Embryonic Stage-Dependent Teratogenicity of Ketamine in Zebrafish

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Embryonic stage-dependent teratogenicity of ketamine in zebrafish (Danio rerio) Luis M. Felix, Cindy Serafim, Ana M. Valentim, Luis M. Antunes, Sónia Campos, Manuela Matos, and Ana M. Coimbra Chem. Res. Toxicol., Just Accepted Manuscript • DOI: 10.1021/acs.chemrestox.6b00122 • Publication Date (Web): 30 Jun 2016 Downloaded from http://pubs.acs.org on July 1, 2016

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Chemical Research in Toxicology

Title: Embryonic stage-dependent teratogenicity of ketamine in zebrafish (Danio rerio)

Author names: Luís M. Félix

†,‡,§,*

, Cindy Serafim ┴, Ana M. Valentim †,‡,§, Luís M.

Antunes †,‡,§,║, Sónia Campos †,‡,§, Manuela Matos ∇,O, Ana M. Coimbra †

Affiliations: †

Centre for the Research and Technology of Agro-Environmental and Biological

Sciences (CITAB), University of Trás-os-Montes and Alto Douro (UTAD), Vila Real, Portugal. ‡

Laboratory Animal Science (LAS), Institute for Molecular and Cell Biology (IBMC),

University of Porto (UP), Porto,Portugal. §

Institute for Research and Innovation in Health (i3S), University of Porto (UP), Porto,

Portugal. ┴

Life Sciences and Environment School (ECVA), University of Trás-os-Montes and

Alto Douro (UTAD), Vila Real, Portugal. ║

School of Agrarian and Veterinary Sciences (ECAV), University of Trás-os-Montes

and Alto Douro (UTAD), Vila Real, Portugal. ∇

Biosystems & Integrative Sciences Institute (BioISI), Faculty of Sciences, University

of Lisboa, Lisboa, Portugal. O Department

of Genetics and Biotechnology (DGB), University of Trás-os-Montes and

Alto Douro (UTAD), Vila Real, Portugal

Corresponding author: Luís Félix, Centre for the Research and Technology of AgroEnvironmental and Biological Sciences (CITAB), University of Trás-os-Montes and Alto Douro (UTAD), Quinta de Prados, 5001-801 Vila Real, Portugal. Phone: 00351 1 ACS Paragon Plus Environment


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259

350 000.

Fax:

00351

259

350 480

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e-mail:

[email protected]

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Abstract Ketamine, a widely used anesthetic, has been shown to have NMDA receptor dependent and

independent

actions

during

zebrafish

(Danio

rerio)

embryogenesis.

Notwithstanding, the effects of developmental toxicity and the mechanisms of ketamine action on fish embryos are still not well understood, and its implications for early vertebrate development remains to be clarified. In this work, zebrafish embryos were exposed to ketamine (0.2, 0.4 and 0.8 mg mL-1) in order to study stage-developmental toxicity of this pharmaceutical. During 256-cell (2.5 hours post fertilization- hpf), 50% epiboly (5.5 hpf) and 1-4 somites (10.5 hpf), embryos were exposed to the referred ketamine concentrations for a period of 20 minutes and were allowed to grow until 144 hpf. Both lethal and non-lethal parameters were evaluated. Skeletal development was assessed by alcian blue and calcein staining. Additionally, the expression of the developmental genes sonic hedgehog a (shh a) and noggin 3 (nog3) was evaluated. Similar to our previous work, bone and cartilage malformations were observed after 256-cell exposure. During 50% epiboly, ketamine exposure induced a concentrationdependent mortality and malformations, such as lordosis and/or kyphosis and microcephaly, namely at higher concentrations. Conversely, exposure during 1-4 somites showed the induction of non-specific effects with no rise in mortality. The quantitative real-time polymerase chain reaction (qRT-PCR) analysis showed differences on shh a and nog3 expressions comparatively to the control group. Overall, this study shows that the ketamine toxic profile is developmental phase-dependent with 256-cell being the most susceptible phase. The effects observed may result from ketamine interaction with cellular signaling pathways that merits further investigation.

Keywords



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Zebrafish development; Ketamine; Toxicity; Developmental genes; Cartilage; Bone.


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1. Introduction Ketamine, a widely used dissociative anesthetic, is frequently consumed in nonmedical situations, including pregnancy 1 with possible consequences for the infant.2 Over the years, increasing experimental evidence has shown that exposure to ketamine may be neurotoxic during specific developmental stages, especially when rapid synaptogenesis occurs.3-7 To date, ambiguous mechanisms involving apoptosis and NMDA receptors have been proposed for ketamine neurotoxicity.7,8 Nevertheless, ketamine has also been shown to act independently from the NMDA receptor exerting toxicity at various types of cell and tissues.9-11 Teratological assays typically conducted in mammalian models, such as rodents and rabbits,12 have shown the transmission of effects from the mother to the embryos.13 However, these studies are generally expensive, time-consuming and require a large number of animals.14 Moreover, these experiments raise concerns regarding animal welfare, protection, and ethics. Currently, these concerns have been overcome by using early life stages of zebrafish (up to 4-5 days post-fertilization), according to the European Union Directive 2010/63/EU.14 In fact, the use of zebrafish embryonic stages in toxicity screening is growing among different areas of research, as it offers a vertebrate model with a high degree of similarity to other high-order vertebrates regarding anatomic, cellular, physiological and genetic processes.15,16 Additionally, the embryo development is a highly coordinated process,17 requiring specific and time-dependent cellular interactions, that can be disrupted by environmental conditions resulting in abnormal development.16 Thus, the use of zebrafish early life stages is of particular interest to study insults during embryogenesis. In adult zebrafish, ketamine has been shown to induce behavioral dose-dependent effects similar to those observed in higher vertebrates.18-20 Additionally, NMDA-related motor neuron toxicity, neurogenesis genetic expression changes, as well as cardiac toxicity during early development has been reported.21,22 In zebrafish, embryos exposed to ketamine during 5 ACS Paragon Plus Environment


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256-cell, a stage where the NMDA receptors are still not expressed,23 we have reported the occurrence of non NMDA-related teratological effects, including the occurrence of malformations that impair and question the occurrence of malformations that decrease survival.9 In the current work, we tested the hypothesis that ketamine exposure during more advanced embryonic stages, also prior to NMDA receptors expression, would display a similar toxicological profile to the observed when embryos were exposed during the 256-cell stage. Thus, the aim of this study was to evaluate ketamine effects on skeletal development after exposure during the 256-cell, 50% epiboly and 1-4 somites stages, and to evaluate potential ketamine-induced deformities during the 50% epiboly and 1-4 somites stages. Moreover, gene expression relative levels of developmental-related genes were evaluated as a tool to identify key events that preceded morphological toxicity.


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2. Experimental Procedures 2.1 Chemicals and Other Reagents Pharmaceutical-grade ketamine hydrochloride (Imalgene1000, 100 mg mL− 1) was obtained from Merial Portuguesa-Saúde Animal Lda (Rio de Mouro, Portugal). All solutions were freshly made with embryo water (28 ± 0.5 °C, 200 mg L− 1 Instant Ocean Salt and 100 mg L−

1

sodium bicarbonate; UV sterilized) prepared from City of Vila Real filtered tap water.

Instant Ocean Salt was obtained from Aquarium Systems Inc. (Sarrebourg, France). All other chemical reagents were either of analytical or HPLC grade and acquired from Sigma-Aldrich (Steinheim, Germany). To establish a standard curve for HPLC analysis, ketamine hydrochloride was purchased from Biotrend (Cologne, Germany).

2.2 Animals and treatments Zebrafish maintenance and embryo collection were performed has previously described.9, 24, 25 Briefly, adult wild-type (AB strain) zebrafish were maintained at the University of Trás-os-Montes and Alto Douro (Vila Real, Portugal) in an open water system supplied with aerated, dechlorinated, charcoal-filtered and UV-sterilized City of Vila Real tap water (pH 7.3– 7.5) at 28 ± 0.5 °C in a 14:10 h light: dark cycle. The fish were fed twice a day with a commercial diet (Sera, Heinsberg, Germany) supplemented with Artemia sp. nauplii. Embryos were obtained by random mating of adult zebrafish, that were collected between 1 and 2 hours post-fertilization (hpf), bleached and rinsed to remove debris, and randomly distributed among treatments. Figure 1 shows a schematic diagram of the experimental design. Exposures of 20 minutes to ketamine concentrations (0.2, 0.4 and 0.8 mg mL−1, respectively 0.84, 1.68 and 3.37 mM) were performed during early 256-cell (2.5 hpf), early 50% epiboly (5.5 hpf) and 1-4 somites (10.5 hpf).9,

17

Test concentrations were selected based on previous work with

ketamine in adult zebrafish where 0.2 mg mL-1 was defined as subthreshold and 0.8 mg mL-1 as above threshold for an anesthetic effect in this species.19 An additional group was maintained 7 ACS Paragon Plus Environment


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without ketamine and used as the control group. After exposure, the embryos were washed three times in embryo water and allowed to develop up to 144 hpf, with daily water renewal. For each treatment at least six independent replicates containing 100 viable embryos were performed to evaluate mortality, lethal and non-lethal parameters. Additional experiments with three independent replicates of 300 viable embryos were used for gene expression studies. HPLC ketamine quantification, alcian blue, and calcein assessments were performed exposing 100 embryos in triplicate. Throughout all procedures, the temperature was kept at 28 ± 0.5 °C. All experimental procedures were conducted in compliance with licenses approved by the National Institutional Animal Care Committee (Direção-Geral de Alimentação e Veterinária, Lisboa, Portugal) and in agreement with European (Directive 2010/63/EU) and Portuguese (Directive 113/2013) legislations on animal welfare ensuring minimal animal distress.

2.3 Analysis of internal ketamine accumulation by HPLC-UV The accumulation of ketamine in embryos was analyzed immediately after exposure (T0) and 24 hours after exposure (T24). Samples were collected as previously described 26 with slight modifications.27,28 Briefly, at each time points (T0 and T24), surviving animals were collected to microcentrifuge tubes, washed three times with ultrapure water and alkalinized with 350 µL 0.2 M borate buffer (pH 13). Samples were then homogenized on TissueLyser II (Qiagen, Hilden, Germany), at 30 Hz for 90 seconds, before a double extraction with 500 µL dichloromethane:ethyl acetate (80:20 v/v). After centrifugation at 1500 xg for 3 minutes at room temperature, the combined organic phase was back-extracted with 2 M HCl and the acidic phase evaporated to dryness at 45 °C in a centrifugal evaporator (Univapo 150ECH; UniEquip, Munich, Germany). The dry residue was reconstituted in 100 µL of the mobile phase which consisted of acetonitrile: 30 mM phosphate buffer (23:77 v/v) at pH 7.2. The chromatographic separation (50 µL) was performed with a Hichrom ACE Ultracore 5 Super C18 (150 × 4.6 mm, 5 µm) at 20 °C and a flow rate of 1.5 mL min −1 in a Dionex UltiMate 3000 HPLC 8 ACS Paragon Plus Environment

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system (Dionex, Olten, Switzerland) coupled with a diode array detector (Dionex PDA 100 photodiode array, Dionex, Olten, Switzerland) at 210 nm and controlled by the Dionex Chromeleon Software 6.70 Build 1820. Control samples were used as blanks and ketamine was quantified by measuring the area under the peak, which was drawn from the analysis of seven known ketamine concentrations (from 0.05 to 10 pmol). The integrated peak areas were plotted in function of the concentration of the standard solutions and from the equation of the linear correlation plot, the amount (pmol) of ketamine in each sample was interpolated. The amount absorbed by each embryo was calculated analytically dividing the amount of ketamine detected by the number of surviving animals collected.

2.4 Zebrafish embryotoxicity test The developmental toxicity of ketamine was evaluated for each phase based on the observable morphological parameters reported previously.9 Briefly, all embryos/larvae were inspected daily and both lethal and sub-lethal effects recorded post exposure at four time points (24, 48, 72 and 144 hpf). At each time point, twelve animals were randomly removed from each replicate and visually inspected in a color digital CCD camera (Color View III, Olympus, Hamburg, Germany) mounted on an inverted microscope (IX 51, Olympus, Antwerp, Belgium) using a 4X Olympus UIS-2 objective lens (Olympus Co., Ltd., Tokyo, Japan) and data acquired using Cell R software (Olympus, Antwerp, Belgium). All scores were performed by a single observer blinded to the treatments. The following parameters were evaluated as categorical measurements: egg coagulation, somite formation, detachment of the tail-bud from the yolk sac, head development including otoliths and eye formation, body pigmentation, pericardial edema and hatching rate. The circulatory system was also analyzed during this period. After hatching, larvae were immobilized in 3% methylcellulose

29

and assayed for

skeletal deformities and standard body size using digital image analysis (Digimizer version 4.1.1.0, MedCalc Software, Mariakerke, Belgium). At 144 hpf, serial zebrafish images were 9 ACS Paragon Plus Environment


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combined, merged and processed with Adobe Photoshop CS6 (Adobe Systems, San Jose, USA) and blind-scored for anomalies using a rating scale (0 to 4 according to increasing severity).9,30 Dead embryos/larvae were removed daily and not taken into consideration for analysis.

2.5 Development of cartilage and bones Zebrafish cartilage evaluation was carried out with alcian blue staining.31 Briefly, at 144 hpf, animals were euthanized with an overdose of tricaine and fixed overnight in 4 % paraformaldehyde. Afterwards, the larvae were washed and dehydrated in 50 % ethanol at room temperature for 10 minutes and stained with 0.02% alcian blue in 70% ethanol with 60 mM MgCl2 overnight. Bleaching (3 % H2O2 and 2 % KOH) was performed to remove excess pigmentation and larvae soft tissues were digested in 0.05% trypsin in PBS.32 Stained animals were then gradually cleared through a series of glycerol/KOH (20%/0.25%; 50%/0.25% and 80%/0.1% for storage). Likewise, at 144 hpf, changes in bone shape were analyzed in vivo using calcein.33 Briefly, twelve larvae were immersed in a 0.2 % solution (pH 7.2 with NaOH) for 10 minutes in the dark and at room temperature. At the end of the staining period, animals were rinsed three times with embryo water, immobilized in 3% methylcellulose and observed. Neurocranial (ethmoid plate, basicranial commissure, trabecula cranii, basal plate and occipital arch) and pharyngeal cartilages (basihyal, ceratobranchial arches , ceratohyal, hyosympletic, interhyal, Meckel’s, palatoquadrate, basibranchial and hypobranchial) as well as vertebrae, cephalic dermal bones (parasphenoid, branchiostegal rays, operculum, entopterygoid, dentary and cleithrum), cartilage-replacement bone (CB5 ossification), occipital and notochord bones were observed under light and fluorescent microscopy. As previously stated, samples were coded and score blinded in a semi-quantitative approach (0 to 3 according to increasing malformations). The head length was measured 34 by digital image analysis (Digimizer version 4.1.1.0, MedCalc Software, Mariakerke, Belgium).


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2.6 Geometric morphometrics At 144 hpf, geometric morphometrics was used to quantify similarities and differences between head shapes of experimental animals. In brief, seven lateral-view landmarks

35

and

twenty-one ventral-view landmarks were used to define the cartilaginous morphology whereas twenty-nine ventral landmarks were defined to study bone morphology of the embryonic head (Fig S1).36 Two-dimensional (x,y) coordinates were manually digitized in ImageJ 1.50e software with Point Picker plugin.37 Raw coordinates were Procrustes-transformed in order to eliminate differences in size, translation, and rotation between the samples.38 To remove the size effects due to allometric growth, a pooled within-group regression on log centroid size was performed and the residuals from this regression were the basis for further analysis. To visualize the position of morphotypes in environmental space, a scatter plot of principal component analysis (PCA) score values was used. All the analyses were made in MorphoJ 1.06d.39

2.7 Developmental-related gene expression Quantitative real time PCR (qRT-PCR) has become the most suitable method to study gene expression in these early stages dues to its sensitivity and specificity that is lost after 2 days of development when using other molecular methods.40 Thus a quantification of the relative expression levels were used to study ketamine influence in gene expression. Embryos/larvae were collected at four time points (8, 24, 72 and 144 hpf) and used for RNA extraction. Total RNA was extracted from independently collected pools of embryos/larvae and synthesized into cDNA by reverse transcription as described before,

24,25

with minor

modifications. Briefly, at each time point, pools of 50 viable embryos were randomly collected from each treatment and stored in RNAlater (Sigma, Steiheim, Germany). Samples were then homogenized on TissueLyser II (Qiagen, Hilden, Germany), at 30 Hz for 90 seconds, before RNA extraction using the Illustra RNAspin kit (GE Healthcare, Munich, Germany) according to the manufacturer's instructions. Treatment with DNase I (RNase-free) was extended for up to 1 11 ACS Paragon Plus Environment


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hour. Total RNA was recovered in 40 µL of RNAse free water and quantified by spectrophotometric (Powerwave XS2, BioTek Instruments, Inc. USA) readings at 260 and 280 nm. RNA integrity and DNA contamination were evaluated by 1 % agarose gel electrophoresis, using Green Safe Premium staining (NZYTech, Ltd., Lisbon, Portugal) and detection under UV light using BioCapt software (v99.02, Vilber Lourmat, France). RNA samples were stored at -80 °C until further use. RNA (500 ng) from each sample were reverse transcribed using the iScript cDNA synthesis (Bio-Rad Laboratories, California, USA) following the manufacturer's instructions and stored at -20 °C. The quantitative real-time PCR (qRT-PCR) reactions were performed in a final volume of 20 µL containing 1 µL of cDNA and 200 nM of each specific primer (Table 1), and using the 5x HOT FIREPol EvaGreen qRT-PCR Mix Plus (Solis Biodyne, Tartu, Estonia) in a Stratagene Mx3005P Real-Time PCR system (Stratagene, Agilent Technologies, Santa Clara, USA). Every set of qRT-PCR was carried out in three independent replicated samples performed in duplicate, including a reaction control without cDNA. The following thermal cycling conditions were used: initial denaturation at 95 °C for 10 minutes followed by 40 cycles of denaturation at 95 °C for 20 seconds, primer annealing for 40 seconds (annealing temperatures in Table 1) and extension at 72 °C for 20 seconds followed by a final extension at 72 °C for 5 minutes. The amplification reaction was followed by a melting curve analysis to exclude the presence of primer-dimers, DNA contaminants and other nonspecific products. The relative expression levels were analyzed using the MxPro QRT-PCR System software (Stratagene, Agilent Technologies, Santa Clara, USA) and were determined by normalization to the β-actin gene used as housekeeping. Values were then normalized to the control average value calculated using the ΔΔCt method with efficiency correction 41 based on standard curves of a 5-fold dilution series prepared from a mix of the used samples.

2.8 Statistical analysis


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Sample size calculation was performed with the G*Power 3 (University of Düsseldorf, Germany) as described in our previous work.9 Normal distribution and homogeneity of the data were confirmed by Kolmogorov–Smirnov and Levene's tests, respectively. When data had no normal distribution, groups were compared by a non-parametric independent samples test, Kruskal–Wallis, followed by Dunn's pairwise comparison tests and data was expressed as median and interquartile range (25th; 75th percentiles). Data normally distributed was analyzed by one-way analysis of variance (ANOVA) followed by Tukey's pairwise comparison tests and data was expressed as mean ± standard deviation. Data following both parametric and non-parametric distributions (gene expression), non-parametric tests were used as they are more conservative and best suited for small datasets. Categorical data, expressed as a percentage, were log transformed (log (x+1)) to meet ANOVA assumptions of normality and homogeneity of variance when necessary and then presented as back-transformed mean of the log transformed variable and upper and lower limits from the 95% confidence interval. Non-detected values were assigned a zero value for statistical purposes. All tests for all analyses described in this section were two-tailed. In all cases, statistical analyses were carried out using SPSS for Windows (Version 22.0; Chicago, IL, USA) and differences were considered significant at p

Embryonic Stage-Dependent Teratogenicity of Ketamine in Zebrafish

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