Neurotoxicology and Teratology 41 (2014) 27–34

Contents lists available at ScienceDirect

Neurotoxicology and Teratology journal homepage: www.elsevier.com/locate/neutera

Ketamine NMDA receptor-independent toxicity during zebrafish (Danio rerio) embryonic development Luís M. Félix a,b,⁎, Luís M. Antunes b,c, Ana M. Coimbra d,e a

Life Sciences and Environment School, University of Trás-os-Montes and Alto Douro, Apartado 1013, 5001-801 Vila Real, Portugal Laboratory Animal Science, Institute for Molecular and Cell Biology, University of Porto, Rua do Campo Alegre, 823, 4150-180 Porto, Portugal Department of Veterinary Sciences, School of Agriculture and Veterinary Sciences, University of Trás-os-Montes and Alto Douro, Apartado 1013, 5001-801 Vila Real, Portugal d Centre for the Research and Technology of Agro-Environmental and Biological Sciences, Life Sciences and Environment School, University of Trás-os-Montes and Alto Douro, Apartado 1013, 5001-801 Vila Real, Portugal e Department of Environmental and Biological Engineering, Life Sciences and Environment School, University of Trás-os-Montes and Alto Douro, Apartado 1013, 5001-801 Vila Real, Portugal b c

a r t i c l e

i n f o

Article history: Received 15 July 2013 Received in revised form 12 November 2013 Accepted 17 November 2013 Available online 26 November 2013 Keywords: Anaesthesia Development Ketamine Teratology Toxicity Zebrafish

a b s t r a c t Concerns have been raised that the effect of anaesthetic drugs on the central nervous system may result in longterm impairment, namely when ketamine is used during embryogenesis. In addition, the cell and molecular basis of anaesthetics teratology and toxicity are still uncertain and its implications in the development remain to be clarified. More recently, the potential risks for human, and animal, exposure through environmental contamination also became an important question. In this study, the effects of sub- and over anaesthetic doses of ketamine were investigated during zebrafish (Danio rerio) embryonic development by exposing zebrafish embryos to ketamine concentrations (0.2, 0.4 and 0.8 mg mL−1) for a period of 20 min during the blastula stage. Ethanol 2% was used as a positive control. Morphological parameters, the overall pattern of cell death using acridine orange and overall degree of oxidative stress levels by 2,7-dichlorodihydrofluorescein-diacetate were determined. Lethality and/or developmental anomalies were measured based on specific time endpoints until 144 h post fertilisation. Results showed a concentration-dependent increase in anomalies and mortality. Cephalic disorders, enlarged organs and tail/spine anomalies were the most prominent deformities observed at 144 hpf. Acridine orange images revealed no differences in cellular death pattern in exposed embryos at 24 hpf. At the same time point, the cellular redox processes were found to be similar among groups. In summary, this study shows that ketamine is teratogen and toxic, interfering with the normal developmental pathways of embryogenesis, suggesting that ketamine exerts an independent NMDA receptor action during the zebrafish blastula stage. © 2013 Elsevier Inc. All rights reserved.

1. Introduction Over the years, anaesthetics have been used in paediatric and obstetric patients without clinical evidence of their toxic effects to the central nervous system (Mellon et al., 2007). Ketamine, a non-competitive inhibitor of N-methyl-D-aspartate (NMDA) glutaminergic receptors (Li et al., 2011), is a chemical commonly used for induction and maintenance of anaesthesia and sedation in human and veterinary medicine (de la Torre, 2010), as well as a drug of abuse (Li et al., 2011). Moreover, ketamine has been detected in surface waters (Lin et al., 2010; Baker and Kasprzyk-Hordern, 2011; Vazquez-Roig et al., 2012). The effects of ketamine exposure during gestation are not clear and experimental studies are needed in order to clarify these. A recent review highlighted the neurotoxicity of ketamine to newborns (Dong and Anand, 2013) and, to our knowledge, only one study has described ⁎ Corresponding author at: Life Sciences and Environment School, University of Trás-osMontes and Alto Douro, Apartado 1013, 5001-801 Vila Real, Portugal. Tel.: +351 259 350 000; fax: +351 259 350 480. E-mail address: [email protected] (L.M. Félix). 0892-0362/$ – see front matter © 2013 Elsevier Inc. All rights reserved. http://dx.doi.org/10.1016/j.ntt.2013.11.005

the occurrence of clinical features in a neonate whose mother was a ketamine drug abuser (Su et al., 2010). Furthermore, studies in monkey proposed the association between high ketamine doses and neuroapoptosis in the foetal brain (Paule et al., 2011; Brambrink et al., 2012). Studies with rat are controversial as some described no ketamine teratological effects (El-Karim and Benny, 1976; AbdelRahman and Ismail, 2000), while others presented dose-dependent effects (Kochhar et al., 1986; Bandazhevskii and Shimanovich, 1991). Despite these findings, the transposition of animal results to humans is unknown and additional studies are currently being performed using cohort studies and different animal models in order to identify possible risks (Bosnjak, 2012). Different aquatic species have been used as models in several lines of human biomedical research (Riehl et al., 2011). Zebrafish embryos emerged as one of such models since embryonic stages of fish are potentially useful as an alternative to mammalian assays and whole embryo culture (Fraysse et al., 2006). Additionally, zebrafish are being increasingly used for the study of vertebrate development (Fleming and Cambridge, 2007) and as a preclinical model for drug screening (Parng et al., 2002) including for ketamine toxicity (Kanungo et al., 2012,

28

L.M. Félix et al. / Neurotoxicology and Teratology 41 (2014) 27–34

2013). Furthermore, 69% of zebrafish genes possess at least one human orthologue, and from these and regarding human disease-related genes 82% have a zebrafish orthologue (Howe et al., 2013), supporting the proximity of zebrafish with higher vertebrates. The first study showing the effects of ketamine in zebrafish larvae dates back to 2007. This showed that ketamine impairs prepulse inhibition (Burgess and Granato, 2007). Recent studies also described ketamine-induced behavioural dose-dependent effects in adult zebrafish (Riehl et al., 2011; Zakhary et al., 2011) and motor neuron toxicity in a transgenic zebrafish embryo line (Kanungo et al., 2013). Kanungo et al. (2012) also showed that 2-hour exposure to ketamine has a dose-dependent effect on the heart rate of 26 and 52 h post fertilisation (hpf) zebrafish dechorionated embryos. Zebrafish NMDA receptor expression was identified at 24 hpf (Cox et al., 2005), but fish can be exposed as early as the zygote stage to aquatic contaminants. Thus, the objective of the present study was to identify the direct effects of sub- and over anaesthetic doses of a ketamine exposure during the early blastula stage, prior to NMDA receptor expression, in zebrafish embryos development. Additionally, as fish and higher vertebrates, including humans, share a high genetic and organ system homology, this study adds knowledge on ketamine toxicity during embryogenesis. 2. Material and methods 2.1. Chemicals Ketamine (Imalgene1000, 100 mg mL−1) was obtained from Merial Portuguesa-Saúde Animal Lda (Rio de Mouro, Portugal). Absolute ethanol was purchased from Panreac (Panreac, Spain). All other chemical reagents were obtained from Sigma-Aldrich (Steinheim, Germany). All solutions were prepared daily with system water (28.5 °C, 200 mg L− 1 Instant Ocean Salt and 100 mg L − 1 sodium bicarbonate; UV sterilised).

were made to ensure minimal animal stress and discomfort. Zebrafish maintenance and embryo collection were performed according to standard procedures (Soares et al., 2009). Briefly, wild-type (AB strain) zebrafish were maintained at 28 ± 0.5 °C, in a 14:10 h light:dark cycle, in a semi-closed water system with both mechanical and biological filtration. The fish were fed twice a day with a commercial diet (Sera, Heinsberg, Germany) supplemented with Artemia sp. nauplii. Zebrafish embryos were obtained from spawning adults grouped in tanks overnight. Spawning was induced in the morning with the beginning of the light period. Newly-fertilised eggs were collected and rinsed several times in system water before being randomly distributed.

2.3. Toxicant and exposure procedure During the early blastula period (Kimmel et al., 1995), embryos were examined and those that were normally developed were selected for the exposure procedure. Fig. 1 shows an overview of the experimental design. Briefly, 100 embryos per group were randomly distributed by beakers containing 50 mL of exposure solutions: negative (system water) and positive (2% ethanol) controls and ketamine concentrations (0.2, 0.4 and 0.8 mg mL−1, respectively 0.84, 1.68 and 3.37 mM). The range of ketamine concentrations was selected based on our pilot studies and on a 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 anaesthetic effect in this species (Zakhary et al., 2011). Embryos were exposed to ketamine for 20 min and to ethanol for 21 h based on previous works where ketamine and ethanol effects were reported in zebrafish (Reimers et al., 2004; Riehl et al., 2011). Five to six independent replicates for each concentration were run, with two beakers per group: one to be used in toxicological endpoint evaluation and another for staining methods. At the end of the exposure periods, embryos were washed three times with system water and allowed to develop until 144 h post-fertilisation (hpf) with daily water changes.

2.2. Maintenance of zebrafish and embryo collection All procedures used were carried out in agreement with European and Portuguese legislations on animal welfare, under personal and project licences approved by the National Institutional Animal Care Committee (Direcção Geral de Veterinária, Lisboa, Portugal). All efforts

2.4. Mortality rate Zebrafish embryo/larvae mortality rate was recorded daily. Embryos were recorded as dead when no movement was detected, if there was a

Fig. 1. Experimental design overview. Between 2 and 3 h post-fertilisation, 100 embryos per group were randomly distributed by beakers containing 50 mL of exposure solutions: negative (system water) and positive (2% ethanol) controls and ketamine solutions (0.2, 0.4 and 0.8 mg mL−1). After 20 min of ketamine and 21 h of ethanol exposure, embryos were washed using system water and set to grow in clean water. Water from the beakers was replaced daily. Randomly, 12 larvae/embryos were microscopically assessed for tail detachment, somite formation, head development, circulatory system (including blood circulation and heartbeat), pigmentation, hatching, lethality, cellular death and reactive oxygen species at 24, 48, 72 and 144 h post-fertilisation. Embryos were scored for malformations (fins, heart/thorax, head, eyes, organs and tail/spine) at 144 h post fertilisation by visual inspection. Scale bar represents 500 μm.

L.M. Félix et al. / Neurotoxicology and Teratology 41 (2014) 27–34

lack of heart beating or if the tissues changed from a transparent to an opaque appearance. Dead animals were removed from the beakers. 2.5. Toxicological endpoint evaluation The development of zebrafish embryos/larvae was monitored at specific time points throughout gastrulation, organogenesis and early larval development (Selderslaghs et al., 2009). At each time point, twelve zebrafish were randomly removed from each beaker and analysed by visual inspection in an inverted microscope (IX 51, Olympus, Antwerp, Belgium) by an investigator who was blinded to treatments. The toxicological endpoints used to assign developmental effects of ketamine included tail detachment, head development including eyes and otoliths, somite formation, circulatory system comprising heart rate and blood circulation, pigmentation, hatching success, oedema, malformations and length of the larvae. Tail detachment, somite formation and head development were analysed daily for presence/absence (0/1). Circulatory system and pigmentation were also assessed for presence/absence (0/1) at 48, 72 and 144 hpf. Embryo/larvae were immobilized in 3% methylcellulose (Muntean et al., 2010) and the heart rate was recorded during 15 s at 48, 72 and 144 hpf and expressed as beats per minute. During this period, embryos were analysed for the presence of oedema. Zebrafish from each replicate were also examined for hatching success and skeletal deformities at 72 and 144 hpf. At the end of the experiment, larvae were also photographed with a colour 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). Serial images were combined, merged and processed with Adobe Photoshop CS6 (Adobe Systems, San Jose, USA). The standard body length of larvae was measured using a digital image analysis software (Digimizer version 4.1.1.0, MedCalc Software, Mariakerke, Belgium). 2.6. Anomalies score At 144 hpf, a rating scale (Padilla et al., 2011) was applied to score zebrafish anomalies. Briefly, each larva was assessed for spine, fins, cranial/facial, thorax and abdominal anomalies by an investigator who was blinded to treatments. Each anatomical feature anomaly was scored from 0 to 4, accordingly to the severity. The total anomaly score was obtained by the sum of the score and by calculating the mean of the individual category scores, with higher scores representing more severely deformed animals. 2.7. Cellular death pattern Acridine orange (AO) staining was used to identify apoptosis in live embryos/larvae (Tucker and Lardelli, 2007). Briefly, at 24, 48, 72 and

29

144 hpf, twelve embryos/larvae were incubated 30 min, in the dark, in a solution of 5 μg mL−1 of acridine orange. Before observation, larvae were rinsed three times in system water. Embryos/larvae were imaged with a colour digital CCD camera (Color View III, Olympus, Hamburg, Germany) through a Fluorescein-Isothiocyanate (FITC) filter using a fluorescence microscope (IX 51, Olympus, Antwerp, Belgium) with a 4X Olympus UIS-2 objective lens (Olympus Co., Ltd., Tokyo, Japan) and visualized within 1 min of exposure as the signal is quenched after that period (Rohde and Heisenberg, 2007). Images were then compiled and assembled in Adobe Photoshop CS6 (Adobe Systems, San Jose, USA) for further analysis. 2.8. Reactive oxygen species generation Estimation of reactive oxygen species accumulation in zebrafish embryos/larvae was analysed using 2,7-dichlorodihydrofluoresceindiacetate (DCFH-DA) (Ko et al., 2011). Briefly, at 24, 48, 72 and 144 hpf, twelve embryos/larvae were incubated 30 min, in the dark, in a solution of 20 μg mL−1 of DCFH-DA. After the incubation period, and before observation, larvae were rinsed three times in system water. Embryos/ larvae were imaged as described previously for the cellular death assay. 2.9. Statistical analysis For statistical purposes, each beaker was considered an experimental unit, thus when twelve embryos were taken from the beaker, their values were averaged and considered as n = 1. Sample size calculation, assuming type II error probability of α = 0.05 and power as 0.90, was based upon the results of our previous study (Soares et al., 2009), where 4 replicates were sufficient to detect effects between negative control and treatment groups. In this study, 5 to 6 replicates were used to decrease variability in the results. Data were checked for normality (Shapiro–Wilk test) and homogeneity of variance (Levene's test). When the assumptions of normality and homogeneity of variances were met, differences among groups were assessed by analysis of variance (ANOVA) followed by the Tukey multiple comparison test and data expressed as mean ± standard deviation. Categorical variables (presence or absence of characteristics 0/1) were expressed as percentages and log transformed (log (x + 1)) before analysis by parametric statistics if normality was passed. Data was then quoted as back transformed mean value of the log transformed variable and upper and lower limits from the 95% confidence interval. Whenever normality was not passed, the Kruskal–Wallis analysis of variance, followed by Dunn's test with a Bonferroni correction for multiple comparisons was used and data expressed as medians and interquartile range (25th; 75th percentiles). Correlation coefficients were calculated using the Pearson test. A chi-square test was performed to analyse categorical variables. The significance level was set for p b 0.05. Power analysis was done

Fig. 2. Dose-response and time course mortality of zebrafish exposed to ketamine and ethanol. Mortality was recorded at 8, 24, 48, 72 and 144 h post-fertilisation. Data are represented as mean ± SD of six independent experiments. Uppercase letters indicate differences between groups within the same time and lowercase letters indicate differences between time within the same group (two-way ANOVA, p b 0.05).

30

L.M. Félix et al. / Neurotoxicology and Teratology 41 (2014) 27–34

Table 1 Frequency (%) of morphological characteristics identified in surviving zebrafish (D. rerio) embryos at 24 and 48 hpf. Group

Neg. control Ketamine

Pos. control Statistical test p

Dose (mg mL−1)

24 hpf No tail detached

No somite formed

No head developed

48 hpf Oedema presence (yolk sac and cardiac)

Heartbeat and blood circulation visible

No eye developed

No otoliths developed

0.0 0.2 0.4 0.8 EtOH 2%

0.00 (0.00–0.00)a 0.56 (0.00–4.40)a 1.44 (0.00–10.2)a 8.33 (8.33–8.33)b 2.06 (0.00–10.0)a F(4,25) = 3.98 0.015

0.00 (0.00–0.00) 0.00 (0.00–0.00) 0.45 (0.00–2.78) 1.11 (0.00–6.06) 0.00 (0.00–0.00) F(4,25) = 1.54 0.22

0.00 (0.00/0.00) 0.61 (0.00–4.52) 1.11 (0.00–6.06) 1.11 (0.00–6.06) 2.06 (0.00–10.0) F(4,25) = 0.93 0.47

0.00 (0.00–0.00)a 0.45 (0.00–2.78)a 1.44 (0.00–10.2)a 10.5 (0.00–27.9)b 72.6 (59.8–88.1)c F(4,25) = 36.04 b0.0001

0.00 (0.00–0.00)a 0.00 (0.00–0.00)a 0.00 (0.00–0.00)a 0.00 (0.00–0.00)a 67.8 (62.5–74.8)b F(4,25) = 12619 b0.0001

0.00 (0.00–0.00) 0.00 (0.00–0.00) 0.00 (0.00–0.00) 0.00 (0.00–0.00) 0.00 (0.00–0.00) np np

0.00 (0.00–0.00) 0.00 (0.00–0.00) 0.00 (0.00–0.00) 0.00 (0.00–0.00) 0.00 (0.00–0.00) np np

Data are quoted as back transformed mean value of the log transformed variable and upper lower limits from the 95% confidence interval. Statistical analysis, in 6 independent replicates of 12 animals each, was performed using one-way ANOVA followed by Tukey's multiple-comparison test. Different lowercase letters indicate significant differences between groups (p b 0.05). Abbreviations used: (np) not possible to calculate descriptive statistics.

using G*Power 3.1.5 (Heinrich Heine University, Dusseldorf, Germany). Statistical analysis was performed using the statistical package SPSS 20.0 for Windows (SPSS Inc., Chicago, USA) including all studied groups. 3. Results 3.1. Dose-dependent increase in mortality with ketamine The mortality of zebrafish embryos and larvae was recorded at five time points (8, 24, 48, 72 and 144 hpf). Zebrafish embryo/larvae mortality increased in all groups between 8 hpf and 48 hpf, remaining thereafter constant in almost all groups (Fig. 2). The statistical analysis showed a significant ketamine concentration [F(4,125) = 136.1, p b 0.001] and time [F(4,125) = 29.79, p b 0.001] effect, but no correlation was detected between both [F(16,125) = 0.71, p = 0.778]. The negative control group showed the lowest mortality values at all times, ranging from 11.5 ± 0.50% at 8 hpf and 16.0 ± 2.28% at 24 hpf (p N 0.05). The group exposed to 0.2 mg mL−1 presented mortality rates similar to the negative control group at 8 hpf and 24 hpf and significant higher values subsequently. Zebrafish embryo exposed to 0.4 mg mL− 1 displayed at all times significantly higher values when compared to the negative control, showing along the experiment, identical values to the positive control (ethanol 2%). The group exposed to 0.8 mg mL−1 of ketamine showed always the highest mortality values, ranging from 26.8 ± 2.01% at 8 hpf to 35.0 ± 2.61% at 48 hpf.

26% of ethanol exposed embryos (Fig. 3A). By 72 hpf, the hatching rates (Table 2) among surviving embryos were not statistically different between ketamine groups and negative control group despite a decreasing tendency with increasing ketamine dose. Compared to the negative control, ethanol exposure resulted in a statistically significant decrease of the hatching success (p b 0.001). Zebrafish larvae skeletal deformities were analysed at 72 and 144 hpf. The cumulative outcome is shown in Fig. 3B. The statistical analysis showed significant effect of dose [F(4,50) = 44.04, p b 0.001] but no statistical difference between time points [F(1,50) = 0.85, p = 0.362], or an interaction between them, F(4,50) = 0.51, p = 0.727. The incidence of skeletal deformities at 72 and 144 hpf in the negative control was the lowest observed. At 72 hpf, an increase with the ketamine doses was observed. The 0.8 mg mL− 1 group showed more skeletal deformities; however, this was not significantly different from the other groups. At 144 hpf, the rate of skeletal deformities among groups showed the same pattern. However, group 0.8 mg mL−1 (p = 0.004) showed significantly more skeletal deformities. The positive control (ethanol 2%) showed, at both time points, the highest incidence (72 hpf p b 0.001; 144 hpf p b 0.001) when compared to the negative control. The Pearson correlation coefficient between exposure groups and the skeletal deformities observed, at 72 hpf and 144 hpf, were 0.79 (p b 0.001) to 0.89 (p b 0.001), respectively. At the end of the experiment, by 144 hpf, it was observed that the total body length was only affected by the ethanol exposure (p = 0.002) as no statistical differences were observed for ketamine groups comparatively to the negative control group.

3.2. Ketamine-induced developmental toxicity General outcomes from toxicological endpoint analyses of embryos/ larvae are summarised in Table 1. At 24 hpf, somite formation and head development in the exposed embryos were similar between the negative control group and all tested groups. Tail detachment was the only parameter significantly affected by the ketamine exposure causing a significant lack of detachment of the tailbud from the yolk sac (data log transformed, p = 0.008). No differences were observed between control and ethanol animals. At 48 hpf, the occurrence of yolk sac and cardiac oedema was significantly higher for 0.8 mg mL− 1 ketamine (data log transformed, p = 0.0007) and ethanol (data log transformed, p b 0.0001) animals when compared to control. No significant effects were observed in ketamine groups and ethanol group regarding eye development and otoliths presence. Detection of blood circulation and heart-beat in the ethanol group, at 48 hpf, could only be observed in one third of the embryos (p b 0.001). At this time, ketamine groups did not show alterations of the cardiovascular system (Table 2). Pigmentation was not different between groups, however weak, dispersed and abnormal chromatophore morphology was observed in 6% of the animals exposed to the highest ketamine concentration and in

Table 2 Quantitative analysis of zebrafish embryos/larvae developmental parameters at 48, 72 and 144 hpf. Group

Neg. control Ketamine

Pos. control Statistical test p

Dose (mg mL−1)

48 hpf

72 hpf

144 hpf

Heart rate (bpm)1

Hatching rate (%)

Body length (cm)

0.0 0.2 0.4 0.8 EtOH 2%

161 ± 17 157 ± 18 157 ± 19 152 ± 14 142 ± 31 F(4,25) = 0.732 0.579

94 ± 2a 91 ± 3a 91 ± 1a 83 ± 3a 65 ± 8b F(4,25) = 7.631 b0.001

4.38 ± 0.14a 4.30 ± 0.13a 4.23 ± 0.05a 4.25 ± 0.13a 3.71 ± 0.07b F(4,25) = 5.882 0.002

Data presented as mean ± SD of six independent replicates. Statistical analysis was performed using one-way ANOVA followed by Tukey's multiple-comparison test. Different lowercase letters indicate significant differences between groups (p b 0.05). Abbreviations used: (bpm) beats per minute. 1 Mean of individuals that presented heartbeat.

L.M. Félix et al. / Neurotoxicology and Teratology 41 (2014) 27–34

31

Fig. 3. A. Malformation in zebrafish exposed to ketamine and ethanol. Anomalies observed were evident in the higher ketamine dose and even more extensive in the ethanol exposed larvae. The major anomalies (arrowheads) observed were: cardiac enlargement (ce); brachycephaly (b); undershot jaw (j); microcephaly (mc); microphthalmia (mp); enlarged organs (eo) and lordosis (l) or kyphosis (k). Note that the melanocytes (m) on these groups present a dispersed and abnormal morphology forming a pavemented layer along the body. Scale bar represents 500 μm. B. Cumulative tail and spine skeletal deformities of zebrafish at 72 and 144 h post-fertilisation. Data are represented as mean ± SD of six independent experiments. Uppercase letters indicate differences between groups within the same time. No statistical differences were observed between time within the same group (two-way ANOVA, p b 0.05).

3.3. Ketamine associated with developmental anomalies General outcomes of morphological analyses of 144 hpf larvae are summarised in Table 3. Main anomalies that were observed after the ketamine exposure are illustrated in Fig. 3. By 144 hpf, the control group appeared normal with few observed anomalies which included malformed fins, cardiac enlargement and microphthalmia. Ketamine exposure did not significantly increase the observed malformations; however, it was observed that it had a tendency to increase a wide range of malformations. These were prevalent in the highest dose and included malformed fins, cardiac enlargement, brachycephaly, undershot jaw, microcephaly, microphthalmia, enlarged organs and lordosis or kyphosis. In general, for the analysed morphological parameters, ethanol exposure had a significant effect when compared to the negative control. 3.4. Apoptosis is not affected by ketamine Cellular death was observed using AO staining of embryos/larvae at different time points (24, 48, 72 and 144 hpf). Fig. 4 shows cellular

death pattern in zebrafish embryos at 24 hpf after ketamine and ethanol exposure. At 24 hpf, low natural fluorescence was observed in the brain area, tail region and pericardium zone of the negative control group. The same pattern was observed for ketamine exposed embryos despite an increase in the fluorescence intensity in the pericardium zone with increasing ketamine concentrations. Zebrafish embryos exposed to ethanol showed an increase in the fluorescence, mainly in the tail region. The observed pattern in all groups was maintained at later developmental stages. 3.5. Reactive oxygen species generation did not substantially increase with ketamine Estimation of ROS production was observed with a fluorescence probe, DCFH-DA, in embryos/larvae at different time points (24, 48, 72 and 144 hpf). Fig. 4 shows the spatial distribution of ROS in zebrafish embryos at 24 hpf after ketamine and ethanol exposure. At 24 hpf, low natural fluorescence was found along the body of the negative control group. However, higher fluorescence was observed in the yolk sac and yolk extension. Ketamine exposed embryos presented the same

Table 3 General outcomes of morphological analyses of zebrafish larvae at 144 hpf. Group dose (mg mL−1)

Neg. control 0.0

Ketamine 0.2

0.4

0.8

Pos. control EtOH 2%

Statistical test

p

Fins

0.00 (0.00–0.04)a 0.31 ± 0.06 0.00 (0.00–0.00) 0.19 ± 0.03a 0.00 (0.00–0.02) 0.00 (0.00–0.00) 0.00 (0.00–0.00) 0.00 (0.00–0.04)a 0.00 (0.00–0.08)a 0.00 (0.00–0.02)a 0.00 (0.00–0.00) 0.00 (0.00–0.00) 0.08 ± 0.03a 0.00 (0.00–0.10) 0.00 (0.00–0.02)a 0.00 (0.00–0.13)a 0.00 (0.00–0.06)a 0.67 (0.58–1.19)a

0.00 (0.00–0.00)a 0.35 ± 0.08 0.00 (0.00–0.00) 0.21 ± 0.04a 0.00 (0.00–0.00) 0.00 (0.00–0.04) 0.00 (0.00–0.00) 0.00 (0.00–0.00)a 0.00 (0.00–0.04)a 0.00 (0.00–0.02)a 0.00 (0.00–0.00) 0.00 (0.00–0.00) 0.11 ± 0.05a 0.04 (0.00–0.10) 0.17 (0.08–0.21)ab 0.13 (0.06–0.25)ab 0.17 (0.00–0.27)ab 1.25 (0.94–1.60)a

0.00 (0.00–0.30)a 0.45 ± 0.14 0.00 (0.00–0.00) 0.22 ± 0.09a 0.00 (0.00–0.25) 0.00 (0.00–0.00) 0.00 (0.00–0.00) 0.00 (0.00–0.17)a 0.00 (0.00–0.04)a 0.00 (0.00–0.08)a 0.00 (0.00–0.00) 0.00 (0.00–0.00) 0.08 ± 0.04a 0.08 (0.00–0.17) 0.08 (0.08–0.29)ab 0.17 (0.08–0.17)ab 0.08 (0.00–0.29)ab 1.50 (0.88–2.38)ab

0.08 (0.00–0.21)ab 0.58 ± 0.14 0.00 (0.00–0.00) 0.35 ± 0.09a 0.00 (0.00–0.00) 0.00 (0.00–0.02) 0.00 (0.00–0.00) 0.13 (0.06–0.21)ab 0.08 (0.00–0.10)ab 0.04 (0.00–0.17)ab 0.00 (0.00–0.00) 0.00 (0.00–0.00) 0.21 ± 0.07a 0.04 (0.00–0.10) 0.21 (0.17–0.33)ab 0.25 (0.08–0.42)ab 0.04 (0.00–0.29)ab 2.17 (1.17–3.02)ab

0.33 (0.13–0.44)b 0.80 ± 0.17 0.00 (0.00–0.00) 0.72 ± 0.07b 0.00 (0.00–0.10) 0.00 (0.00–0.02) 0.00 (0.00–0.00) 0.50 (0.42–0.92)b 0.33 (0.25–0.40)b 0.29 (0.25–0.38)b 0.00 (0.00–0.00) 0.00 (0.00–0.00) 0.83 ± 0.16b 0.00 (0.00–0.17) 0.75 (0.56–0.89)b 0.46 (0.08–0.65)b 0.42 (0.15–0.79)b 5.67 (5.40–6.17)b

X2(4) = 11.34 F(4,25) = 2.782 np F(4,25) = 11.34 X2(4) = 5.607 X2(4) = 1.982 np X2(4) = 20.85 X2(4) = 18.09 X2(4) = 18.21 np np F(4,25) = 14.49 X2(4) = 0.484 X2(4) = 22.59 X2(4) = 12.31 X2(4) = 9.822 X2(4) = 19.23

0.023 0.053 np b0.001 0.230 0.739 np b0.001 0.001 0.001 np np b0.001 0.975 b0.001 0.015 0.044 0.001

Heart/thorax

Head

Eyes Organs Tail/spine

Stunted Malformed Missing Cardiac enlarg. Bradycardia Tachycardia Dist. thoracic region Microcephaly Brachycephalic Undershot jaw Enlarged otoliths Dolichocephalic Microphthalmia Ocular oedema Enlarged organs Lordosis/kyphosis Kink in tail Total

Parametric data presented as mean ± SD of six independent replicates. Statistical analysis was performed using one-way ANOVA followed by Tukey's multiple-comparison test. Nonparametric data presented as median (25th–75th quartile) of six independent replicates. Statistical analysis was performed using the Kruskal–Wallis test followed by Dunn's test. Different lowercase letters indicate significant differences between groups (p b 0.05). Abbreviations used: (np) not possible to calculate descriptive statistics.

32

L.M. Félix et al. / Neurotoxicology and Teratology 41 (2014) 27–34

Fig. 4. Cellular death pattern and reactive oxygen species spatial distribution in zebrafish embryos at 24 hpf. Apoptotic staining by acridine orange (AO), from 5 independent experiments was visualized in the negative control, namely in the midbrain (mb), dorsal hindbrain (hb) and around the pericardium zone (pc). Ketamine exposed embryos showed more fluorescence intensity in the pericardium zone with increasing concentrations when compared to the negative control. Ethanol exposure resulted in an increase in the fluorescence intensity namely in the tail region (t). ROS accumulation, assessed using a fluorescent probe, 2′,7′-dicholorfluorescein-diacetate (DCFH-DA), was observed in the negative control mainly in the yolk sac (yk) and in the yolk extension (ys). With ketamine increasing concentrations, an increase in the fluorescence in the pericardium zone (pc) was observed. No differences were observed in the ethanol exposed group. Scale bar represents 500 μm.

pattern of fluorescence distribution. In addition, an increase in the fluorescence was observed in the pericardium zone with increasing ketamine concentrations. Fluorescence in the ethanol group was not significantly higher than that observed in the other groups. The ROS distribution pattern remained the same in all groups at later developmental stages. 4. Discussion The objective of the present study was to evaluate the potential toxicological effects of sub- and over clinical anaesthetic doses of ketamine in vivo using the zebrafish embryo, prior to its action as NMDA receptor inhibitor. The results show that the ketamine exposure induces developmental toxicity as well as increases mortality during blastula stage.

Ketamine, a N-Methyl-D-aspartate (NMDA) receptor antagonist, has been used as a common anaesthetic for surgery, emergency, paediatric and veterinary medicine. However, due to its dissociative and hallucinogenic effects, its use in adults has declined. Nevertheless, ketamine has become one of the most frequently used anaesthetics worldwide (Green et al., 1996), including in obstetric and paediatric anaesthesia procedures (Dong and Anand, 2013). Furthermore, ketamine surfaced into recreational settings which has led to increasing concerns about the harmful consequences of its use (Morgan et al., 2012) particularly by pregnant women (Su et al., 2010). As the placenta is not a barrier to ketamine and it can enter the foetal circulation (Craven, 2007), this potentiates the risk of teratogenic effects after in utero exposure (Su et al., 2010). More recently, the potential risks for human exposure through environmental contamination have become an important

L.M. Félix et al. / Neurotoxicology and Teratology 41 (2014) 27–34

question, since ketamine was detected in aquatic ecosystems in concentrations ranging from 0.05 to 0.4 ng mL−1 (Lin et al., 2010; Baker and Kasprzyk-Hordern, 2011; Vazquez-Roig et al., 2012). Regarding studies using zebrafish as an animal model, behavioural dose-dependent effects after 20 minute exposure to concentrations up to 0.04 mg mL−1 (Riehl et al., 2011; Zakhary et al., 2011) and inducement of motor neuron toxicity in 28 hpf embryos of a transgenic zebrafish line, after 20 hour exposure to a ketamine concentration of 2 mM (=0.55 mg mL−1) (Kanungo et al., 2013) were reported. Another study showed that ketamine can reduce heart rate of zebrafish after a 2 h-treatment with doses of 5 (= 1.37 mg mL−1) and 10 mM (= 2.74 mg mL−1) in 26 and 52 hpf embryos (Kanungo et al., 2012). However, the effect of ketamine during early zebrafish development phases has not been explored. Based on the advantages of this model and physiological similarities to higher vertebrates, it can be used to identify embryogenic ketamine target pathways. In the present study, exposure to ketamine concentrations during the blastula period, which resembles the blastocyte period in higher vertebrate (Richardson et al., 1997), delayed the detachment of the tailbud from the yolk sac and showed a tendency to delay the head development. The highest concentration of ketamine used resulted in a significant increase in pericardial and yolk sac oedema. Also, it was observed, although not significant, a tendency to a heart rate decrease with the increasing ketamine concentration. This heart beat decline was previously observed in zebrafish embryos (Kanungo et al., 2012), although in that study higher doses and longer and different exposure periods were used. In zebrafish, the cardiac precursors have only been identified at the late blastula stage before the start of gastrulation (Stainier et al., 1993) thus, our observed results indicate a non-specific ketamine action. Indeed, in this study, the exposure of embryos to 20 min of ketamine during zebrafish blastula period, before ketamine action as NMDA receptor inhibitor, resulted in development toxicity. This was more evident in the highest ketamine dose used and suggests that ketamine affects the normal zebrafish development through a mechanism independent of NMDA receptors, whose expression begins around 24 hpf during the pharyngula period (Cox et al., 2005). Moreover, it has already been shown that ketamine action is not reserved to the NMDA receptor and also acts via suppression of F-actin polymerisation and microtubular cytoskeletons in HepG2 cells (Chang et al., 2009). Furthermore, the activation of apoptotic pathways in human lymphocytes and neuronal cells was described for ketamine, when used at millimolar concentrations, which is unlikely to be mediated through the NMDA receptor (Braun et al., 2010). Thus, as during the blastula stage the transition from maternal to zygotic expression is occurring, it can be hypothesized that ketamine may act independently from NMDA receptor by silencing the zygotic genome by one of the strategies proposed by Schier (Schier, 2007), resulting in the teratogenic effects observed. In addition, in the present study, zebrafish cell death was assessed using acridine orange and accumulation of reactive oxygen species in embryonic and larval zebrafish was evaluated using a fluorescent probe, DCFH-DA. The results showed no significant effect of ketamine neither on the cellular death pattern nor the ROS accumulation in the developing zebrafish, by 24 hpf. This may be explained by the fact that the ketamine exposure was performed previous to the epiboly stage (between 4 and 7 hpf), and zebrafish embryos are known to resist to apoptosis until reaching that stage, where the activation of the zygotic genome starts to be the primary source of mRNA in the embryo (Ikegami et al., 1999; Negron and Lockshin, 2004). Several studies have showed that zebrafish embryos treated with anticancer drugs (camptothecin, nocodazole and staurosporine) prior to the end of the epiboly stage survived until the mid-gastrula stage, approximately by 7 hpf, at which point they undergo a rapid and synchronous cell death (Eimon and Ashkenazi, 2010). This was also observed in the present study, where mortality rates increased significantly between 8 and 24 hpf, with increasing ketamine concentrations. As no previous studies

33

examined ketamine effects on similar stages of development, further studies should be undertaken in order to elucidate its biological mechanisms of action. 5. Conclusions This study shows that ketamine exposure during blastula phase is teratogenic and toxic to the zebrafish embryos, interfering with the normal developmental zebrafish pathways which ultimately result in a number of morphological anomalies and increased mortality. All evidences from this study indicate that ketamine exerts an independent NMDA receptor action during the zebrafish blastula stage, which should be further studied. Furthermore, as zebrafish displays sufficient genetic similarities to superior vertebrates, this study could provide information on the potential mechanisms of ketamine-induced toxicity during embryogenesis. Acknowledgements This research was financially supported by the Portuguese Foundation for Science and Technology (Lisboa, Portugal) and co-funded by the COMPETE: -01-0124-FEDER-009497 through the project grant: PTDC/CVT/099022/2008. Appendix A. Supplementary data Supplementary data to this article can be found online at http://dx. doi.org/10.1016/j.ntt.2013.11.005. References Abdel-Rahman MS, Ismail EE. Teratogenic effect of ketamine and cocaine in CF-1 mice. Teratology 2000;61(4):291–6. Baker DR, Kasprzyk-Hordern B. Multi-residue analysis of drugs of abuse in wastewater and surface water by solid-phase extraction and liquid chromatography–positive electrospray ionisation tandem mass spectrometry. J Chromatogr A 2011;1218(12): 1620–31. Bandazhevskii I, Shimanovich AI. The characteristics of the embryotoxic action of kalipsol. Farmakol Toksikol 1991;54(5):58–9. Bosnjak ZJ. Developmental neurotoxicity screening using human embryonic stem cells. Exp Neurol 2012;237(1):207–10. Brambrink AM, Evers AS, Avidan MS, Farber NB, Smith DJ, Martin LD, et al. Ketamine-induced neuroapoptosis in the fetal and neonatal rhesus macaque brain. Anesthesiology 2012;116(2):372–84. Braun S, Gaza N, Werdehausen R, Hermanns H, Bauer I, Durieux ME, et al. Ketamine induces apoptosis via the mitochondrial pathway in human lymphocytes and neuronal cells. Br J Anaesth 2010;105(3):347–54. Burgess HA, Granato M. Sensorimotor gating in larval zebrafish. J Neurosci 2007;27(18): 4984–94. Chang HC, Chen TL, Chen RM. Cytoskeleton interruption in human hepatoma HepG2 cells induced by ketamine occurs possibly through suppression of calcium mobilization and mitochondrial function. Drug Metab Dispos 2009;37(1):24–31. Cox JA, Kucenas S, Voigt MM. Molecular characterization and embryonic expression of the family of N-methyl-D-aspartate receptor subunit genes in the zebrafish. Dev Dyn 2005;234(3):756–66. Craven R. Ketamine. Anaesthesia 2007;62:48–53. de la Torre R. Commentary on Morgan et al. (2010): ketamine abuse: first medical evidence of harms we should confront. Addiction 2010;105(1):134–5. Dong C, Anand KJS. Developmental neurotoxicity of ketamine in pediatric clinical use. Toxicol Lett 2013;220(1):53–60. Eimon PM, Ashkenazi A. The zebrafish as a model organism for the study of apoptosis. Apoptosis 2010;15(3):331–49. El-Karim AHB, Benny R. Embryotoxic and teratogenic action of ketamine hydrochloride in rats. Ain Shams Med J 1976;27:459–63. Fleming A, Cambridge C. Zebrafish as an alternative model organism for disease modelling and drug discovery: implications for the 3Rs, 10. NC3Rs (National Centre for the Replacement, Refinement and Reduction of Animals in research); 20071–7. Fraysse B, Mons R, Garric J. Development of a zebrafish 4-day toxicity of embryo-larval bioassay to assess chemicals. Ecotoxicol Environ Saf 2006;63(2):253–67. Green SM, Clem KJ, Rothrock SG. Ketamine safety profile in the developing world: survey of practitioners. Acad Emerg Med 1996;3(6):598–604. Howe K, Clark MD, Torroja CF, Torrance J, Berthelot C, Muffato M, et al. The zebrafish reference genome sequence and its relationship to the human genome. Nature 2013;496:498–503. Ikegami R, Hunter P, Yager TD. Developmental activation of the capability to undergo checkpoint-induced apoptosis in the early zebrafish embryo. Dev Biol 1999;209(2): 409–33.

34

L.M. Félix et al. / Neurotoxicology and Teratology 41 (2014) 27–34

Kanungo J, Cuevas E, Ali SF, Paule MG. L-Carnitine rescues ketamine-induced attenuated heart rate and MAPK (ERK) activity in zebrafish embryos. Reprod Toxicol 2012;33(2):205–12. Kanungo J, Cuevas E, Ali SF, Paule MG. Ketamine induces motor neuron toxicity and alters neurogenic and proneural gene expression in zebrafish. J Appl Toxicol 2013;33(6): 410–7. Kimmel CB, Ballard WW, Kimmel SR, Ullmann B, Schilling TF. Stages of embryonic development of the zebrafish. Dev Dyn 1995;203:253–310. Ko SC, Cha SH, Heo SJ, Lee SH, Kang SM, Jeon YJ. Protective effect of Ecklonia cava on UVB-induced oxidative stress: in vitro and in vivo zebrafish model. J Appl Phycol 2011;23(4):697–708. Kochhar MM, Aykac I, Davidson PP, Fraley ED. Teratologic effects of d,1-2-(o-chlorophenyl)2-(methylamino) cyclohexanone hydrochloride (ketamine hydrochloride) in rats. Res Commun Chem Pathol Pharmacol 1986;54(3):413–6. Li J, Vicknasingam B, Cheung Y, Zhou W, Nurhidayat AW, Des Jarlais DC, et al. To use or not to use: an update on licit and illicit ketamine use. Subst Abuse Rehabil 2011;2:11–20. Lin AY, Wang XH, Lin CF. Impact of wastewaters and hospital effluents on the occurrence of controlled substances in surface waters. Chemosphere 2010;81(5):562–70. Mellon RD, Simone AF, Rappaport BA. Use of anesthetic agents in neonates and young children. Anesth Analg 2007;104(3):509–20. Morgan CJ, Curran HV, Independent Scientific Committee on Drugs. Ketamine use: a review. Addiction 2012;107(1):27–38. Muntean BS, Horvat CM, Behler JH, Aboualaiwi WA, Nauli AM, Williams FE, et al. A comparative study of embedded and anesthetized zebrafish in vivo on myocardiac calcium oscillation and heart muscle contraction. Front Pharmacol 2010;1:1–9. Negron JF, Lockshin RA. Activation of apoptosis and caspase-3 in zebrafish early gastrulae. Dev Dyn 2004;231(1):161–70. Padilla SD, Hunter DL, Padnos B, Frady S, MacPhail RC. Assessing locomotor activity in larval zebrafish: influence of extrinsic and intrinsic variables. Neurotoxicol Teratol 2011;33(6):624–30. Parng C, Seng WL, Semino C, McGrath P. Zebrafish: a preclinical model for drug screening. Assay Drug Dev Technol 2002;1(1):41–8.

Paule MG, Li M, Allen RR, Liu F, Zou X, Hotchkiss C, et al. Ketamine anesthesia during the first week of life can cause long-lasting cognitive deficits in rhesus monkeys. Neurotoxicol Teratol 2011;33(2):220–30. Reimers MJ, Flockton AR, Tanguay RL. Ethanol- and acetaldehyde-mediated developmental toxicity in zebrafish. Neurotoxicol Teratol 2004;26(6):769–81. Richardson MK, Hanken J, Gooneratne ML, Pieau C, Raynaud A, Selwood L, et al. There is no highly conserved embryonic stage in the vertebrates: implications for current theories of evolution and development. Anat Embryol 1997;196:91–106. Riehl R, Kyzar E, Allain A, Green J, Hook M, Monnig L, et al. Behavioral and physiological effects of acute ketamine exposure in adult zebrafish. Neurotoxicol Teratol 2011;33(6):658–67. Rohde LA, Heisenberg CP. Zebrafish gastrulation: cell movements, signals, and mechanisms. Int Rev Cytol 2007;261:159–92. Schier AF. The maternal–zygotic transition: death and birth of RNAs. Science 2007;316(5823):406–7. Selderslaghs IWT, Van Rompay AR, De Coen W, Witters HE. Development of a screening assay to identify teratogenic and embryotoxic chemicals using the zebrafish embryo. Reprod Toxicol 2009;28(3):308–20. Soares J, Coimbra AM, Reis-Henriques MA, Monteiro NM, Vieira MN, Oliveira JM, et al. Disruption of zebrafish (Danio rerio) embryonic development after full life-cycle parental exposure to low levels of ethinylestradiol. Aquat Toxicol 2009;95(4):330–8. Stainier DY, Lee RK, Fishman MC. Cardiovascular development in the zebrafish. I. Myocardial fate map and heart tube formation. Development 1993;119(1):31–40. Su PH, Chang YZ, Chen JY. Infant with in utero ketamine exposure: quantitative measurement of residual dosage in hair. Pediatr Neonatol 2010;51(5):279–84. Tucker B, Lardelli M. A rapid apoptosis assay measuring relative acridine orange fluorescence in zebrafish embryos. Zebrafish 2007;4(2):113–6. Vazquez-Roig P, Andreu V, Blasco C, Morillas F, Pico Y. Spatial distribution of illicit drugs in surface waters of the natural park of Pego–Oliva Marsh (Valencia, Spain). Environ Sci Pollut Res Int 2012;19(4):971–82. Zakhary SM, Ayubcha D, Ansari F, Kamran K, Karim M, Leheste JR, et al. A behavioral and molecular analysis of ketamine in zebrafish. Synapse 2011;65(2):160–7.

Ketamine NMDA receptor-independent toxicity during zebrafish (Danio rerio) embryonic development.

Concerns have been raised that the effect of anaesthetic drugs on the central nervous system may result in long-term impairment, namely when ketamine ...
791KB Sizes 0 Downloads 0 Views