A category approach to predicting the developmental (neuro) toxicity of organotin compounds: The value of the zebrafish (Danio rerio) embryotoxicity test (ZET)

Reproductive Toxicology 41 (2013) 35–44

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Reproductive Toxicology journal homepage: www.elsevier.com/locate/reprotox

A category approach to predicting the developmental (neuro) toxicity of organotin compounds: The value of the zebrafish (Danio rerio) embryotoxicity test (ZET) Anna Beker van Woudenberg a,∗ , André Wolterbeek a , Lindsey te Brake a , Cor Snel b , Aswin Menke b , Carina Rubingh a , Didima de Groot a , Dinant Kroese a a b

TNO, Research Group Risk Analysis for Products In Development (RAPID), Zeist, The Netherlands TNO Triskelion B.V., Zeist, The Netherlands

a r t i c l e

i n f o

Article history: Received 26 March 2013 Received in revised form 31 May 2013 Accepted 7 June 2013 Available online 21 June 2013 Keywords: Zebrafish embryotoxicity test (ZET) Alternative model Organotins Developmental (neuro) toxicity Grouping Read across Biological verification

a b s t r a c t Zebrafish embryos were exposed to different organotin compounds during very early development (<100 h post fertilization). Morphology, histopathology and swimming activity (in a motor activity test) were the endpoints analyzed. DBTC was, by far, the most embryotoxic compound at all time points and endpoints studied. In fact, we observed a clear concordance between the effects observed in our zebrafish embryo model, and those observed with these compounds in full rodent in vivo studies. All organotin compounds classified as developmental (neuro) toxicants in vivo, were correctly classified in the present assay. Together, our results support the ZET model as a valuable tool for providing biological verification for a grouping and a read-across approach to developmental (neuro) toxicity. © 2013 Elsevier Inc. All rights reserved.

1. Introduction Toxicology testing strategies are constantly being challenged, especially in the past decade when important regulatory changes occurred. For instance, in 2006 the EU approved the legislation known as REACH (Registration, Evaluation, Authorization and restriction of CHemicals), a new law covering the production and use of chemical substances and their potential impacts on both human health and the environment [1]. This regulation requires a rapid hazard and risk assessment of thousands of new and existing chemicals. For this purpose and especially for reproductive and developmental toxicity assessment, a great number of test animals will be required [2]. Additionally, REACH also requires that animal testing should be used only as a last resort, i.e. only after having evaluated all available information about the chemical of interest. Therefore, alternative strategies that replace and/or reduce animal testing are urgently needed. Substantial efforts have already been undertaken to develop alternative methods for the assessment of reproductive and development toxicity [3]. However, none of these assays alone can cover

∗ Corresponding author. Tel.: +31 888661696. E-mail address: [email protected] (A. Beker van Woudenberg). 0890-6238/$ – see front matter © 2013 Elsevier Inc. All rights reserved. http://dx.doi.org/10.1016/j.reprotox.2013.06.067

the whole mammalian reproductive cycle due to its inherent complexity [4]. Currently, only a few alternatives have been validated for developmental toxicity, such as whole embryo culture (WEC), the embryonic stem cells test (EST) and the mammalian micromass (MM) test [5]. Consequently, the most realistic strategy is to use a tiered approach that combines non-testing methods (such as (Q)SAR and read across) with a battery of in vitro techniques and use in vivo studies only as last resource [6,7]. It has been estimated that applying combinations of alternatives, collectively called “intelligent testing strategies” (ITS), could reduce the need for in vivo tests by up to 70% for individual endpoints, resulting in significant savings in testing costs and use of animals [2]. Read across and the grouping of chemicals into categories is one of the ITS approaches and it has already been used in national and international procedures for the safety and risk assessment of chemicals [8,9]. It is also expected to play an important role in the REACH system [10]. Additionally, the grouping of chemicals into categories demonstrating similar biological responses may be one of the first steps toward the identification of structural alerts for different endpoints, and to identifying and building adverse outcome pathways. In aiming for an even faster and cheaper strategy, which would contribute to the 3R’s concept (replacement, reduction and refinement of animal use), the zebrafish (Danio rerio) embryotoxicity test (ZET) might be an excellent alternative as it offers a rapid

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A. Beker van Woudenberg et al. / Reproductive Toxicology 41 (2013) 35–44

Table 1 Overview of organotin compounds used in this study [23].

CAS nr.

Monomethyltin trichloride (MMTC)

Dimethyltin dichloride (DMTC)

Monobutyltin trichloride (MBTC)

Dibutyltin dichloride

Dioctyltin dichloride

(DBTC)

Monooctyltin trichloride (MOTC)

993-16-8

753-73-1

1118-46-3

683-18-1

3091-25-6

3542-36-7

240.8 1 × 105 −2.15 Aldrich 97%

219.7 1 × 105 −2.18 to −3.1 Aldrich 97%

282.2 8.2 0.18 Aldrich 95%

303.8 36 1.89 Aldrich 96%

338.3 0.1 2.14 ABCR 95%

416 1.0 5.82 ABCR 95%

(DOTC)

Molecular structurea

Mw (g/mole) Solubility (g/L) Log P(ow) Supplier Purity

a Characteristically, organotin compounds are described with the following general formula: Rx Sn(L)(4−x) , with R representing an organic alkyl or aryl group and L a relatively weaker organic (or sometimes inorganic) ligand [23]. In this study, tin exists in either divalent (Sn2+ ) or tetravalent (Sn4+ ) cationic (positively charged) ions to form either SnX2 or SnX4 .

external development and the unique advantage of detecting compound effects during the entire period of organogenesis. Zebrafish have also been demonstrated to share many common features with humans in: development, anatomy, physiological responses, metabolism and chemical-induced organ/tissue responses and many molecular pathways are evolutionarily conserved between zebrafish and humans (see reviews [11,12]). At 96 h post fertilization (hpf) all major organs, as well as the sensory organs, are well developed – comparable to a three-month old human embryo [13] – and larvae start swimming at approximately 72 hpf. Therefore ZET screening for developmental (neuro) toxicity can be performed at early stages [14]. According to the new EU Directive 2010/63/EU on the protection of animals used for scientific purposes [15] the earliest life-stages of animals do not fall into the regulatory frameworks dealing with animal experimentation. Independent feeding is considered to be the stage from which free-living larvae are subjected to regulations on animal experimentation, which in zebrafish occurs approximately 120 hpf [16,17]. Therefore, the developing zebrafish can be used as an alternative to in vivo (rodent) models for developmental toxicity and (developmental) neurotoxicity, with the advantages of relatively high throughput and minimizing costs and amount of test compound needed [18]. Moreover, with recent state-of-the-art developments, it has been suggested that ZET could play a role in bridging the gap between in vitro cell-based models and in vivo mammalian models [19]. For these reasons, ZET may be a powerful tool when combined with the category approach, presuming that the ZET ranking of a series of analogs would correspond to the in vivo mammalian ranking both in types of effect and potencies. In that case, there is a strong likelihood that the toxic potency of new compounds within the same structural class, with unknown developmental effects, can be correctly predicted with this test system [20–22]. Therefore, a series of mono- and disubstituted organotins were selected as a compound class of interest because in vivo data showed differences in developmental toxicity and (developmental) neurotoxicity. The aim of the present study was not only to verify whether the ZET model would be able to identify all these compounds as developmental toxic and/or (developmental) neurotoxic, but also to show the different profiles of these compounds

Table 2 Concentration range (␮M) obtained from the DRF experiments. Concentrations within these ranges were used in the subsequent main experiments. MMTC 100–600 a

DMTC 100–1050

MBTC 100–600

DBTC 1–14

MOTC

DOTC a

20–60

a

20–70

Selected test concentrations were limited by substance maximum solubility.

as observed in in vivo rodent studies [23]. The applicability of the ZET model to obtain supplementary data on toxicity to support the category approach in hazard (and risk) evaluation is discussed. 2. Materials and methods 2.1. Fish husbandry and egg production Zebrafish (Wildtype AB line Zebrafish, Zebrafish International Resource Center, Eugene, OR, USA) were housed in self-regulating aquaria (Tecniplast, Tecnilab-BMI, Someren, The Netherlands) under standard laboratory conditions of temperature 28 ◦ C, pH 7.5 and water conductivity 500 ␮S/cm, according to standard zebrafish breeding protocols [24]. Lighting was controlled by a timer to provide a 12/12 h light/dark cycle. Adults were fed with dry flake food (SDS) twice daily and additionally with live food (brine shrimp; Artemia) once daily as recommended by [24] and OECD guidelines [25]. To optimize fecundity, females were housed separately and fed only Artemia for three days before breeding. On the evening prior to the day of breeding, adult fish at a sex ratio of 1:1 were placed into divided spawning boxes with egg traps to prevent egg predation. The next morning, spawning was triggered by the onset of light and removal of the partition. Fertilized eggs were collected approximately 60 min after spawning using 800 ␮m meshes and were rinsed with MilliQ water to remove impurities. Only eggs from batches with a fertilization rate of at least 90% were selected for the experiments. Obtained eggs were transferred to Petri dishes containing aquarium water spiked with 0.05% methylene blue to color non-fertilized eggs or eggs with a damaged membrane [25]. Colored eggs were discarded. The fertilized eggs were then directly transferred into different Petri dishes containing the test compounds at selected concentrations. Subsequently, embryos within the 8–16 cells stage, defined by Kimmel et al. [18], were selected and transferred to a 12-well plate containing 3 mL of the test medium per well. Direct transfer from aquarium water to the test wells was avoided not only to minimize dilution errors, but also to expose the embryo to the test compound as early as possible, so starting with embryos at the same developmental stage (8–16 cells). 2.2. Compounds The organotin compounds used in the present study were: monomethyltin trichloride (MMTC), dimethyltin dichloride (DMTC), monobutyltin trichloride (MBTC), dibutyltin dichloride (DBTC), monooctyltin trichloride (MOTC) and dioctyltin dichloride

A. Beker van Woudenberg et al. / Reproductive Toxicology 41 (2013) 35–44

(DOTC). All reagents were purchased from Sigma–Aldrich (St.Louis, MO, USA) and ABCR (Karlsruhe, Germany). Test solutions were freshly prepared on the morning of each experiment. First, a stock solution of each compound was prepared in DMSO. Subsequently, the stock solution was further diluted with aquarium water to the desired concentrations. The final concentration of DMSO in the water supplemented with the compounds was 0.3%. As negative controls, aquarium water and 0.3% DMSO in aquarium water was used. An overview of the organotin compounds used in this study and their physical/chemical properties is shown in Table 1. 2.3. Experimental design 2.3.1. Dose range finding (DRF) experiments In the DRF studies, the embryos (n = 12 per concentration) were exposed to a relatively wide range of concentrations of each test compound in order to determine the concentration range of interest for further testing. The maximum substance solubility was also taken into account. The concentration range used in the subsequent main experiment is shown in Table 2. The embryos, in the DRF studies, were assessed for lethal and/or developmental toxic effects at 24 and 48 hpf following the criteria described in Table 3. 2.3.2. Main experiments The main studies (MS) were performed after the concentration range of interest had been established in the DRF studies. At least five test concentrations per compound were systematically assigned to all 12 well plates, with one plate containing, as far as possible, all test concentrations and two control groups – aquarium water and 0.3% DMSO. This schedule was followed to assure that, on average, all test concentrations were assessed for potential effects at the same time point of development of an individual embryo/larva. Embryos (n = 12 per concentration) were kept at 28 ± 1 ◦ C with a 12/12 h light/dark cycle and test solutions were not refreshed during the exposure period. Morphological evaluation of the embryos was performed (see Section 2.4) and termination was done shortly after 100 hpf (after motor activity assessment, see Section 2.5), by euthanizing the larvae with a tricaine solution (1000 mg tricaine per liter aquarium water). Subsequently, larvae were processed for histo-pathological investigation (see Section 2.6). For each test concentration at least one replicate was available, but in general there were three. 2.4. Morphological scoring Morphological changes were evaluated according to the method described by Fraysse and coworkers [27] (see Table 3); the evaluation was performed at 24, 48, 72 and 96 hpf. Endpoints were assessed using binary responses (yes/no) as proposed in the DIN standard [28], except for the number of spontaneous movements per minute at 24 hpf which was also quantified. Embryos and larvae were considered dead when coagulation had occurred and/or no heartbeat was detected. Assessment was performed by one observer with the use of a Zeiss Achromat S microscope (Carl Zeiss, Sliedrecht, The Netherlands). Experiments were considered valid if survival rates in controls were >90% [25]. Per experiment, the data of the morphological scores was normalized and expressed as a percentage compared to controls. Per compound, the morphology scores were then combined and presented as mean ± standard error of the mean (SEM). The EC50 was defined as the concentration at which there was a 50% increase (or decrease) in the incidence of a given parameter in comparison with the control (see Section 2.7).

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2.5. Motor activity (MA) test At day 4 (96 hpf), a motor activity (MA) test was performed to assess developmental neurotoxicity. The MA test was performed as described previously [29]. Briefly, MA testing was carried out using a ViewPoint behavior recording system including the accompanying movement tracking and analysis software (Zebrabox, ViewPoint, Lyon, France), which consists of a light box platform and an infrared-sensitive camera within a closed box. Just before the start of the MA test, larvae were transferred to square 96-wells, flat bottom plates (Whatman, GE Healthcare, 7701-1651) in their own test solution (±500 ␮L). The MA test was performed in the afternoon, to ensure steady activity by the zebrafish [30]. Larvae were allowed to acclimatize, for 15 min on average, in the Zebrabox in visible light conditions. Subsequently, larvae were exposed to the following light conditions: 15 min of darkness (i.e. infrared light), 15 min of light (i.e. visible light plus infrared light) and a final 15 min period of darkness (i.e. infrared light), further called sessions Dark I, Light and Dark II respectively. To measure the intensity in the locomotor apparatus, a spherical sensor was placed in the Zebrabox with a closed lid. The light, turned on at the same intensity as used during the experiments, resulted in a measured light intensity of 700 lx, while the infrared light resulted in a measured intensity of 0 lx. Larvae movement was tracked with ViewPoint software. Thermostatically controlled, continuous water flow (27 ◦ C ± 1 ◦ C) into the closed Zebrabox was used to maintain larvae at a constant environmental temperature. The larvae were equally distributed over two plates in a random fashion and the two plates were subsequently tested. All concentrations from the same compound and the respective control were tested the same day. Larvae with a positive score on any of the developmental toxicity endpoints were also measured in the MA test but excluded from data analysis in order not to confound MA results. For each MA test performed, individual larvae tracks were analyzed using the analysis software (Zebrabox) for the Distance Moved (DM) (given in mm) as a measure of locomotor activity. Tracks were analyzed per unit time (1-min or 5-min intervals per 15-min session) as described previously [29]. In the present study, developmental neurotoxicity of zebrafish larvae was studied using the following parameters (see Fig. 1): (i) the Distance Moved (DM) per unit time, i.e. DM15 min , -DM5 min , DM1 min ; (ii) the Slope (SL) of Habituation (H), i.e. HSL; and (iii) the Startle Response (StR). All parameters, except the Startle Response, were measured within the three periods of 15-min test session each, i.e.: Dark I (0–15th min); Light (16th–30th min) and Dark II (31st–45th min). The Startle Response was defined as the immediate reaction of the larvae to a sudden change of the light condition, such as from the Dark I to the Light period (i.e. the Lightness Startle Response; L-StR) and from the Light to the Dark II (i.e. the Darkness Startle Response; D-StR). The latter transition (D-StR), especially results in an instantaneous and drastic change of the motility pattern. D-StR was quantified by calculating per individual the difference (in mm) between: average Distance Moved (avgDM) over the last 5 min in the Light period (L-avgDM26th to 30th min ) and the avgDM of the first 5 min in the Dark II period (DII-avg DM31st to 35th min ). The Lightness Startle Response L-StR was derived along similar lines. Thus, respective group means for the Darkness D-StR and Lightness L-StR Startle Responses were calculated per group as follows, where Ng = number of individuals per group and g = 1 to total number of groups: D StRg =



 i=1 to Ng

(DII avgDM(31st to 35th min) − L avgDM(25th to 30th min) )i

/Ng

and L StRg =



 i=1 to Ng

(L avgDM(16th to 20th min) − DI avgDM(11th to 15th min) )i

/Ng

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Table 3 Overview of the toxicological endpoints assessed for each life stage in the ZET. Toxicological endpoints Time (life stagea ) of evaluation

Lethal effects

Developmental effects

a b c d

Coagulated eggsb Development of somites Tail detachment Presence of heartbeat Hatching Spontaneous movements per minutec Developmental delay Decreased pigmentation Malformation of eyes/otholits/heart/yolksac/chorda/tail Malformation of chorda/tail

24 hpf

48 hpf

72 hpf

x x x

x x x x

x

x x

x

x x x

x xd xd

Early life stage terminology according to Baxter [26]: embryo, stage before hatching; pro-larva, stage between hatching and yolk-sac resorption. Coagulated eggs are milky white and appear dark under the microscope. Parameter is quantified. Evaluation of those parameters was also done at 96 hpf when they were not conclusive at 72 hpf (e.g. due to delayed hatching).

2.6. Histopathology

2.7. Statistical analysis

After the MA test, the larvae were euthanized and subsequently fixed overnight in Bouin’s fixative (Klinipath; Duiven, The Netherlands), embedded in paraffin and sectioned down completely into serial 4 ␮m thick saggital sections. After staining with hematoxylin and eosin (H&E), all sections were analyzed using an Axioskop 2 plus microscope (Zeiss). In each experiment, five to six larvae from each concentration were analyzed.

For morphology examination, a dose-response curve was fitted for each relevant endpoint (Table 3) and EC50 concentrations were calculated using Sigma Plot (Version 9.01, Systat Software, Chicago IL, USA) by curve fitting to the relative incidence data. The following 4-parameter logistic equation was used for curve fitting: y = A + ((B − A)/(1+((C/x)D))); where A = minimum y (constrained to > 0), B = maximum y (constrained to < 100), C = the EC50 value, and D = slope factor.

Fig. 1. Distance Moved (given in mm) per minute in the motor activity test of larvae (96 hpf) kept in 0.3% DMSO vehicle and larvae kept in aquarium water (Mean ± SEM). For illustration, all parameters analyzed in the motor activity test are also indicated in this figure. Swimming activity of the larvae is used to study normal behavior of the larvae, and the effects of chemicals thereon. The swimming activity is tested during three sequential periods (sessions) – alternating dark and light – of 15 min duration each, i.e. Dark I (0–15th min); Light (16th–30th min) and Dark II (31st–45th min). The test is preceded by an acclimatization period in light (duration on average 15 min (not shown)). Parameters measured are: (i) Distance Moved (given in mm) per unit time, i.e. per total session of 15 min DM15 min , per minute DM1 min , to enable analysis of activity over a session with time such as e.g. Habituation, and per 5 min DM5 min (1) (2) (3) (4), to enable calculation of the startle response at the transition zones from Dark I to Light (1) (2) and from Light to Dark II (3) (4). (ii) Habituation H over time is calculated from the slope SL of the curve HSL per 15 min session, i.e. for the Dark I period DI-HSL, and the Dark II period DII-HSL. (iii) Darkness and Lightness Startle Response (D-StR and L-StR, respectively). The Startle Response was defined as the immediate reaction of the larvae to a sudden change of the light condition, such as from the Dark I to the Light period (i.e. the Lightness Startle Response L-StR and from the Light to the Dark II (i.e. the Darkness Startle Response D-StR). The Darkness Startle Response is quantified by calculating – per individual – the difference (in mm) between: average distance moved (avgDM) over the last 5 min in the Light period (L-avg DM26th to 30th min (3)) and the avgDM of the first 5 min in the Dark II period (DII-avg DM31st to 35th min (4)). The Lightness Startle Response L-StR was derived along similar lines (1 and 2). Notice that, to calculate the startle response per individual, the average DM per min, calculated over 5 consecutive (steady) minutes, was chosen instead of taking the value of the last minute only. For further details the reader is referred to the text in Section 2.5.

A. Beker van Woudenberg et al. / Reproductive Toxicology 41 (2013) 35–44

All of the derived parameters studied (Distance Moved, Startle Response and Slope of Habituation) were analyzed using a mixed model. Compound/concentration was used as a fixed factor and plate and row within plate was used as a random factor. In all tests performed, the hypothesis was rejected at the level of 5% of probability (alpha = 0.05). A Dunnett’s test was used for post hoc testing. 3. Results 3.1. Morphology Only experiments where the embryo survival rate of both control groups (aquarium water and 0.3% DMSO) were above 90% were used. At 72 hpf, the mean survival rate for control embryos in 0.3% DMSO vehicle (n = 10) was 97.5 ± 1.3% and for control embryos in aquarium water (n = 10) it was 99.2 ± 0.8%. The critical effects (based on EC50 ) on morphology of embryos exposed to organotins at different time points is presented in Table 4. For MMTC, the most critical effect was the survival rate as measured by heart beat or coagulation (at 24 and 48 hpf). The decreased survival rate was concentration-dependent. Also, a decreased hatching rate at 72 hpf was observed, with an EC50 of 299 ␮M for MMTC. In embryos that survived and were hatched at higher concentrations (>300 ␮M), an increased incidence of malformations, such as tail malformations, was observed at 96 hpf. In the group of embryos exposed to DMTC, the most critical effect observed was a decrease in hatching rate at 72 hpf. No significant decrease in survival or increase in malformations was observed at 24 and 48 hpf. Similar to the effects of MMTC, survival rate was the most critical effect for embryos exposed to MBTC at 24 and 48 hpf. Moreover, a decreased hatching rate was observed with an EC50 of 311 ␮M at 72 hpf. Also, similarly to MMTC, embryos that survived exposure to higher concentrations of MBTC (>300 ␮M) showed an increased incidence of malformations, such as tail malformations at 96 hpf. In contrast to the other organotins, embryos exposed to DBTC scored positive on multiple lethal and developmental toxicity endpoints in a concentration-related way. DBTC was by far the most developmental toxic of all the compounds investigated. Exposure to DBTC resulted in a decreased number of embryos displaying normal spontaneous movement (>2/min) at 24 hpf (Fig. 2a). Similar DBTC concentrations also resulted in an increased incidence of embryos showing decreased pigmentation at 48 hpf (Fig. 2b). Embryos exposed to ≥8 ␮M DBTC had reduced hatching rates and the EC50 for this parameter was 10 ␮M (Fig. 2c). Further, an increase in tail malformations for larvae exposed to DBTC at 96 hpf is shown (Fig. 2d), with an EC50 of 9 ␮M. Overall, for DBTC, developmental toxicity endpoints are positively assessed at much lower concentrations than decreased survival rate – as indicated by an increased percentage of embryos that coagulated or had no heartbeat (Fig. 2). The tested range of MOTC and DOTC concentrations (highest dose 60 and 70 ␮M, respectively) was found to have no effects on the survival or development of the embryos and larvae (data not shown). Higher concentrations could not be tested, since the maximum solubility had already been reached. 3.2. Motor activity test The MA was always performed in the afternoon after the last morphological examination. The results of the MA test performed

39

for larvae (96 hpf ± 2) kept in vehicle, i.e. 0.3% DMSO or aquarium water are shown in Fig. 1. Significant differences between the 0.3% DMSO and aquarium water groups were not found for any of the parameters studied, i.e. (1) the Darkness and Lightness Startle Response (D-StR and L-StR, respectively); (2) the Habituation Slope (HSL) of the Dark I (DI-HSL) and Dark II session (DII-HSL); 3) the Distance Moved during the 15 min of the: Dark I session (DI-DM15 min ), the Light session (L-DM15 min ) and the Dark II session (DII-DM15 min ). In the control groups (aquarium water and 0.3% DMSO), a typical overall activity pattern was observed: high activity (DM15 min ) during the Dark I and Dark II periods, and low activity during the Light period (Fig. 1). During the Dark I and Dark II sessions the activity starts out high (150–200 mm/min) and gradually decreases over the 15 min test session, i.e. the Habituation Slope (HSL). Typically, low activity (<25 mm/min) was observed during the Light period. The change in activity at the transition zone from the Light session to Dark II, i.e. the Darkness Startle Response (D-StR), occurs rather suddenly and is well pronounced; in contrast, the Lightness Startle Response (L-StR) at the transition from the Dark I to Light session is small. Fig. 3 shows the effects of 0.3% DMSO (vehicle control group) and different concentrations of MMTC (Fig. 3a), DMTC (Fig. 3b), MBTC (Fig. 3c) and DBTC (Fig. 3d) on the different MA parameters studied for the Dark I, Light and Dark II 15 min test sessions, as described in Section 2.5. Results of the statistical assessment are presented in the right panel of Fig. 3. Evaluation of these results showed the following: MMTC (Fig. 3a), showed a concentration-related increase of activity (DM15 min ) in both the Dark I and Dark II sessions. In the groups of larvae exposed to 200, 250 or 300 ␮M MMTC the DM15 min was significantly different than the control group. The Habituation Slope was only significant in Dark I (DI-HSL) and only for the 300 ␮M MMTC group. It was noticed that, both at the start and at the end of the Dark I and Dark II sessions, the activity was concentrationrelated increased. The activity change – Startle Response – from the Light to the Dark II period, the Darkness Startle Response D-StR, was not affected; in contrast, the Lightness Startle Response, L-StR, was affected significantly in the 250 ␮M MMTC group. DMTC (Fig. 3b), showed concentration-related changes in the Slope of Habituation in Dark I (DI-HSL) for the 450 and 600 ␮M DMTC groups. Also, the Darkness Startle Response D-StR was significantly reduced in the 600 ␮M DMTC group. In fact, the overall activity – DM15 min – of the 600 ␮M DMTC group was reduced in Dark I (non-significant, p = 0.9726) and Dark II (p < 0.01). MBTC (Fig. 3c), at the concentrations and in the circumstances here, did not affect any of the MA parameters investigated. DBTC (Fig. 3d) reduced significantly (p < 0.05) the overall activity (DM15 min ) in both darkness periods for the groups with higher concentrations (6 and 8 ␮M). The Darkness Startle Response (D-StR) and the Slope of Habituation in Dark I (DI-HSL) were also significantly (p < 0.05) reduced for the 6 and 8 ␮M DBTC groups. However, for the Habituation Slope in Dark II (DII-HSL) only 8 ␮M DBTC group was significantly different. The 6 ␮M DBTC group showed a clear decrease for DII-HSL, but this did not reach the level of statistical significance. 3.3. Histopathology Histopathological analyses were performed on larvae sampled at 100 hpf, after the MA test. There were no significant differences in histopathological analysis between larvae kept in aquarium water or in 0.3% DMSO (data not shown). The main histopathological findings of larvae exposed to the different organotins are presented in Fig. 4. Larvae exposed to 150 ␮M MMTC revealed no histopathological abnormalities (not shown). At 250 ␮M MMTC, only 16.6% of the larvae (n = 6) examined

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A. Beker van Woudenberg et al. / Reproductive Toxicology 41 (2013) 35–44

Table 4 The most critical effects (based on EC50 ) on morphology of zebrafish embryos observed in the ZET at 24, 48 and 72 hpf for MMTC, DMTC, MBTC and DBTC. 48 hpf Most critical effect EC50 (␮M)

24 hpf Most critical effect EC50 (␮M) MMTC DMTC MBTC DBTC

Survival No effect observed Survival Spontaneous movements

(395) – (408) (9)

Survival No effect observed Survival Decreased pigmentation

histopathologically, were not hatched. However, at 350 ␮M MMTC the majority of the larvae (67%) were not hatched (Fig. 4.1b); in the other 33%, the development was disrupted completely, showing a proteinaceous blob with some cells in it. At 450 ␮M MMTC, development was disrupted completely in all larvae studied (n = 5) (Fig. 4.1c). Larvae exposed to 300 ␮M or 450 ␮M DMTC revealed no histopathological abnormalities at 100 hpf (not shown). However, at 600 ␮M DMTC, 83% of the exposed larvae showed degeneration

b)

24 hours post fertilization following DBTC exposure

100

80

60

40

20

0 0.000

(389) – (391) (11)

0.002

0.004

0.006

0.008

0.010

0.012

0.014

0.016

100

80

60

40

20

0 0.000

0.002

% of alive larvae concerned relative to controls

% of alive larvae concerned relative to controls

d)

100

80

60

40

20

0.004

0.006

0.008

0.010

0.012

0.006

0.008

0.010

0.012

0.014

0.016

% of alive embryos with decreased pigmentation % survival of embryos at 48 hpf

72 hours post fertilization following DBTC exposure

0.002

0.004

Concentration DBTC (mM)

% of alive embryos with normal spontaneous movement (> 2/min) % survival of embryos at 24 hpf

0 0.000

(299) (697) (311) (9)/(10)

48 hours post fertilization following DBTC exposure

Concentration DBTC (mM)

c)

Hatching Hatching Hatching Tail malformation/hatching

of cells in the eye and in the brain (Fig. 4.2b). Exposure to 750 ␮M DMTC (Fig. 4.2c) or higher concentrations (900 and 1050 ␮M) revealed no degeneration of cells in these regions. Instead, development of the larvae appears to have been delayed and/or arrested since the larvae had not hatched. Regarding MBTC, except for the non-hatching effect (Table 4), no other histopathological abnormalities were observed at any of the concentrations studied; therefore, no figure for MBTC is presented.

% of alive embryos concerned relative to controls

% of alive embryos concerned relative to controls

a)

72 hpf Most critical effect EC50 (␮M)

0.014

Concentration DBTC (mM) ___ % of alive larvae that have hatched at 72 hpf _ _ % larvae of embryos at 72 hpf

0.016

96 hours post fertilization following DBTC exposure

100

80

60

40

20

0 0.000

0.002

0.004

0.006

0.008

0.010

0.012

0.014

0.016

Concentration DBTC (mM) % of alive larvae with a malformed tail at 96 hpf

Fig. 2. Results of the morphology assessment of embryos exposed to DBTC at 24 (a), 48 (b), 72 (c) and 96 (d) hpf. Results are expressed in mean ± SEM.

A. Beker van Woudenberg et al. / Reproductive Toxicology 41 (2013) 35–44

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Fig. 3. Effect of organotins on the swimming activity of zebrafish larvae in the motor activity test. Left panel shows the graphics for the Distance Moved DM- per 1-min intervals for larvae exposed to MMTC (a), DMTC (b), MBTC (c) and DBTC (d) (mean ± SEM; n = 12/group/concentration). Right panel presents, respectively, the statistical evaluation of the data for the motor activity.♦ Concentration, *p < 0.05, **p < 0.01 and ***p < 0.001.

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Fig. 4. Histopathology of zebrafish larvae (100 hpf) exposed to: (1) MMTC, (2) DMTC and (3) DBTC. *Degeneration in the eye and in brain regions; **apoptosis in the brain.

Larvae exposed to 2, 4 or 6 ␮M DBTC revealed no histopathological abnormalities at 100 hpf (not shown). However, 80% and 100% of the larvae exposed to 8 and 10 ␮M DBTC, respectively, showed areas of apoptosis in the brain (Fig. 4.3b and 3c, respectively). 4. Discussion In the present study, the aim was to verify whether the investigated compounds within the organotin group exhibit comparable developmental and/or neurotoxic profiles in the ZET as found in in vivo studies in rodents. Morphology was performed aiming to detect developmental toxic effects; the MA test was carried out in order to identify (developmental) neurotoxicity and histopathology was performed to confirm the effects at microscopically levels. The objective behind this investigation is that: if the ZET would provide a correct prediction of the type of effects, as well as their relative potencies, there is a high likelihood that it will also correctly predict the toxic profiles of new compounds within this group of organotin compounds not yet tested for developmental effects. In

this way, the ZET could be a valuable alternative model in the hazard assessment of substances that have structural analogs with adequate in vivo developmental toxicity test data. This could avoid of full in vivo (guideline) testing of these new substances. From the present results, a number of conclusions can be made. First of all, the tin-octyl derivatives (DOTC and MOTC) could not be identified as developmental toxicants. This is most probably due to their low solubility. As with other alternative (in vitro) tests, the solubility and Log P(ow) of test compounds need to be taken into account, since they might be the bottleneck for testing sufficiently high concentrations. Also, like the absorption barriers in rodents in vivo, the chorion might hamper the absorption of some classes of compounds, potentially resulting in false negative results. Second, and most importantly, the ZET correctly predicted all other organotin compounds tested. All compounds were correctly classified for developmental toxicity, but they were also correctly predicted with regard to their developmental toxicity and (developmental) neurotoxicity profiles. DBTC was, by far, the most embryotoxic compound at all time points studied, being approximately 30 times

A. Beker van Woudenberg et al. / Reproductive Toxicology 41 (2013) 35–44 Table 5 Comparison of the ranking of the organotins, for developmental (neuro) toxicity, between in vivo studies and the ZET model. (a) In vivoa (Developmental) neurotoxicity Developmental toxicity

DMTC > DBTC > MMTC yes yes Ambiguous

MBTC No data

DBTC > DMTC » MBTC yes yes No

MMTC No data

(b) Zebrafish embryotoxicity test (ZET) Developmental neurotoxicityb

DBTC > DMTC > MMTC > MBTC yes yes Ambiguous No

Developmental toxicityc

DBTC > DMTC yes yes

MBTC No

= MMTC No

a The in vivo effects of various mono- and disubstituted tin compounds on (neuro) developmental toxicity were reviewed by WHO [23]. In this review, DMTC was reported to be developmentally toxic. Severe developmental effects (reduced fetal weights, fetal death, cleft palate) were observed at maternal toxic dose levels. A NOAEL of 10 mg/kg body weight was derived for developmental toxicity. In a recent evaluation of DMTC by the ECHA Committee for Risk Assessment (RAC), DMTC was classified as a reproductive toxicant for effects seen on development [36]. In the WHO report [23], DMTC was considered a neurotoxicant (NOAEL of 0.6 mg/kg body weight for neurotoxicity) based on neuropathological lesions observed in the hippocampus and surrounding cortical regions. At higher dose levels, convulsions and tremors were also reported. In the evaluation of DMTC by RAC [36,37], inconsistent neurodevelopmental effects of DMTC were reported (effects observed in runaway and water maze tests). In the WHO report [23] and in an IUCLID (http://iuclid.echa.europa.eu/) evaluation of DBTC, a review of animal data consistently showed DBTC to cause dose-dependent developmental toxicity, such as fetal deaths, birth defects and decreased fetal weights. An overall NOAEL of 2.5 mg/kg body weight was derived for developmental toxicity[23]. In the WHO report [23] no neurotoxic effects for DBTC were described but in a study by Jenkins et al. [31] in utero exposure to DBTC caused an increased incidence of apoptosis in the hippocampus and neocortex at postnatal day 38. A NOAEL of 2.5 mg/kg body weight was found in this study. Both in the WHO report [23] and in a proposal for harmonized classification and labeling [37], there are no data on the prenatal developmental toxicity of MMTC. In a study by Noland et al. [38] in utero and lactational exposure of male rats resulted in slight cognitive deficits but these results could not be confirmed in a series of studies by Moser et al. [39]. Therefore, it was concluded that the effects of MMTC on neurodevelopment were ambiguous. Based on a series of prenatal developmental toxicity studies in rats not showing effects on fetuses or only at maternally toxic dose levels, it was concluded that MBTC was not developmentally toxic [23]. A NOAEL of >400–2000 mg/kg body weight was derived. No data were available on the (developmental) neurotoxic potential of MBTC. b Ranking based on the results of the motor activity test (Section 3.2 and Fig. 3) and histopathology (Section 3.3 and Fig. 4). c Ranking based on the results of morphology (Section 3.1 and Table 4) and histopathology (Section 3.3 and Fig. 4).

more potent than the other analogs studied. The MA test also showed similar results. At a concentration as low as 6 ␮M DBTC, a significant difference on the Startle Response for Dark II was already observed. Together, the MA results indicate that of the organotin compounds studied here, DBTC is the most potent (developmental) neurotoxic compound. For the same parameter, DMTC showed effects only at concentrations 100× higher (600 ␮M). This is in agreement with Jenkins et al. [31], who also showed that DBTC was a neurotoxicant 40 times more potent than its analog substitute trimethyltin (TMT).Thus, if we compare the organotin compounds classification resulting from in vivo rodent studies with the classification obtained in the present study using the ZET (see Table 5), we observe a clear accordance in the classification: all (organotin) compounds classified in vivo as developmental toxic and (developmental) neurotoxic were correctly classified, although developmental neurotoxicity ranking between DMTC and DBTC differs slightly between in vivo (WHO) and ZET. With regard to this ranking it should be recognized that different potency values for the same compound can occur, depending on the dosing regimens used (absolute and interval values), (animal) strain, and/or just response variability. For instance, Janer et al. [32], retrospectively compared the (rat) two-generation reproductive toxicity study with the (rat)

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subchronic toxicity study and found that NOAELs observed in these types of studies may differ by an order of magnitude for a single substance. Thus, observed small absolute potency differences of two substances in vivo should not be taken as representative of their absolute in vivo ranking. The intrinsic complexities of the anatomy and physiology of the nervous system and the reproductive cycle in mammals are the main reasons why none of the currently available (in vitro) alternatives can fully replace the in vivo studies in mammals for assessing developmental (neuro) toxicity, whether as stand alone test or when combined. However, within the context of a chemical grouping (in vitro) alternatives potentially might replace in vivo studies when for one or more of the analogs adequate in vivo (guideline) test data are available: i.e. when in this situation it can be shown that the (in vitro) alternative(s) reflect observed in vivo responses. This category approach has already been used to assess reproductive toxicity for compounds such as phthalates [10], but their data also show that the structural properties alone are insufficient for use as a basis for read across. Therefore, some complementary biological verification model would be important in supporting the read across: i.e. to show that structural similarity is reflected in a similar response in an adequate biological model. The ZET has already demonstrated a high degree of sensitivity in identifying developmentally toxic substances [33,34]. Hagenaars et al. [35], also using the ZET, investigated the developmental toxicity of perfluorinated chemicals (PFC’s) with different functional groups and chain lengths and although general effects were observed in all classes, some specific effects observed were related to the structure of the tested chemicals, showing that the ZET model was also able to identify such specific effects. Recently, Hermsen et al. [22], showed that the ZET correctly predicted the in vivo potencies for developmental toxicity for glycol ethers and triazoles. With the results presented in the present paper, and those published by others, it is clear that the ZET may be a very valuable tool to further substantiate read across within chemical groups in which at least one of the members has adequate (guideline) in vivo test data. When testing both the chemicals of interest, and the source chemical with in vivo test data (to which read across is to be proposed) in the ZET, they should show similar effect profiles for providing this support. In order to serve such a function, the ZET model ideally should be able to pick up all different types of critical effects which can be detected by in vivo studies. We assume that assessment of developmental neurotoxicity might be improved in the future once the MA part in the ZET has been evaluated and standardized for a number of typical neuro read-outs representing different aspects of the developing nervous system (this is beyond the scope of the present paper). Taken together, our data showed that, the ZET seems to be a very sensitive and accurate model in detecting developmental toxicity and (developmental) neurotoxicity. It is recognized that the solubility and Log P(ow) of the compounds tested are to be considered, as they may give rise to false negative conclusions. Extension of the ZET model with a motor activity test would further improve the predictability of this model for developmental neurotoxicity. Based on this, we believe that the ZET model represents a potential tool for providing biological verification for a grouping and read across proposal.

Acknowledgements This work was partially financed by the NTC (Netherlands Toxicogenomics Center) and also supported by the Commission of the European Communities, the collaborative project ChemScreen (GA244236).

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References [1] European Commission (EC). Regulation (EC) No 1907/2006 of the European Parliament and of the Council of 18 December 2006 concerning the Registration, Evaluation, Authorization and Restriction of Chemicals (REACH), establishing a European Chemicals Agency, amending Directive No 1488/94 as well as Council Directive 76/769/EEC and Comission Directives 91/155/EEC, 93/105/EC and 2000/21/EC. Official Journal of the European Union L 2007 2007;136:3–280. [2] van der Jagt K, Munn S, Tørsløv J, de Bruijn J. Alternative approaches can reduce the use of animals under REACH 2004; EUR 21 405 EN:1-31. [3] Adler S, Basketter D, Creton S, Pelkonen O, Van Benthem J, Zuang V, et al. Alternative (non-animal) methods for cosmetics testing: current status and future prospects-2010. Archives of Toxicology 2011;85:367–485. [4] Schenk B, Weimer M, Bremer S, van der Burg B, Cortvrindt R, Freyberger A, et al. The ReProTect Feasibility Study, a novel comprehensive in vitro approach to detect reproductive toxicants. Reproductive Toxicology 2010;30:200–18. [5] Pamies D, Martinéz CE, Sogorb MA, Vilanova E. Mechanism-based models in reproductive and development toxicology. In: Gupta RC, editor. Reproductive and developmental toxicology. San Diego, CA, USA: Academic Press–Elsevier; 2011. p. 135–46. [6] van der Burg B, Kroese ED, Piersma AH. Towards a pragmatic alternative testing strategy for the detection of reproductive toxicants. Reproductive Toxicology 2011;31:558–61. [7] Piersma AH. Innovations in testing strategies in reproductive toxicology. Methods in Molecular Biology 2013;947:327–41. [8] Hanway RH, Evans PF. Read-across of toxicological data in the notification of new chemicals. Toxicology Letters 2000;116:61. [9] Combes R, Barratt M, Balls M. An overall strategy for the testing of chemicals for human hazard and risk assessment under the EU REACH system. ATLA Alternatives to Laboratory Animals 2003;31:7–19. [10] Fabjan E, Hulzebos E, Mennes W, Piersma AH. A category approach for reproductive effects of phthalates. Critical Reviews in Toxicology 2006;36:695–726. [11] Goldsmith JR, Jobin C. Think small: Zebrafish as a model system of human pathology. Journal of Biomedicine and Biotechnology 2012;201:2, http://dx.doi.org/10.1155/2012/817341. [12] Lieschke GJ, Currie PD. Animal models of human disease: zebrafish swim into view. Nature Reviews Genetics 2007;8:353–67. [13] Parng C, Seng WL, Semino C, McGrath P. Zebrafish: a preclinical model for drug screening. Assay and Drug Development Technologies 2002;1:41–8. [14] Fleming A. Zebrafish as an alternative model organism for disease modelling and drug discovery: implications for the 3Rs. NC3Rs 2007;10:1–7 www.nc3rs.org.uk [15] European Commission (EC). EU. Directive 2010/63/EU of the European parliament and of the council of 22 September 2010 on the protection of animals used for scientific purposes. Official Journal of the European Union L 2010 2010;276:33–79. [16] Belanger SE, Balon EK, Rawlings JM. Saltatory ontogeny of fishes and sensitive early life stages for ecotoxicology tests. Aquatic Toxicology 2010;97:88–95. [17] Strähle U, Scholz S, Geisler R, Greiner P, Hollert H, Rastegar S, et al. Zebrafish embryos as an alternative to animal experiments – a commentary on the definition of the onset of protected life stages in animal welfare regulations. Reproductive Toxicology 2012;33:128–32. [18] Kimmel CB, Ballard WW, Kimmel SR, Ullmann B, Schilling TF. Stages of embryonic development of the zebrafish. Developmental Dynamics 1995;203:253–310. [19] Sukardi H, Chng HT, Chan ECY, Gong Z, Lam SH. Zebrafish for drug toxicity screening: bridging the in vitro cell-based models and in vivo mammalian models. Expert Opinion on Drug Metabolism and Toxicology 2011;7:579–89.

[20] de Jong E, Louisse J, Verwei M, Blaauboer BJ, van de Sandt JJM, Woutersen RA, et al. Relative developmental toxicity of glycol ether alkoxy acid metabolites in the embryonic stem cell test as compared with the in vivo potency of their parent compounds. Toxicological Sciences 2009;110:117–24. [21] Hefter R, Hernandez O, Sayre P. Development of chemical categories in the HPV challenge program. US Environmental Protection Agency 1999:1–16 http://www.epa.gov/chemrtk/categuid.htm [22] Hermsen SAB, van den Brandhof E, van der Ven LTM, Piersma AH. Relative embryotoxicity of two classes of chemicals in a modified zebrafish embryotoxicity test and comparison with their in vivo potencies. Toxicology In Vitro 2011;25:745–53. [23] World Health Organization (WHO). Mono-and disubstituted methyltin, butyltin, and octyltin compounds. In: Concise international chemical assessment document; 2006. p. 73. [24] Westerfield M. The zebrafish book: a guide for the laboratory use of zebrafish (Danio rerio). 5th ed. Eugene, OR, USA: University of Oregon press; 2007. [25] OECD. Guideline for the testing of chemicals. Draft proposal for a new guideline Fish Embryo Toxicity (FET) test; 2006. [26] Baxter J. Pattern and variety in development. In: Hoar WS, Randall DJ, editors. Fish physiology. San Diego, USA: Academic press; 1988. p. 1–58. [27] Fraysse B, Mons R, Garric J. Development of a zebrafish 4-day embryo-larval bioassay to assess toxicity of chemicals. Ecotoxicology and Environmental Safety 2006;63:253–67. [28] DIN. Basivalidierung genormter verfahre zur wasser-,abwasser-,und schlammuntersuchung validierungsdokument (38415-T6); 2002. p. 27. [29] de Esch C, van der Linde H, Slieker R, Willemsen R, Wolterbeek A, Woutersen R, et al. Locomotor activity assay in zebrafish larvae: influence of age, strain and ethanol. Neurotoxicology and Teratology 2012;34:425–33. [30] MacPhail RC, Brooks J, Hunter DL, Padnos B, Irons TD, Padilla S. Locomotion in larval zebrafish: influence of time of day, lighting and ethanol. Neurotoxicology 2009;30:52–8. [31] Jenkins SM, Ehman K, Barone Jr S. Structure-activity comparison of organotin species: dibutyltin is a developmental neurotoxicant in vitro and in vivo. Development in Brain Research 2004;151:1–12. [32] Janer G, Hakkert BC, Piersma AH, Vermeire T, Slob W. A retrospective analysis of the added value of the rat two-generation reproductive toxicity study versus the rat subchronic toxicity study. Reproductive Toxicology 2007;24: 103–13. [33] Ton C, Lin Y, Willett C. Zebrafish as a model for developmental neurotoxicity testing. Birth Defects Research Part A: Clinical and Molecular Teratology 2006;76:553–67. [34] McGrath P, Li C. Zebrafish: a predictive model for assessing drug-induced toxicity. Drug Discovery Today 2008;13:394–401. [35] Hagenaars A, Vergauwen L, De Coen W, Knapen D. Structure–activity relationship assessment of four perfluorinated chemicals using a prolonged zebrafish early life stage test. Chemosphere 2011;82:764–72. [36] European Chemicals Agency (ECHA) – Committee for Risk Assessment RAC. Annex 1:Background document to the Opinion proposing harmonised classification and labelling at Community level of Dimethyltin dichloride. 2012;CLH-O-0000001701-83-03/A1. [37] European Chemicals Agency ECHA. CLH report: proposal for harmonised classification and labelling; Trichloromethylstannane(MMTC); 2010. [38] Noland EA, Taylor DH, Bull RJ. Monomethyl- and trimethyltin compounds induce learning deficiencies in young rats. Neurobehavioral Toxicology and Teratology 1982;4:539–44. [39] Moser VC, Barone Jr S, Phillips PM, McDaniel KL, Ehman KD. Evaluation of developmental neurotoxicity of organotins via drinking water in rats: Monomethyl tin. Neurotoxicology 2006;27:409–20.