to maintain 26 ± 1 °C in wells (or test chambers);

• (f) pH-meter;

• (g) Oxygen meter;

• (h) Equipment for determination of hardness of water and conductivity;

• (i) Spawn trap: instrument trays of glass, stainless steel or other inert materials; wire mesh (grid size 2 ± 0,5 mm) of stainless steel or other inert material to protect the eggs once laid; spawning substrate (e.g. plant imitates of inert material) (TM C.48, Appendix 4a (23));

• (j) Pipettes with widened openings to collect eggs;

• (k) Glass vessels to prepare different test concentrations and dilution water (beakers, graduated flasks, graduated cylinders and graduated pipettes) or to collect zebrafish eggs (e.g. beakers, crystallisation dishes);

- (l) If alternative exposure systems, such as flow-through (24) or passive dosing (25) are used for the conduct of the test, appropriate facilities and equipment are needed.

Test chambers

- 11.Glass or polystyrene test chambers should be used (e.g. 24-well plates with a 2,5-5 ml filling capacity per well). In case adsorption to polystyrene is suspected (e.g., for non-polar, planar substances with high KOW), inert materials (glass) should be used to reduce losses due to adsorption (26). Test chambers should be randomly positioned in the incubator.

Water and test conditions

- 12.Dilution of the maintenance water is recommended to achieve hardness levels typical of a wide variety of surface waters. Dilution water should be prepared from reconstituted water (27). The resulting degree of hardness should be equivalent to 100-300 mg/l CaCO3 in order to prevent excessive precipitation of calcium carbonate. Other well-characterised surface or well water may be used. The reconstituted water may be adapted to maintenance water of low hardness by dilution with deionised water up to a ratio of 1:5 to a minimum hardness of 30-35 mg/l CaCO3. The water is aerated to oxygen saturation prior to addition of the test chemical. Temperature should be kept at 26 ± 1 °C, in the wells, throughout the test. The pH should be in a range between pH 6,5 and 8,5, and not vary within this range by more than 1,5 units during the course of the test. If the pH is not expected to remain in this range, then pH adjustment should be done prior to initiating the test. The pH adjustment should be made in such a way that the stock solution concentration is not changed to any significant extent and that no chemical reaction or precipitation of the test chemical is caused. Use of hydrogen chloride (HCl) and sodium hydroxide (NaOH) to correct pH in the solutions containing the test chemical is recommended.

Test solutions

- 13.Test solutions of the selected concentrations can be prepared, e.g. by dilution of a stock solution. The stock solutions should preferably be prepared by simply mixing or agitating the test chemical in the dilution water by mechanical means (e.g. stirring and/or ultra-sonification). If the test chemical is difficult to dissolve in water, procedures described in the OECD Guidance Document No. 23 for handling difficult substances and mixtures should be followed (28). The use of solvents should be avoided, but may be required in some cases in order to produce a suitably concentrated stock solution. Where a solvent is used to assist in stock solution preparation, its final concentration should not exceed 100 μl/l and should be the same in all test vessels. When a solvent is used, an additional solvent control is required.

Maintenance of brood fish

- 14.A breeding stock of unexposed, wild-type zebrafish with well-documented fertilisation rate of eggs is used for egg production. Fish should be free of macroscopically discernible symptoms of infection and disease and should not have undergone any pharmaceutical (acute or prophylactic) treatment for 2 months before spawning. Breeding fish are maintained in aquaria with a recommended loading capacity of 1 l water per fish and a fixed 12 - 16 hour photoperiod (29)(30)(31)(32)(33). Optimal filtering rates should be adjusted; excess filtering rates causing heavy perturbation of the water should be avoided. For feeding conditions, see Appendix 2. Surplus feeding should be avoided, and water quality and cleanness of the aquaria should be monitored regularly and be reset to the initial state, if necessary.

Proficiency Testing

- 15.As a reference chemical, 3,4-dichloroaniline (used in the validation studies (1)(2)), should be tested in a full concentration-response range to check the sensitivity of the fish strain used, preferably twice a year. For any laboratory initially establishing this assay, the reference chemical should be used. A laboratory can use this chemical to demonstrate their technical competence in performing the assay prior to submitting data for regulatory purposes.

Egg production

• 16.Zebrafish eggs may be produced via spawning groups (in individual spawning tanks) or via mass spawning (in the maintenance tanks). In the case of spawning groups, males and females (e.g., at a ratio of 2:1) in a breeding group are placed in spawning tanks a few hours before the onset of darkness on the day prior to the test. Since spawning groups of zebrafish may occasionally fail to spawn, the parallel use of at least three spawning tanks is recommended. To avoid genetic bias, eggs are collected from a minimum of three breeding groups, mixed and randomly selected.

- 17.For the collection of eggs, spawn traps are placed into the spawning tanks or maintenance tanks before the onset of darkness on the day prior to the test or before the onset of light on the day of the test. To prevent predation of eggs by adult zebrafish, the spawn traps are covered with inert wire mesh of appropriate mesh size (approx. 2 ± 0,5 mm). If considered necessary, artificial plants made of inert material (e.g., plastic or glass) can be fixed to the mesh as spawning stimulus (3)(4)(5)(23)(35). Weathered plastic materials which do not leach (e.g., phthalates) should be used. Mating, spawning and fertilisation take place within 30 min after the onset of light and the spawn traps with the collected eggs can be carefully removed. Rinsing eggs with reconstituted water after collection from spawning traps is recommended.

Egg differentiation

- 18.At 26 °C, fertilised eggs undergo the first cleavage after about 15 min and the consecutive synchronous cleavages form 4, 8, 16 and 32 cell blastomers (see Appendix 3)(35). At these stages, fertilised eggs can be clearly identified by the development of a blastula.

PROCEDURE

Conditions of exposure

- 19.Twenty embryos per concentration (one embryo per well) are exposed to the test chemical. Exposure should be such that ± 20 % of the nominal chemical concentration are maintained throughout the test. If this is not possible in a static system, a manageable semi-static renewal interval should be applied (e.g. renewal every 24 hrs). In these cases exposure concentrations need to be verified as a minimum in the highest and lowest test concentrations at the beginning and the end of each exposure interval (see paragraph 36). If an exposure concentration of ± 20 % of the nominal concentrations cannot be maintained, all concentrations need to be measured at the beginning and the end of each exposure interval (see paragraph 36). Upon renewal, care should be taken that embryos remain covered by a small amount of old test solutions to avoid drying. The test design can be adapted to meet the testing requirements of specific substances (e.g,. flow-through (24) or passive dosing systems (25) for easily degradable or highly adsorptive substances (29), or others for volatile substances (36)(37)). In any case, care should be taken to minimise any stress to the embryos. Test chambers should be conditioned at least for 24 hrs with the test solutions prior to test initiation. Test conditions are summarised in Appendix 2.

Test concentrations

• 20.Normally, five concentrations of the test chemical spaced by a constant factor not exceeding 2,2 are required to meet statistical requirements. Justification should be provided, if fewer than five concentrations are used. The highest concentration tested should preferably result in 100 % lethality, and the lowest concentration tested should preferably give no observable effect, as defined in paragraph 28. A range-finding test before the definitive test allows selection of the appropriate concentration range. The range-finding is typically performed using ten embryos per concentration. The following instructions refer to performing the test in 24-well plates. If different test chambers (e.g. small Petri dishes) are used or more concentrations are tested, instructions have to be adjusted accordingly.

- 21.Details and visual instructions for allocation of concentrations across 24-well plates are available in paragraph 27 and Appendix 4, Figure 1.

Controls

• 22.Dilution water controls are required both as negative control and as internal plate controls. If more than 1 dead embryo is observed in the internal plate control, the plate is rejected, thus reducing the number of concentrations used to derive the LC50. If an entire plate is rejected the ability to evaluate and discern observed effects may become more difficult, especially if the rejected plate is the solvent control plate or a plate in which treated embryos are also affected. In the first case the test must be repeated. In the second one the loss of an entire treatment group(s) due to internal control mortality may limit the ability to evaluate effects and determine LC50 values.

• 23.A positive control at a fixed concentration of 4 mg/l 3,4-dichloroaniline is performed with each egg batch used for testing.

- 24.In case a solvent is used, an additional group of 20 embryos is exposed to the solvent on a separate 24-well plate, thus serving as a solvent control. To consider the test acceptable, the solvent should be demonstrated to have no significant effects on time to hatch, survival, nor produce any other adverse effects on the embryos (cf. paragraph 8c).

Start of exposure and duration of test

• 25.The test is initiated as soon as possible after fertilisation of the eggs and terminated after 96 hrs of exposure. The embryos should be immersed in the test solutions before cleavage of the blastodisc commences, or, at latest, by the 16 cell-stage. To start exposure with minimum delay, at least twice the number of eggs needed per treatment group are randomly selected and transferred into the respective concentrations and controls (e.g. in 100 ml crystallisation dishes; eggs should be fully covered) not later than 90 minutes post fertilisation.

- 26.Viable fertilised eggs should be separated from unfertilised eggs and be transferred to 24-well plates pre-conditioned for 24 hrs and refilled with 2 ml/well freshly prepared test solutions within 180 minutes post fertilisation. By means of stereomicroscopy (preferably ≥30-fold magnification), fertilised eggs undergoing cleavage and showing no obvious irregularities during cleavage (e.g. asymmetry, vesicle formation) or injuries of the chorion are selected. For egg collection and separation, see Appendix 3, Fig. 1 and 3 and Appendix 4, Fig. 2.

Distribution of eggs over the 24-well plates

• 27.Eggs are distributed to well plates in the following numbers (see also Appendix 4, Fig. 1)

• — 20 eggs on one plate for each test concentration;

• — 20 eggs as solvent control on one plate (if necessary);

• — 20 eggs as positive control on one plate;

• — 4 eggs in dilution water as internal plate control on each of the above plates;

- — 24 eggs in dilution water as negative control on one plate.

Observations

- 28.Apical observations performed on each tested embryo include: coagulation of embryos, lack of somite formation, non-detachment of the tail, and lack of heartbeat (Table 1). These observations are used for the determination of lethality: Any positive outcome in one of these observations means that the zebrafish embryo is dead. Additionally, hatching is recorded in treatment and control groups on a daily basis starting from 48 hrs. Observations are recorded every 24 hrs, until the end of the test.

Table 1

Apical observations of acute toxicity in zebrafish embryos 24-96 hrs post fertilisation

	Exposure times	Exposure times	Exposure times	Exposure times
	24 hrs	48 hrs	72 hrs	96 hrs
Coagulated embryos	+	+	+	+
Lack of somite formation	+	+	+	+
Non-detachment of the tail	+	+	+	+
Lack of heartbeat		+	+	+

• 1. Coagulation of the embryo: Coagulated embryos are milky white and appear dark under the microscope (see Appendix 5, Fig. 1). The number of coagulated embryos is determined after 24, 48, 72 and 96 hrs.

• 1. Lack of somite formation: At 26 ± 1 °C, about 20 somites have formed after 24 hrs (see Appendix 5, Figure 2) in a normally developing zebrafish embryo. A normally developed embryo shows spontaneous movements (side-to-side contractions). Spontaneous movements indicate the formation of somites. The absence of somites is recorded after 24, 48, 72 and 96 hrs. Non-formation of somites after 24 hrs might be due to a general retardation of development. At latest after 48 hrs, the formation of somites should be developed. If not, the embryos are considered dead.

• 1. Non-detachment of the tail: In a normally developing zebrafish embryo, detachment of the tail (seeAppendix 5, Figure 3) from the yolk is observed following posterior elongation of the embryonic body. Absence of tail detachment is recorded after 24, 48, 72 and 96 hrs.

• 1. Lack of heartbeat: In a normally developing zebrafish embryo at 26 ± 1 °C, the heartbeat is visible after 48 hrs (see Appendix 5, Figure 4). Particular care should be taken when recording this endpoint, since irregular (erratic) heartbeat should not be recorded as lethal. Moreover, visible heartbeat without circulation in aorta abdominalis is considered non-lethal. To record this endpoint, embryos showing no heartbeat should be observed under a minimum magnification of 80x for at least one minute. Absence of heartbeat is recorded after 48, 72 and 96 hrs.

• 33.Hatching rates of all treatment and control groups should be recorded from 48 hrs onwards and reported. Although hatching is not an endpoint used for the calculation of the LC50, hatching ensures exposure of the embryo without a potential barrier function of the chorion, and as such may help data interpretation.

- 34.Detailed descriptions of the normal (35) and examples of abnormal development of zebrafish embryos are illustrated in Appendixes 3 and 5.

Analytical measurements

• 35.At the beginning and at the end of the test, pH, total hardness and conductivity in the control(s) and in the highest test chemical concentration are measured. In semi-static renewal systems the pH should be measured prior to and after water renewal. The dissolved oxygen concentration is measured at the end of the test in the negative controls and highest test concentration with viable embryos, where it should be in compliance with the test validity criteria (see paragraph 8f). If there is concern that the temperature varies across the 24-well plates, temperature is measured in three randomly selected vessels. Temperature should be recorded preferably continuously during the test or, as a minimum, daily.

- 36.In a static system, the concentration of the test chemical should be measured, as a minimum, in the highest and lowest test concentrations, but preferably in all treatments, at the beginning and end of the test. In semi-static (renewal) tests where the concentration of the test chemical is expected to remain within ± 20 % of the nominal values, it is recommended that, as a minimum, the highest and lowest test concentrations be analysed when freshly prepared and immediately prior to renewal. For tests where the concentration of the test chemical is not expected to remain within ± 20 % of nominal, all test concentrations must be analysed when freshly prepared and immediately prior to renewal. In case of insufficient volume for analysis, merging of test solutions, or use of surrogate chambers being of the same material and having the same volume to surface area ratios as 24-well plates, may be useful. It is strongly recommended that results be based on measured concentrations. When the concentrations do not remain within 80-120 % of the nominal concentration, the effect concentrations should be expressed relative to the geometric mean of the measured concentrations; see Chapter 5 in the OECD Guidance Document on Aquatic Toxicity Testing of Difficult Substances and Mixtures for more details (28).

LIMIT TEST

- 37.Using the procedures described in this test method, a limit test may be performed at 100 mg/l of test chemical or at its limit of solubility in the test medium (whichever is the lower) in order to demonstrate that the LC50 is greater than this concentration. The limit test should be performed using 20 embryos in the treatment, the positive control and –if necessary- in the solvent control and 24 embryos in the negative control. If the percentage of lethality at the concentration tested exceeds the mortality in the negative control (or solvent control) by 10 %, a full study should be conducted. Any observed effects should be recorded. If mortality exceeds 10 % in the negative control (or solvent control), the test becomes invalid and should be repeated.

DATA AND REPORTING

Treatment of results

- 38.In this test, the individual wells are considered independent replicates for statistical analysis. The percentages of embryos for which at least one of the apical observations is positive at 48 and/or 96 hrs are plotted against test concentrations. For calculation of the slopes of the curve, LC50 values and the confidence limits (95 %), appropriate statistical methods should be applied (38) and the OECD Guidance Document on Current Approaches in the Statistical Analysis of Ecotoxicity Data should be consulted (39).

Test report

• 39.The test report should include the following information:

Test chemical:

Mono-constituent substance

• — physical appearance, water solubility, and additional relevant physicochemical properties;

• — chemical identification, such as IUPAC or CAS name, CAS number, SMILES or InChI code, structural formula, purity, chemical identity of impurities as appropriate and practically feasible, etc. (including the organic carbon content, if appropriate).

Multi-constituent substance, UVCBs and mixtures:

• — characterised as far as possible by chemical identity (see above), quantitative occurrence and relevant physicochemical properties of the constituents.

Test organisms:

• — scientific name, strain, source and method of collection of the fertilised eggs and subsequent handling.

Test conditions:

• — test procedure used (e.g., semi-static renewal);

• — photoperiod;

• — test design (e.g., number of test chambers, types of controls);

• — water quality characteristics in fish maintenance (e.g. pH, hardness, temperature, conductivity, dissolved oxygen);

• — dissolved oxygen concentration, pH, total hardness, temperature and conductivity of the test solutions at the start and after 96 hrs;

• — method of preparation of stock solutions and test solutions as well as frequency of renewal;

• — justification for use of solvent and justification for choice of solvent, if other than water;

• — the nominal test concentrations and the result of all analyses to determine the concentration of the test chemical in the test vessels; the recovery efficiency of the method and the limit of quantification (LoQ) should also be reported;

• — evidence that controls met the overall survival validity criteria;

• — fertilisation rate of the eggs;

• — hatching rate in treatment and control groups.

Results:

• — maximum concentration causing no mortality within the duration of the test;

• — minimum concentration causing 100 % mortality within the duration of the test;

• — cumulative mortality for each concentration at the recommended observation times;

• — the LC50 values at 96 hrs (and optionally at 48 hrs) for mortality with 95 % confidence limits, if possible;

• — graph of the concentration-mortality curve at the end of the test;

• — mortality in the controls (negative controls, internal plate controls, as well as positive control and any solvent control used);

• — data on the outcome of each of the four apical observations;

• — incidence and description of morphological and physiological abnormalities, if any (see examples provided in Appendix 5, Figure 2);

• — incidents in the course of the test which might have influenced the results;

• — statistical analysis and treatment of data (probit analysis, logistic regression model and geometric mean for LC50);

• — slope and confidence limits of the regression of the (transformed) concentration-response curve.

Any deviation from the test method and relevant explanations.

Discussion and interpretation of results.

LITERATURE

• (1) OECD (2011) Validation Report (Phase 1) for the Zebrafish Embryo Toxicity Test: Part I and Part II. Series on Testing and Assessment No. 157, OECD, Paris.

• (2) OECD (2012) Validation Report (Phase 2) for the Zebrafish Embryo Toxicity Test: Part I and Part II (Annexes). Series on Testing and Assessment No. 179, OECD, Paris.

• (3) Braunbeck, T., Böttcher, M., Hollert, H., Kosmehl, T., Lammer, E., Leist, E., Rudolf, M. and Seitz, N. (2005) Towards an alternative for the acute fish LC50 test in chemical assessment: The fish embryo toxicity test goes multi-species-an update. ALTEX 22: 87-102.

• (4) ISO (2007) International Standard Water quality — Determination of the acute toxicity of waste water to zebrafish eggs (Danio rerio). ISO 15088:2007(E) International Organization for Standardization.

• (5) Nagel, R. (2002) DarT: The embryo test with the zebrafish (Danio rerio) — a general model in ecotoxicology and toxicology. ALTEX 19: 38-48.

• (6) Schulte, C. and Nagel, R. (1994) Testing acute toxicity in embryo of zebrafish, Brachydanio rerio as alternative to the acute fish test — preliminary results. ATLA 22, 12-19.

• (7) Bachmann, J. (2002) Development and validation of a teratogenicity screening test with embryos of the zebrafish (Danio rerio). PhD-thesis, Technical University of Dresden, Germany.

• (8) Lange, M., Gebauer, W., Markl, J. and Nagel, R. (1995) Comparison of testing acute toxicity on embryo of zebrafish (Brachydanio rerio), and RTG-2 cytotoxicity as possible alternatives to the acute fish test. Chemosphere 30/11: 2087-2102.

• (9) Knöbel, M., Busser, F.J.M., Rico-Rico, A., Kramer, N.I., Hermens, J.L.M., Hafner, C., Tanneberger, K., Schirmer, K., Scholz, S. (2012). Predicting adult fish acute lethality with the zebrafish embryo: relevance of test duration, endpoints, compound properties, and exposure concentration analysis. Environ. Sci. Technol. 46, 9690-9700.

• (10) Kammann, U., Vobach, M. and Wosniok, W. (2006) Toxic effects of brominated indoles and phenols on zebrafish embryos. Arch. Environ. Contam. Toxicol., 51: 97-102.

• (11) Groth, G., Kronauer, K. and Freundt, K.J. (1994) Effects of N,N-diemethylformamide and its degradation products in zebrafish embryos. Toxicol. In Vitro 8: 401-406.

• (12) Groth, G., Schreeb, K., Herdt, V. and Freundt, K.J. (1993) Toxicity studies in fertilized zebrafish fish eggs treated with N-methylamine, N,N-dimethylamine, 2-aminoethanol, isopropylamine, aniline, N-methylaniline, N,N-dimethylaniline, quinone, chloroacetaldehyde, or cyclohexanol. Bull. Environ. Contam. Toxicol. 50: 878-882.

• (13) Nguyen, L.T. and Janssen, C.R. (2001) Comparative sensitivity of embryo-larval toxicity assays with African catfish (Clarias gariepinus) and zebra fish (Danio rerio). Environ. Toxicol. 16: 566-571.

• (14) Cheng, S.H., Wai, A.W.K., So, C.H. and Wu, R.S.S. (2000) Cellular and molecular basis of cadmium-induced deformities in zebrafish embryos. Environ. Toxicol. Chem. 19: 3024-3031.

• (15) Belanger, S. E., Rawlings J. M. and Carr G. J. (2013). Use of Fish Embryo Toxicity Tests for the Prediction of Acute Fish Toxicity to Chemicals. Environmental Toxicology and Chemistry 32: 1768-1783. .

• (16) Lammer, E., Carr, G. J., Wendler, K., Rawlings, J. M., Belanger, S. E., Braunbeck, T. (2009) Is the fish embryo toxicity test (FET) with the zebrafish (Danio rerio) a potential alternative for the fish acute toxicity test? Comp. Biochem. Physiol. C Toxicol. Pharmacol.: 149 (2), 196-209

• (17) Chapter C.4 of this Annex: Ready Biodegradability.

• (18) Chapter C.29 of this Annex: Ready Biodegradability, CO2 in sealed vessels.