- Email: [email protected]
Thiamethoxam induces meiotic arrest and reduces the quality of oocytes in cattle
Toxicology in Vitro 61 (2019) 104635
Contents lists available at ScienceDirect
Toxicology in Vitro journal homepage: www.elsevier.com/locate/toxinvit
Thiamethoxam induces meiotic arrest and reduces the quality of oocytes in cattle ⁎
Zheng-Wen Niea, Ying-Jie Niua, Wenjun Zhoua, Ju-Yeon Kima, Sun A. Ockb, , Xiang-Shun Cuia, a b
T
⁎
Department of Animal Sciences, Chungbuk National University, Chungbuk, Cheongju 361-763, Republic of Korea Animal Biotechnology Division, National Institute of Animal Science, Rural Development Administration, Jeonju 55365, Republic of Korea
A R T I C LE I N FO
A B S T R A C T
Keywords: Thiamethoxam Bovine oocyte ROS Mitochondria Apoptosis
Thiamethoxam (TMX) is a neonicotinoid insecticide, the residues of which have been detected on various crops. In addition to its specific acetylcholine toxicity to insects, TMX was also found to be toxic to mammals. Moreover, oocytes are vulnerable to reactive oxygen species (ROS). Excessive ROS production can override antioxidant defenses and produce oxidative stress and DNA damage that trigger apoptosis and necrosis in organisms. In this study, we exposed bovine oocytes to TMX during maturation. Microscopic examination showed that 1.6 mM TMX significantly inhibited maturation at the germinal vesicle (GV) and metaphase I (MI) stages. Immunofluorescence staining and enzyme activity analysis revealed that TMX induced a reduction in CDC25 and CDC2 activity. Furthermore, time-lapse tracking and immunofluorescence staining indicated the maintenance of cyclin B in the cytoplasm, persistence of Bub3 at kinetochores, and absence of actin caps after TMX-exposed oocytes reached the MI stage. In addition, metaphase II (MII) oocytes exposed to TMX showed disordered chromosomes and spindles. These oocytes accumulated excess ROS and showed significantly decreased mitochondrial membrane potential and increased apoptotic signals. Parthenogenetic embryos from these oocytes showed decreased percentages of morulae and blastocysts. These results indicate that TMX delays bovine oocyte progression to MI stage, blocks them at the MI stage, triggers disordered chromosomes and spindles at MII stage, and ultimately results in MII oocytes with poor cleavage ability and inhibited development to morulae and blastocysts.
1. Introduction Neonicotinoid insecticides are pesticides widely used on agriculture crops that mediate their toxic effects by binding to nicotinic acetylcholine receptors (nAChRs). These insecticides have a high affinity for insect nAChRs and low toxicity to mammals, lacking teratogenic and mutagenic effects, and therefore can be extensively applied (Jeschke and Nauen, 2008). Imidacloprid, thiamethoxam (TMX), thiacloprid (THI), nitenpyram, acetamiprid, and clothianidin (CLO) are currently the most commonly used neonicotinoids (Simon-Delso et al., 2015). Among these neonicotinoids, TMX has highest IC50 for insect and vertebrate α4β2 receptors (Taillebois et al., 2018). This indicates that TMX has a relatively low toxicity to non-target insects and is safer for vertebrates compared to other neonicotinoids. However, it is very likely that this characteristic of TMX will result in its excessive use, which could pose a danger to mammals. TMX residue has been found in sheep and cow milk when TMX was used widely (Fedrizzi et al., 2019). Besides, a previous study indicated that among neonicotinoids, TMX had
⁎
the highest cytotoxicity to SH-SY5Y and HepG2 cells and TMX and THI more readily induced DNA damage in HepG2 cells compared to that with other neonicotinoids (Senyildiz et al., 2018). With regards to cytotoxicity, TMX produces metabolites such as CLO, desmethyl-CLO, desmethyl-TMX, and formaldehyde, which are harmful to cell metabolism These products are generated through catalysis or the inhibition of metabolic enzymes (Wang et al., 2018). For example, formaldehyde derived from TMX, dmTMX, or CLO results in mouse-specific hepatotoxicity/hepatocarcinogenicity (Swenson and Casida, 2013). In terms of DNA damage, TMX can also exert other toxic effects via different molecular mechanisms. Neonicotinoids can perturb mitochondrial function and induce increased levels of cytochrome P450 (CYP450) and aldehyde oxidase (AOX), resulting in excess ROS production as a consequence of mitochondrial dysfunction. Excess ROS can in turn induce oxidative stress, which might result in the oxidation of lipids, proteins, and DNA (Wang et al., 2018). Meanwhile, TMX inhibits normal apoptosis in the liver of rabbits (El Okle et al., 2018). In addition, TMX has a tertiary and specific effect on cells. Using co-cultures of
Corresponding authors. E-mail addresses: [email protected] (S.A. Ock), [email protected] (X.-S. Cui).
https://doi.org/10.1016/j.tiv.2019.104635 Received 8 July 2019; Received in revised form 18 August 2019; Accepted 29 August 2019 Available online 31 August 2019 0887-2333/ © 2019 Published by Elsevier Ltd.
Toxicology in Vitro 61 (2019) 104635
Z.-W. Nie, et al.
(caption on next page)
2
Toxicology in Vitro 61 (2019) 104635
Z.-W. Nie, et al.
Fig. 1. Effect of thiamethoxam (TMX) toxicity on bovine oocyte maturation. Maturation rates of bovine oocytes exposed to different concentrations of TMX (A) showed that 1.6 mM TMX significantly decreased the rate of oocyte maturation. Spindles and chromosomes at different stages in the development of oocytes treated with or without TMX are shown (B). Oocytes showed a spindle ball and disordered chromosomes at the germinal vesicle breakdown (GVBD) stage, a bipolar spindle and chromosomes in a belt at the pre-metaphase I (MI) stage, or a bipolar spindle and aligned chromosomes at the pre-MI stage. Metaphase II (MII) oocytes had two parts of spindles and chromosomes, which belonged to the secondary oocyte and first polar body, respectively. Normal secondary oocytes had the same spindle and chromosomes as MI oocytes, and abnormal secondary oocytes had disordered or cohesive spindles or chromosomes. Portions of GVBD, MI, and MII oocytes after different durations of in vitro maturation (C) and changes in the percentages of oocytes at the GVBD, MI, and MII stages during maturation (D), showing delayed or arrested stages. Percentages of oocytes with abnormal spindles and chromosomes (E) showing increased rates of abnormal MII oocytes in the TMX group. The scale bar in the merged figures = 30 μm. The scale bar in enlarged figures = 8 μm. “n” represents the total number of oocytes in each stage.
mL epidermal growth factor (E4127; Sigma), 100 IU/mL penicillin/ streptomycin, 22 μg/mL sodium pyruvate (P5280; Sigma), and 95 μg/ mL L-cysteine (C7352; Sigma) at 38.5 °C in a humidified atmosphere of 5% CO2 and 95% air.
H295R and BeWo cells, it was shown that TMX induced CYP3A7 (cytochrome P450 family 3 subfamily A member 7) expression, which enhanced the activity of aromatase, in turn resulting in increased synthesis of estradiol (Caron-Beaudoin et al., 2017). Similarly, research has indicated that TMX induces PII- and I.3-mediated CYP19 expression and aromatase activity (Caron-Beaudoin et al., 2016). MtDNA and copy number of mitochondria increase during oocyte maturation (Cecchino et al., 2018), and their number in matured oocytes is high (St John, 2014). Oocyte maturation is associated with high levels of mitochondrial activity, during which, the mitochondria concentrate around the spindle and are involved in chromosome separation (Dalton and Carroll, 2013). During polar body extrusion, the levels of ATP generated by mitochondria peak (Dalton et al., 2014). In addition, mitochondria are an important source of ROS owing to the production of superoxide by the respiratory chain (Garcia-Ruiz and FernandezCheca, 2018). In normal cells, peroxide can be removed; however, for oocytes with dysfunctional mitochondria or unbalanced redox reactions, ROS are generated, which has a detrimental effect on oocyte maturation. In this regard, it has previously been found that bovine oocytes extracted from ovaries at the germinal vesicle (GV) stage initially have relatively high levels of ROS, which gradually decrease during the first 4 h of in vitro maturation, and are then maintained at a relatively low level in the subsequent stages of maturation (Morado et al., 2009). This suggests that bovine oocytes have anti-oxidative capacity and that certain levels of ROS are necessary for meiosis to proceed in oocytes. Despite the residual TMX found on crops, of which of stalks are usually used to feed cattle (Telo et al., 2015; Veysset et al., 2014), the presence excess ROS derived from TMX and the specific anti-oxidative capacity of bovine oocytes, and specifically whether excess ROS from TMX has toxicity to bovine oocytes, are unknown. In the present study, we hypothesized that TMX induces the excess production of ROS and mitochondrial dysfunction. ROS-triggered peroxidation and defects in mitochondrial energy generation would then damage biomolecules involved in maturation, blocking bovine oocytes at the MI (metaphase I) stage, trapping them at the anaphase I (AI)/ telophase I (TI) stage, and inducing chromosome and spindle disorder at the metaphase II (MII) stage. Additionally, the maternal-derived materials required for embryo development in oocytes would also be disturbed. Consequently, these MII oocytes with poor potential for embryo development would lose their ability to undergo cleavage and develop into morulae and blastocysts. Collectively, the damage caused by TMX would finally result in apoptosis.
2.2. Parthenogenetic activation and in vitro embryo culture After maturation for 24 h, cumulus cells surrounding oocytes were removed by dispersion with 0.1% hyaluronidase and repeated pipetting for approximately 1 min. For parthenogenetic activation, oocytes with first polar bodies were selected, incubated in CR1-aa (6.7 mg/mL NaCl (S5886; Sigma), 0.23 mg/mL KCl (P5405; Sigma), 2.2 mg/mL NaHCO3 (S5761; Sigma), 0.55 mg/mL hemicalcium lactate (L4388; Sigma), 1% MEM non-essential amino acids (M7145), 2% BMEM essential amino acids (B6766)) supplemented with 3 mg/mL BSA (A8806; Sigma) and 5 mM ionomycin (I0634; Sigma) for 7 min, and then in CR1-aa supplemented with 2 mM 6-dimethylaminopurine (D2629; Sigma) and 7.5 mg/mL cytochalasin B (C6762; Sigma) for 3 h. The oocytes were then washed with IVC-1 (CR1-aa supplemented with 10% BSA, 0.31 mg/mL glutathione (G6013; Sigma), 100 IU/mL penicillin/streptomycin, 44 μg/mL sodium pyruvate, and 0.146 mg/mL glutamine), cultured in IVC-1 for 4 d and then in IVC-2 (IVC-1 in which 10% BSA was replaced with 10% fetal calf serum) for 3 d in a humidified atmosphere of 5% CO2 and 95% air. 2.3. Immunofluorescence staining and confocal microscopy Oocytes were fixed in 3.7% paraformaldehyde for 1 h at room temperature, washed three times with phosphate-buffered saline/ polyvinyl alcohol (PBS/PVA), permeabilized with PBS/PVA containing 1.0% triton X-100 at room temperature for 1 h, and washed three times with PBS/PVA. The oocytes were then blocked in PVA-PBS containing 1.0% bovine serum albumin at room temperature for 1 h and subsequently incubated overnight at 4 °C with an anti-α-tubulin-FITC antibody (1:100; Sigma; USA), anti-γH2AX antibody (1:100, pS139; Cell Signaling Technology, Danvers, MA, USA), anti-CDC25C antibody (1:100, sc-5620; Santa Cruz Biotechnology, Dallas, TX, USA), anti-CDC2 antibody (1:100, pT161; Cell Signaling; USA), anti-actin antibody (1:100, sc-1616; Santa Cruz Biotechnology), or an anti-Bub3 antibody (1:50, sc-28,258; Santa Cruz Biotechnology). After incubation with the primary antibody (except for anti-α-tubulin-FITC antibody), the oocytes were washed three times with PBS/PVA, incubated at 37 °C for 1 h with a secondary IgG antibody, stained with Hoechst 33342 (bisBenzimide H33342 trihydrochloride, 1:2000; Sigma Life Science) for 15 min, washed three times in PBS/PVA, mounted on slides, and examined under a confocal microscope (Zeiss LSM 710 META; Jena, Germany). Images were processed using Zen software (version 8.0, Zeiss).
2. Materials and methods 2.1. Oocyte collection and in vitro maturation Ovaries from female Hanwoo cattle were obtained from a local slaughterhouse, maintained in saline at 38.5 °C, and transported to the laboratory. Cumulus oocyte complexes were extracted from follicles using an injection syringe, washed three times in Tyrode's lactate-4-(2hydroxyethyl)-1-piperazineethanesulfonic acid, and cultured in tissue culture medium 199 (TCM 199) supplemented with 10% fetal calf serum (10082–147; Gibco, USA), 1 μg/mL β-estradiol (E2758; Sigma, USA), 10 μg/mL follicle-stimulating hormone (F2293; Sigma), 10 ng/
2.4. ROS staining The ROS contents of the treated and untreated oocytes were measured after oocytes had been cultured for 24 h. ROS content was quantified using the dichlorodihydrofluorescein diacetate (DCHFDA, Molecular Probes, Invitrogen, Carlsbad, CA, USA) method, as described previously (Yi et al., 1990). Briefly, the cumulus cells were removed from cumulus oocyte complexes with hyaluronidase. Denuded oocytes 3
Toxicology in Vitro 61 (2019) 104635
Z.-W. Nie, et al.
Fig. 2. Molecular regulation from germinal vesicle breakdown (GVBD) to metaphase I (MI) stages. Immunofluorescence staining for CDC25 at the GV and GVBD stages (A) and relative intensity of CDC25 fluorescence (B), showing higher CDC25 activity in the thiamethoxam (TMX) group. Immunofluorescence staining of CDC2 at the GV, pre-MI, and MI stages (C), relative intensity of CDC2 fluorescence (D), and relative activity of CDC2 extracted from oocytes (E), showing lower CDC2 activity in the TMX group between 8 and 12 h. The scale bar in the merged figures = 30 μm. The scale bar in enlarged figures = 8 μm. “n” represents the total number of oocytes in each stage.
2.5. Analysis of mitochondrial membrane potential
were exposed to DCHFDA diluted in PBS/PVA (1:1000) for 15 min and then washed three times with PBS/PVA. Live imaging and quantitation were conducted using a fluorescence microscope (Nikon, Tokyo, Japan) and Photoshop (CS2; Adobe, San Jose, CA, USA). Then Image J software was used to filter out the contaminated spots with strong intensity and calculate the ROS signal of oocytes.
To measure mitochondrial membrane potential, denuded oocytes were washed three times with PBS/PVA and incubated with 5,5′,6,6′tetrachloro-1,1′,3,3′-tetraethyl-imidacarbocyanine iodide (JC-1) (Invitrogen) diluted in culture medium (1:200) for 30 min. The membrane potential was calculated as the ratio of red florescence, 4
Toxicology in Vitro 61 (2019) 104635
Z.-W. Nie, et al.
Fig. 3. Molecular regulation from metaphase I (MI) to anaphase I (AI) stages. Time-lapse of H2B-Cherry and cyclin B-eGFP (A) and relative intensity of cyclin B-eGFP fluorescence (C), showing the maintenance of cyclin B1 from 12 to 18 h (Scale bar = 60 μm). The extrusion rate of polar bodies in the thiamethoxam (TMX) group at 18 h (B) was decreased significantly. Immunofluorescence staining of actin (D) and percentage of oocytes with actin caps (F) at 18 h, showing that actin caps could not be formed in the TMX group (Scale bar = 30 μm). Immunofluorescent staining for BUB3 (E) and percentage of oocytes with BUB3 puncta at 18 h (G), showing the maintenance of puncta in the TMX group (Scale bar = 4 μm). “n” represents the total number of oocytes in each stage.
(50 mM tris-HCl (pH 7.5), 0.5 M NaCl, 5 mM EDTA, 2 mM EGTA, 0.01% Brij-35 (v/v), 1 mM PMSF, 0.05 mg/mL leupeptin, 50 mM 2-mercaptoethanol, 25 mM β-glycerophosphate, and 1 mM sodium orthovanadate) and stored at −80 °C. Sample extracts were quantified by ELISA using a MESACUP cdc2 Kinase Assay Kit (MBL International, Nagoya, Japan), and the optical density of each well was read at 492 nm using a microplate reader.
corresponding to activated mitochondria (J-aggregates), to green fluorescence, corresponding to less-activated mitochondria (J-monomers). Fluorescence was visualized using a Zeiss inverted confocal microscope equipped with a × 40 oil immersion objective (Zeiss, Jena, Germany). Images were processed using ZEN software (Zen Software, Manchester, UK). 2.6. Detection of early apoptosis
2.8. Statistical analysis
Apoptosis was detected using an Annexin V-FITC-PI Apoptosis Detection kit (Vazyme, A211-01) according to the manufacturer's instructions. Briefly, 20 oocytes were washed in PBS/PVA and then incubated in 100 mL binding buffer containing 5 mL of Annexin V-FITC for 10 min in the dark and washed twice in PBS/PVA. The green FITC signal was detected using a fluorescent microscope with 450–490 nm (excitation) and 520 nm (emission) filters.
All results were subjected to a one-way analysis of variance, and differences between the treatment groups were assessed using a least significant difference test in the Statistical Package for Social Sciences (SPSS) software. Each experiment was conducted at least three times, and differences were considered significant when P < .05 or highly significant when P < .01. The relative value of each repeat in the control group was set to 1, and the values of the treatment group were standardized to those of the control.
2.7. In vitro p34cdc2 kinase assay Thirty oocytes, as one sample, were lysed with 5 μL of sample buffer 5
Toxicology in Vitro 61 (2019) 104635
Z.-W. Nie, et al.
Fig. 4. Reactive oxygen species (ROS), mitochondrial membrane potential (MMP), and apoptosis induced by thiamethoxam (TMX) at 24 h. ROS staining (A) and relative levels of ROS (B), showing increased ROS in the TMX group (Scale bar = 120 μm). JC-1 staining (C) and relative intensity of red to green staining (D), showing decreased MMP in the TMX group (Scale bar = 120 μm). Immunofluorescence staining for γH2AX (E) and relative intensity of fluorescence (F), showing no significant difference in γH2AX between the TMX group and control groups (Scale bar = 30 μm). AnnexinV staining (G) and relative intensity (H), showing early apoptosis in the TMX group (Scale bar = 120 μm). “n” represents the total number of oocytes in each stage. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)
6
Toxicology in Vitro 61 (2019) 104635
Z.-W. Nie, et al.
Fig. 5. Development of embryos derived from oocytes treated with or without thiamethoxam (TMX). Embryos derived from oocytes treated with or without TMX at day 7 (A) and percentages at each stage (B), showing arrest before the 8-cell stage. Scale bar = 240 μm. “MO” and “BL”, respectively, represent morula and blastocyst stage. “n” represents the total number of oocytes in each stage.
3. Results 3.1. TMX affects bovine oocyte maturation To determine the toxicity of TMX during bovine oocyte maturation, we examined the effects of a concentration gradient of this agent according to its lowest effective dose in previous study (Green et al., 2005). GV oocytes were cultured and exposed to various concentrations of TMX for 24 h to determine the appropriate concentration of TMX for subsequent analyses. Compared to 0 mM TMX, 1.6 mM TMX significantly reduced the maturation rate of oocytes (65.27% ± 3.57% vs. 51.68% ± 4.70%; P < .05) and that 3.2 mM TMX resulted in a highly significant reduction (65.27% ± 3.57% vs. 39.67% ± 4.74%; P < .01; Fig. 1A). Therefore, we established 1.6 mM TMX as an appropriate dose to study the effect of TMX on oocyte maturation. To further study the process of bovine oocyte maturation, oocytes cultured for 0, 8, 12, and 24 h, which corresponds to the GV, germinal vesicle breakdown (GVBD), MI, and MII stages of maturation, respectively (Tosti et al., 2000), were exposed to 1.6 mM TMX and subsequently analyzed by immunofluorescence staining. The characteristics of oocytes at each stage were determined by tracking spindles and chromosomes, as shown in Fig. 1B. Oocytes contained a nucleus at the GV stage, a ball of microtubes and scattered chromosomes at the beginning of the GVBD stage, a dipolar spindle and aligned chromosomes at the MI stage, and two sets of spindles and chromosomes at the MII stage. The pattern of maturation was altered when oocytes were exposed to TMX (Fig. 1C). For the convenience of analysis, oocytes at each stage were individually tracked during the course of culture (Fig. 1D and Table S1). At 8 h, the proportion of GVBD oocytes was highly significantly decreased compared to that in the control group (84.00% ± 2.40% vs. 24.90% ± 3.02%; P < .01) but recovered to
Fig. 6. Model of the mechanism to explain how thiamethoxam (TMX) affects bovine oocyte maturation. During bovine oocyte maturation, TMX induces excess accumulation of reactive oxygen species (ROS), a decrease in mitochondrial membrane potential (MMP), and apoptosis. These changes affect the maturation and developmental potential of bovine oocytes, reducing the level of active CDC25, and subsequently retarding the phosphorylation of CDC2. Consequently, there is a delay in the progression of oocytes to the germinal vesicle breakdown stage. After this stage, the TMX-induced effects cause damage including inhibition of the inactivation of the spindle assembly checkpoint (SAC), suppression of cyclin B degradation, and reduced formation of actin caps, which are essential for oocytes to successfully enter the AI stage. Exposure to TMX also diminishes the quality of metaphase II oocytes due to chromosomal abnormalities, which reduce the ability of oocytes to develop into embryos at all stages.
7
Toxicology in Vitro 61 (2019) 104635
Z.-W. Nie, et al.
3.4. TMX induces increases in ROS and apoptosis
control levels at 12 h. The proportion of MI oocytes in the TMX group was significantly lower than that in the control group only at 12 h (63.51% ± 6.71% vs. 39.50 ± 9.28%; P < .01), whereas the percentage of MII oocytes exposed to TMX decreased only at 24 h. At the long pre-MI stage, it was difficult to quantify the proportions based on two-dimensional images because of the specific three-dimensional structure of chromosomes in the cricoid metaphase band at this long stage (Kitajima et al., 2011). However, the oocytes at both GVBD (including all stages except the GV stage) and the MII stage showed no significant changes. Based on these observations, we concluded that TMX blocked oocytes before the MI stage. Similarly, at 24 h, TMX blocked oocytes prior to the AI/TI stage. We subsequently selected all MII oocytes and examined them for abnormal spindle and chromosome development. Compared to that in the control group, the percentage of abnormal MII oocytes, in which the spindle and chromosomes were disordered, increased in the TMX group during the maturation process (14.27 ± 10.39% vs. 31.89 ± 11.75%; P < .01); those MII oocytes with cohesive chromosomes (6.50 ± 1.80%) only appeared in the TMX group, but the rate was not significantly different from that in the control group (Fig. 1B and E).
We initially detected ROS using DCHFDA at 24 h. The average relative intensity of the ROS signal in the TMX group was significantly higher than that in the control group (1 vs. 1.33 ± 0.07; P < .05; Fig. 4A and B). A cellular imbalance of redox reactions is one factor contributing to the production of ROS, and this process is related to mitochondrial dysfunction. We therefore examined mitochondrial membrane potential (MMP), which revealed that the MMP of oocytes exposed to TMX was significantly decreased (1% vs. 0.81% ± 0.04%; P < .05; Fig. 4C and D). We subsequently evaluated the damage induced by ROS. Previous studies (Perez et al., 2007; Yuh et al., 2010) have indicated the possibility that ROS production induced by TMX triggers DNA damage. Therefore, to assess potential DNA damage, we selected γH2AX as a marker to stain for (Mayer et al., 2016; Omata and Kato, 2019). However, we observed no significant difference in γH2AX between the control and TMX groups (1 vs. 1.20 ± 0.21; P > .05; Fig. 4E and F), indicating that exposure to TMX does not result in DNA damage. Nevertheless, the higher intensity of the Annexin-V signals in the membrane of TMX-treated oocytes than that in the control indicated that TMX does induce early apoptosis at 24 h (2.33 ± 1.45 vs. 24.88 ± 4.99; P < .05; Fig. 4G and H).
3.2. TMX delays the activation of maturation promoting factor before the MI stage
3.5. TMX results in poor early embryonic development To assess the potential for embryonic development, we evaluated parthenogenetic embryos at each stage. We accordingly found that the percentages of TMX-treated embryos at 1-cell and 2–8-cell stages were significantly increased compared to those in respective controls (Fig. 5A and B, Table S2). In contrast, we observed significant decreases at the morula and blastocyst stage.
To determine how oocytes are arrested, we performed immunofluorescence staining for active CDC25 and CDK2. We accordingly found that oocytes in the TMX group had higher CDC25 activity compared to that in the control group at 12 h (1 vs. 1.97 ± 0.21; P < .05; Fig. 2A and B). Moreover, as described in previous studies, we found that the phosphorylation of CDC2 decreased from 8 to 12 h in the control group; however, the phosphorylation showed a marginal increase in the TMX group and was invariably lower (8 h: 1 vs. 0.71 ± 0.05; P < .05; 12 h: 0.90 ± 0.13 vs. 0.74 ± 0.10; P < .05; Fig. 2C and D). This suggested increased activity of CDC2 in the control group and decreased activity in the TMX group. Here, the specificity of the P-CDC2 antibody was tested by western blotting and a single band for P-CDC2 indicated the accuracy of assessing P-CDC2 in Fig. 2C and D (Fig. 2E). Additionally, we assessed substrate catalysis and the results showed that enzymatic activities decreased significantly compared to those in the control group at 8 h (1 vs. 0.71 ± 0.03; P < .05; Fig. 2F), but were not significantly changed at 12 h (1 vs. 0.99 ± 0.10; P > .05; Fig. 2F).
4. Discussion Neonicotinoid including TMX has anti-reproductive cytotoxic effects on animals and increases ROS or RNS formation in vivo (Wang et al., 2018). Similarly, it has previously been shown that TMX induces oxidative stress and an antioxidant response in the zebrafish liver (Yan et al., 2016). Consistent with these previous studies, we found that TMX induces high levels of ROS, as determined by DCHFDA levels. Only TMX at a concentration of > 1.6 mM could disturb bovine oocyte maturation, which indicates that oocytes have an effective antioxidant control system. The inhibition of GVBD induced by TMX could be overcome at 12 h; however, thereafter, the arrest between the MI and MII stages could not be recovered. In normal in vitro mature cultures, bovine oocytes show a rapid reduction in ROS, which are subsequently maintained at low levels until 12 h, after which the levels gradually increase (Morado et al., 2009). This indicates that bovine oocytes have effective antioxidant capacity prior to the MI stage, which subsequently wanes. ROS and mitochondrial dysfunction generally occur concomitantly. Peroxide can perturb mitochondrial function and trigger a transition in mitochondrial permeability (Vercesi et al., 1997). In addition, although ROS generated by mitochondria can be removed by antioxidative enzymes, mitochondrial dysfunction such as the inhibition of complex I and a high NADH:NAD+ ratio results in the accumulation of excess ROS, which is damaging to cells (Adam-Vizi and Chinopoulos, 2006). An important change observed in dysfunctional mitochondria is a decrease in MMP. The mechanism underlying the adverse effects of neonicotinoids has been evaluated and it was suggested that these chemicals can disrupt mitochondrial function and that impaired mitochondria consequently produce increased amounts of ROS (Wang et al., 2018). Indeed, in the present study, we also demonstrated the association of low MMP and excess ROS. These changes can promote cell apoptosis, which has been observed in many previous studies (Girish et al., 2013; Li et al., 2016; Yang et al., 2017). During the early phase of apoptosis, no apoptotic bodies form; however, cells in this phase exhibit exposed
3.3. TMX arrests oocytes at the MI stage To determine whether some oocytes were arrested between the MI and MII stages when exposed to TMX, we tracked chromosomes and cyclin B by time-lapse from 12 to 18 h. As shown in Fig. 3A–C, whereas oocytes in the control group developed a polar body, those exposed to TMX did not, as determined by the pole body listing percentages at 18 h (58.12% ± 5.06% vs. 52.98% ± 6.10%; P < .05; Fig. 3B). Consistent with this observation, in the control group, the cyclin B1-EGFP signal gradually diminished during pole body listing, whereas in the TMX group, the signal was maintained (Fig. 3A–C, Movies S1 and S2). The progression of oocytes from the AI/TI stage not only requires degradation of cyclin B1 but also the coordination of cytoplasmic and nuclear division. To examine these processes, staining for actin and Bub3 was performed at 18 h, and the results showed that compared to those in the control group, actin caps did not form in oocytes exposed to TMX (23.74% ± 2.58% vs. 15.22% ± 3.30%; P < .05; Fig. 3D and E). In addition, in contrast to that observed in the control group, the mottled signal for Bub3 in the TMX group indicated that the spindle assembly checkpoint was still active (Fig. 3F and G).
8
Toxicology in Vitro 61 (2019) 104635
Z.-W. Nie, et al.
Given the marked decrease in the proportion of morulae observed in the TMX group, these materials are assumed to be vital for major genome activation.
phosphatidylserine on the outer surface of the plasma membrane, to which Annexin-V can bind (Wuest et al., 2019). ROS can trigger apoptosis by disrupting mitochondria (Lee and Lee, 2018). Consistently, in the present study, we detected positive Annexin-V signals only in the membranes of oocytes exposed to TMX. However, although it has previously been found that increased oxidative stress can result in DNA damage (Driessens et al., 2009; Stone and Yang, 2006), our results indicate that ROS induced by TMX does not trigger DNA damage in bovine oocytes. Given that there have been no previous reports of the detection of γH2AX (a marker of DNA damage) in bovine oocytes, DNA damage is probably an infrequent event in these oocytes. Events during oocyte maturation are controlled by a series of proteins, among which the maturation promoting factor (MPF), consisting of CDC2 and cyclin B, regulates meiosis. Although the timing of events during bovine oocyte maturation might differ depending upon the medium in which they are cultured, the activity of MPF shows a significant increase at the GVBD stage, peaks at the MI stage, decreases at the AI stage, and subsequently increases at the MII stage (Wehrend and Meinecke, 2001; Wu et al., 1997). When MPF activity is inhibited, bovine oocytes become arrested at the GV stage (Beker-van Woudenberg et al., 2006). Cdc25 dephosphorylates and activates cdc2 kinase at Thr14 and Tyr15 (Nurse, 1997). In addition, it has been found that Cdc25C, in conjunction with Zn2+, dephosphorylates its substrate MPF/cdk1 in Xenopus oocytes (Lincoln et al., 2002). Accordingly, it can be reasoned that the delay in GVBD observed in the present study is due to the inactivation of CDC25C and the resulting inactivation of cdc2 kinase. In addition, degradation of cyclin B is required for the transition from the MI to the MII stage. It is not cyclin B2 but cyclin B1 that is responsible for progression from the MI stage and entry into the AI stage in mouse oocytes (Ledan et al., 2001). Consistently, we found that cyclin B1 was degraded in normal bovine oocytes. However, this process was inhibited by a significant degree in the TMX group. This suggests that TMX results in failed events related to the AI stage. At the MI stage, the spindle assembly checkpoint (SAC) in the active state can ensure the correct alignment of chromosomes and all the requisite connections of kinetochores with microtubules. Inactive SAC triggers the anaphase-promoting complex/cyclosome (APC/C)–CDC20-dependent degradation of cyclin B1. In this context, it has previously been shown that disturbances in Bub3, a member of the SAC, result in the incorrect attachment of microtubules to kinetochores, misaligned chromosomes, abnormal polar bodies, and aneuploidy (Li et al., 2009). Furthermore, it has been shown that the actin cap, which is under the control of actin-capping proteins, plays a role in extrusion of the first polar body during the maturation of mouse oocytes (Jo et al., 2015). Therefore, exit from the MI stage is controlled by multiple molecular mechanisms. Consistent with the aforementioned findings, the results obtained in the present study indicate that TMX-induced ROS blocks the inactivation of SAC and formation of an actin cap. Oocytes store mRNAs and proteins such as cyclin A2 and histone deacetylases for early embryonic development (Kanka et al., 2012). During bovine preimplantation development, embryos gradually lose the control exerted by maternal materials and show increasing expression of zygotic genes (Kanka et al., 2009). The commencement of major genome activation in bovine embryos is assumed to occur at the 8-cell stage because the expression pattern of genes at this stage is significantly different from that at the previous stage (Kanka et al., 2009; Memili and First, 2000). Consequently, the development of bovine embryos prior to the 8-cell stage mainly depends on the influence of maternal materials from oocytes. In the present study, we observed that embryos at all stages sustained damage resulting from the exposure of oocytes to TMX. It is noteworthy that blocks in embryo development gradually increased prior to the morula stage, whereas thereafter the differences between the TMX and control groups declined. Considering aforementioned points, we can conclude that TMX-induced ROS might destroy materials stored in oocytes that are required for further embryonic development, which results in arrest prior to the 8-cell stage.
5. Conclusion TMX exerts multiple effects on bovine oocyte maturation and quality. It reduces the level of active CDC25 and subsequently blocks the phosphorylation of CDC2. As a result, the development of bovine oocytes is arrested at the GVBD stage. However, progression of the maturation process can be recovered by certain repair mechanisms. In addition, this results in the maintenance of SAC activity at the MI stage or the inhibition of cyclin B degradation and the formation of actin caps. Consequently, oocytes are arrested at the MI stage or fail to enter the AI stage. However, even those oocytes that do progress from the MI stage and develop to the MII stage show evidence of TMX-induced damage, including abnormal chromosomes, apoptosis, the accumulation of ROS, decreased MMP, and poor embryonic developmental potential (Fig. 6). Supplementary data to this article can be found online at https:// doi.org/10.1016/j.tiv.2019.104635. Declaration of Competing Interest The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper. Acknowledgments This research was supported by the Basic Science Research Program through the National Research Foundation of Korea (NRF) funded by the Ministry of Education (No. 2018R1A2B6001173), Republic of Korea. References Adam-Vizi, V., Chinopoulos, C., 2006. Bioenergetics and the formation of mitochondrial reactive oxygen species. Trends Pharmacol. Sci. 27, 639–645. Beker-van Woudenberg, A.R., Zeinstra, E.C., Roelen, B.A., Colenbrander, B., Bevers, M.M., 2006. Developmental competence of bovine oocytes after specific inhibition of MPF kinase activity: effect of estradiol supplementation and follicle size. Anim. Reprod. Sci. 92, 231–240. Caron-Beaudoin, E., Denison, M.S., Sanderson, J.T., 2016. Effects of neonicotinoids on promoter-specific expression and activity of aromatase (CYP19) in human adrenocortical carcinoma (H295R) and primary umbilical vein endothelial (HUVEC) cells. Toxicol. Sci. 149, 134–144. Caron-Beaudoin, E., Viau, R., Hudon-Thibeault, A.A., Vaillancourt, C., Sanderson, J.T., 2017. The use of a unique co-culture model of fetoplacental steroidogenesis as a screening tool for endocrine disruptors: the effects of neonicotinoids on aromatase activity and hormone production. Toxicol. Appl. Pharmacol. 332, 15–24. Cecchino, G.N., Seli, E., Alves da Motta, E.L., Garcia-Velasco, J.A., 2018. The role of mitochondrial activity in female fertility and assisted reproductive technologies: overview and current insights. Reprod. BioMed. Online 36, 686–697. Dalton, C.M., Carroll, J., 2013. Biased inheritance of mitochondria during asymmetric cell division in the mouse oocyte. J. Cell Sci. 126, 2955–2964. Dalton, C.M., Szabadkai, G., Carroll, J., 2014. Measurement of ATP in single oocytes: impact of maturation and cumulus cells on levels and consumption. J. Cell. Physiol. 229, 353–361. Driessens, N., Versteyhe, S., Ghaddhab, C., Burniat, A., De Deken, X., Van Sande, J., Dumont, J.E., Miot, F., Corvilain, B., 2009. Hydrogen peroxide induces DNA singleand double-strand breaks in thyroid cells and is therefore a potential mutagen for this organ. Endocr. Relat. Cancer 16, 845–856. El Okle, O.S., El Euony, O.I., Khafaga, A.F., Lebda, M.A., 2018. Thiamethoxam induced hepatotoxicity and pro-carcinogenicity in rabbits via motivation of oxidative stress, inflammation, and anti-apoptotic pathway. Environ. Sci. Pollut. Res. Int. 25, 4678–4689. Fedrizzi, G., Altafini, A., Armorini, S., Al-Qudah, K.M., Roncada, P., 2019. LC-MS/MS analysis of five neonicotinoid pesticides in sheep and cow Milk samples collected in Jordan Valley. Bull. Environ. Contam. Toxicol. 102, 347–352. Garcia-Ruiz, C., Fernandez-Checa, J.C., 2018. Mitochondrial oxidative stress and antioxidants balance in fatty liver disease. Hepatol Commun 2, 1425–1439. Girish, K.S., Paul, M., Thushara, R.M., Hemshekhar, M., Shanmuga Sundaram, M., Rangappa, K.S., Kemparaju, K., 2013. Melatonin elevates apoptosis in human
9
Toxicology in Vitro 61 (2019) 104635
Z.-W. Nie, et al.
Simon-Delso, N., Amaral-Rogers, V., Belzunces, L.P., Bonmatin, J.M., Chagnon, M., Downs, C., Furlan, L., Gibbons, D.W., Giorio, C., Girolami, V., Goulson, D., Kreutzweiser, D.P., Krupke, C.H., Liess, M., Long, E., McField, M., Mineau, P., Mitchell, E.A., Morrissey, C.A., Noome, D.A., Pisa, L., Settele, J., Stark, J.D., Tapparo, A., Van Dyck, H., Van Praagh, J., Van der Sluijs, J.P., Whitehorn, P.R., Wiemers, M., 2015. Systemic insecticides (neonicotinoids and fipronil): trends, uses, mode of action and metabolites. Environ. Sci. Pollut. Res. Int. 22, 5–34. St John, J., 2014. The control of mtDNA replication during differentiation and development. Biochim. Biophys. Acta 1840, 1345–1354. Stone, J.R., Yang, S., 2006. Hydrogen peroxide: a signaling messenger. Antioxid. Redox Signal. 8, 243–270. Swenson, T.L., Casida, J.E., 2013. Neonicotinoid formaldehyde generators: possible mechanism of mouse-specific hepatotoxicity/hepatocarcinogenicity of thiamethoxam. Toxicol. Lett. 216, 139–145. Taillebois, E., Cartereau, A., Jones, A.K., Thany, S.H., 2018. Neonicotinoid insecticides mode of action on insect nicotinic acetylcholine receptors using binding studies. Pestic. Biochem. Physiol. 151, 59–66. Telo, G.M., Senseman, S.A., Marchesan, E., Camargo, E.R., Jones, T., McCauley, G., 2015. Residues of thiamethoxam and chlorantraniliprole in rice grain. J. Agric. Food Chem. 63, 2119–2126. Tosti, E., Boni, R., Cuomo, A., 2000. Ca(2+) current activity decreases during meiotic progression in bovine oocytes. Am J Physiol Cell Physiol 279, C1795–C1800. Vercesi, A.E., Kowaltowski, A.J., Grijalba, M.T., Meinicke, A.R., Castilho, R.F., 1997. The role of reactive oxygen species in mitochondrial permeability transition. Biosci. Rep. 17, 43–52. Veysset, P., Lherm, M., Bebin, D., Roulenc, M., 2014. Mixed crop-livestock farming systems: a sustainable way to produce beef? Commercial farms results, questions and perspectives. Animal 8, 1218–1228. Wang, X., Anadon, A., Wu, Q., Qiao, F., Ares, I., Martinez-Larranaga, M.R., Yuan, Z., Martinez, M.A., 2018. Mechanism of neonicotinoid toxicity: impact on oxidative stress and metabolism. Annu. Rev. Pharmacol. Toxicol. 58, 471–507. Wehrend, A., Meinecke, B., 2001. Kinetics of meiotic progression, M-phase promoting factor (MPF) and mitogen-activated protein kinase (MAP kinase) activities during in vitro maturation of porcine and bovine oocytes: species specific differences in the length of the meiotic stages. Anim. Reprod. Sci. 66, 175–184. Wu, B., Ignotz, G., Currie, W.B., Yang, X., 1997. Dynamics of maturation-promoting factor and its constituent proteins during in vitro maturation of bovine oocytes. Biol. Reprod. 56, 253–259. Wuest, M., Perreault, A., Richter, S., Knight, J.C., Wuest, F., 2019. Targeting phosphatidylserine for radionuclide-based molecular imaging of apoptosis. Apoptosis 24, 221–244. Yan, S.H., Wang, J.H., Zhu, L.S., Chen, A.M., Wang, J., 2016. Thiamethoxam induces oxidative stress and antioxidant response in zebrafish (Danio Rerio) livers. Environ. Toxicol. 31, 2006–2015. Yang, H., Xie, Y., Yang, D., Ren, D., 2017. Oxidative stress-induced apoptosis in granulosa cells involves JNK, p53 and Puma. Oncotarget 8, 25310–25322. Yi, M., Rosin, M.P., Anderson, C.K., 1990. Response of fibroblast cultures from ataxiatelangiectasia patients to oxidative stress. Cancer Lett. 54, 43–50. Yuh, H.S., Yu, D.H., Shin, M.J., Kim, H.J., Bae, K.B., Lee, D.S., Lee, H.C., Chang, W.K., Park, S.B., Lee, S.G., Park, H.D., Ha, J.H., Hyun, B.H., Ryoo, Z.Y., 2010. The effects of various antioxidants on the development of parthenogenetic porcine embryos. In Vitro Cellular and Developmental Biology: Animal 46, 148–154.
platelets via ROS mediated mitochondrial damage. Biochem. Biophys. Res. Commun. 438, 198–204. Green, T., Toghill, A., Lee, R., Waechter, F., Weber, E., Noakes, J., 2005. Thiamethoxam induced mouse liver tumors and their relevance to humans. Part 1: mode of action studies in the mouse. Toxicol. Sci. 86, 36–47. Jeschke, P., Nauen, R., 2008. Neonicotinoids-from zero to hero in insecticide chemistry. Pest Manag. Sci. 64, 1084–1098. Jo, Y.J., Jang, W.I., Namgoong, S., Kim, N.H., 2015. Actin-capping proteins play essential roles in the asymmetric division of maturing mouse oocytes. J. Cell Sci. 128, 160–170. Kanka, J., Kepkova, K., Nemcova, L., 2009. Gene expression during minor genome activation in preimplantation bovine development. Theriogenology 72, 572–583. Kanka, J., Nemcova, L., Toralova, T., Vodickova-Kepkova, K., Vodicka, P., Jeseta, M., Machatkova, M., 2012. Association of the transcription profile of bovine oocytes and embryos with developmental potential. Anim. Reprod. Sci. 134, 29–35. Kitajima, T.S., Ohsugi, M., Ellenberg, J., 2011. Complete kinetochore tracking reveals error-prone homologous chromosome biorientation in mammalian oocytes. Cell 146, 568–581. Ledan, E., Polanski, Z., Terret, M.E., Maro, B., 2001. Meiotic maturation of the mouse oocyte requires an equilibrium between cyclin B synthesis and degradation. Dev. Biol. 232, 400–413. Lee, W., Lee, D.G., 2018. Reactive oxygen species modulate itraconazole-induced apoptosis via mitochondrial disruption in Candida albicans. Free Radic. Res. 52, 39–50. Li, M., Li, S., Yuan, J., Wang, Z.B., Sun, S.C., Schatten, H., Sun, Q.Y., 2009. Bub3 is a spindle assembly checkpoint protein regulating chromosome segregation during mouse oocyte meiosis. PLoS One 4, e7701. Li, Y., Liu, X., Jiang, D., Lin, Y., Wang, Y., Li, Q., Liu, L., Jin, Y.H., 2016. Betulin induces reactive oxygen species-dependent apoptosis in human gastric cancer SGC7901 cells. Arch. Pharm. Res. 39, 1257–1265. Lincoln, A.J., Wickramasinghe, D., Stein, P., Schultz, R.M., Palko, M.E., De Miguel, M.P., Tessarollo, L., Donovan, P.J., 2002. Cdc25b phosphatase is required for resumption of meiosis during oocyte maturation. Nat. Genet. 30, 446–449. Mayer, A., Baran, V., Sakakibara, Y., Brzakova, A., Ferencova, I., Motlik, J., Kitajima, T.S., Schultz, R.M., Solc, P., 2016. DNA damage response during mouse oocyte maturation. Cell Cycle 15, 546–558. Memili, E., First, N.L., 2000. Zygotic and embryonic gene expression in cow: a review of timing and mechanisms of early gene expression as compared with other species. Zygote 8, 87–96. Morado, S.A., Cetica, P.D., Beconi, M.T., Dalvit, G.C., 2009. Reactive oxygen species in bovine oocyte maturation in vitro. Reprod. Fertil. Dev. 21, 608–614. Nurse, P., 1997. Checkpoint pathways come of age. Cell 91, 865–867. Omata, C., Kato, T.A., 2019. In situ DNA damaging foci analysis on metaphase chromosomes. Methods Mol. Biol. 1984, 87–93. Perez, G.I., Acton, B.M., Jurisicova, A., Perkins, G.A., White, A., Brown, J., Trbovich, A.M., Kim, M.R., Fissore, R., Xu, J., Ahmady, A., D'Estaing, S.G., Li, H., Kagawa, W., Kurumizaka, H., Yokoyama, S., Okada, H., Mak, T.W., Ellisman, M.H., Casper, R.F., Tilly, J.L., 2007. Genetic variance modifies apoptosis susceptibility in mature oocytes via alterations in DNA repair capacity and mitochondrial ultrastructure. Cell Death Differ. 14, 524–533. Senyildiz, M., Kilinc, A., Ozden, S., 2018. Investigation of the genotoxic and cytotoxic effects of widely used neonicotinoid insecticides in HepG2 and SH-SY5Y cells. Toxicol. Ind. Health 34, 375–383.
10
Contents lists available at ScienceDirect
Toxicology in Vitro journal homepage: www.elsevier.com/locate/toxinvit
Thiamethoxam induces meiotic arrest and reduces the quality of oocytes in cattle ⁎
Zheng-Wen Niea, Ying-Jie Niua, Wenjun Zhoua, Ju-Yeon Kima, Sun A. Ockb, , Xiang-Shun Cuia, a b
T
⁎
Department of Animal Sciences, Chungbuk National University, Chungbuk, Cheongju 361-763, Republic of Korea Animal Biotechnology Division, National Institute of Animal Science, Rural Development Administration, Jeonju 55365, Republic of Korea
A R T I C LE I N FO
A B S T R A C T
Keywords: Thiamethoxam Bovine oocyte ROS Mitochondria Apoptosis
Thiamethoxam (TMX) is a neonicotinoid insecticide, the residues of which have been detected on various crops. In addition to its specific acetylcholine toxicity to insects, TMX was also found to be toxic to mammals. Moreover, oocytes are vulnerable to reactive oxygen species (ROS). Excessive ROS production can override antioxidant defenses and produce oxidative stress and DNA damage that trigger apoptosis and necrosis in organisms. In this study, we exposed bovine oocytes to TMX during maturation. Microscopic examination showed that 1.6 mM TMX significantly inhibited maturation at the germinal vesicle (GV) and metaphase I (MI) stages. Immunofluorescence staining and enzyme activity analysis revealed that TMX induced a reduction in CDC25 and CDC2 activity. Furthermore, time-lapse tracking and immunofluorescence staining indicated the maintenance of cyclin B in the cytoplasm, persistence of Bub3 at kinetochores, and absence of actin caps after TMX-exposed oocytes reached the MI stage. In addition, metaphase II (MII) oocytes exposed to TMX showed disordered chromosomes and spindles. These oocytes accumulated excess ROS and showed significantly decreased mitochondrial membrane potential and increased apoptotic signals. Parthenogenetic embryos from these oocytes showed decreased percentages of morulae and blastocysts. These results indicate that TMX delays bovine oocyte progression to MI stage, blocks them at the MI stage, triggers disordered chromosomes and spindles at MII stage, and ultimately results in MII oocytes with poor cleavage ability and inhibited development to morulae and blastocysts.
1. Introduction Neonicotinoid insecticides are pesticides widely used on agriculture crops that mediate their toxic effects by binding to nicotinic acetylcholine receptors (nAChRs). These insecticides have a high affinity for insect nAChRs and low toxicity to mammals, lacking teratogenic and mutagenic effects, and therefore can be extensively applied (Jeschke and Nauen, 2008). Imidacloprid, thiamethoxam (TMX), thiacloprid (THI), nitenpyram, acetamiprid, and clothianidin (CLO) are currently the most commonly used neonicotinoids (Simon-Delso et al., 2015). Among these neonicotinoids, TMX has highest IC50 for insect and vertebrate α4β2 receptors (Taillebois et al., 2018). This indicates that TMX has a relatively low toxicity to non-target insects and is safer for vertebrates compared to other neonicotinoids. However, it is very likely that this characteristic of TMX will result in its excessive use, which could pose a danger to mammals. TMX residue has been found in sheep and cow milk when TMX was used widely (Fedrizzi et al., 2019). Besides, a previous study indicated that among neonicotinoids, TMX had
⁎
the highest cytotoxicity to SH-SY5Y and HepG2 cells and TMX and THI more readily induced DNA damage in HepG2 cells compared to that with other neonicotinoids (Senyildiz et al., 2018). With regards to cytotoxicity, TMX produces metabolites such as CLO, desmethyl-CLO, desmethyl-TMX, and formaldehyde, which are harmful to cell metabolism These products are generated through catalysis or the inhibition of metabolic enzymes (Wang et al., 2018). For example, formaldehyde derived from TMX, dmTMX, or CLO results in mouse-specific hepatotoxicity/hepatocarcinogenicity (Swenson and Casida, 2013). In terms of DNA damage, TMX can also exert other toxic effects via different molecular mechanisms. Neonicotinoids can perturb mitochondrial function and induce increased levels of cytochrome P450 (CYP450) and aldehyde oxidase (AOX), resulting in excess ROS production as a consequence of mitochondrial dysfunction. Excess ROS can in turn induce oxidative stress, which might result in the oxidation of lipids, proteins, and DNA (Wang et al., 2018). Meanwhile, TMX inhibits normal apoptosis in the liver of rabbits (El Okle et al., 2018). In addition, TMX has a tertiary and specific effect on cells. Using co-cultures of
Corresponding authors. E-mail addresses: [email protected] (S.A. Ock), [email protected] (X.-S. Cui).
https://doi.org/10.1016/j.tiv.2019.104635 Received 8 July 2019; Received in revised form 18 August 2019; Accepted 29 August 2019 Available online 31 August 2019 0887-2333/ © 2019 Published by Elsevier Ltd.
Toxicology in Vitro 61 (2019) 104635
Z.-W. Nie, et al.
(caption on next page)
2
Toxicology in Vitro 61 (2019) 104635
Z.-W. Nie, et al.
Fig. 1. Effect of thiamethoxam (TMX) toxicity on bovine oocyte maturation. Maturation rates of bovine oocytes exposed to different concentrations of TMX (A) showed that 1.6 mM TMX significantly decreased the rate of oocyte maturation. Spindles and chromosomes at different stages in the development of oocytes treated with or without TMX are shown (B). Oocytes showed a spindle ball and disordered chromosomes at the germinal vesicle breakdown (GVBD) stage, a bipolar spindle and chromosomes in a belt at the pre-metaphase I (MI) stage, or a bipolar spindle and aligned chromosomes at the pre-MI stage. Metaphase II (MII) oocytes had two parts of spindles and chromosomes, which belonged to the secondary oocyte and first polar body, respectively. Normal secondary oocytes had the same spindle and chromosomes as MI oocytes, and abnormal secondary oocytes had disordered or cohesive spindles or chromosomes. Portions of GVBD, MI, and MII oocytes after different durations of in vitro maturation (C) and changes in the percentages of oocytes at the GVBD, MI, and MII stages during maturation (D), showing delayed or arrested stages. Percentages of oocytes with abnormal spindles and chromosomes (E) showing increased rates of abnormal MII oocytes in the TMX group. The scale bar in the merged figures = 30 μm. The scale bar in enlarged figures = 8 μm. “n” represents the total number of oocytes in each stage.
mL epidermal growth factor (E4127; Sigma), 100 IU/mL penicillin/ streptomycin, 22 μg/mL sodium pyruvate (P5280; Sigma), and 95 μg/ mL L-cysteine (C7352; Sigma) at 38.5 °C in a humidified atmosphere of 5% CO2 and 95% air.
H295R and BeWo cells, it was shown that TMX induced CYP3A7 (cytochrome P450 family 3 subfamily A member 7) expression, which enhanced the activity of aromatase, in turn resulting in increased synthesis of estradiol (Caron-Beaudoin et al., 2017). Similarly, research has indicated that TMX induces PII- and I.3-mediated CYP19 expression and aromatase activity (Caron-Beaudoin et al., 2016). MtDNA and copy number of mitochondria increase during oocyte maturation (Cecchino et al., 2018), and their number in matured oocytes is high (St John, 2014). Oocyte maturation is associated with high levels of mitochondrial activity, during which, the mitochondria concentrate around the spindle and are involved in chromosome separation (Dalton and Carroll, 2013). During polar body extrusion, the levels of ATP generated by mitochondria peak (Dalton et al., 2014). In addition, mitochondria are an important source of ROS owing to the production of superoxide by the respiratory chain (Garcia-Ruiz and FernandezCheca, 2018). In normal cells, peroxide can be removed; however, for oocytes with dysfunctional mitochondria or unbalanced redox reactions, ROS are generated, which has a detrimental effect on oocyte maturation. In this regard, it has previously been found that bovine oocytes extracted from ovaries at the germinal vesicle (GV) stage initially have relatively high levels of ROS, which gradually decrease during the first 4 h of in vitro maturation, and are then maintained at a relatively low level in the subsequent stages of maturation (Morado et al., 2009). This suggests that bovine oocytes have anti-oxidative capacity and that certain levels of ROS are necessary for meiosis to proceed in oocytes. Despite the residual TMX found on crops, of which of stalks are usually used to feed cattle (Telo et al., 2015; Veysset et al., 2014), the presence excess ROS derived from TMX and the specific anti-oxidative capacity of bovine oocytes, and specifically whether excess ROS from TMX has toxicity to bovine oocytes, are unknown. In the present study, we hypothesized that TMX induces the excess production of ROS and mitochondrial dysfunction. ROS-triggered peroxidation and defects in mitochondrial energy generation would then damage biomolecules involved in maturation, blocking bovine oocytes at the MI (metaphase I) stage, trapping them at the anaphase I (AI)/ telophase I (TI) stage, and inducing chromosome and spindle disorder at the metaphase II (MII) stage. Additionally, the maternal-derived materials required for embryo development in oocytes would also be disturbed. Consequently, these MII oocytes with poor potential for embryo development would lose their ability to undergo cleavage and develop into morulae and blastocysts. Collectively, the damage caused by TMX would finally result in apoptosis.
2.2. Parthenogenetic activation and in vitro embryo culture After maturation for 24 h, cumulus cells surrounding oocytes were removed by dispersion with 0.1% hyaluronidase and repeated pipetting for approximately 1 min. For parthenogenetic activation, oocytes with first polar bodies were selected, incubated in CR1-aa (6.7 mg/mL NaCl (S5886; Sigma), 0.23 mg/mL KCl (P5405; Sigma), 2.2 mg/mL NaHCO3 (S5761; Sigma), 0.55 mg/mL hemicalcium lactate (L4388; Sigma), 1% MEM non-essential amino acids (M7145), 2% BMEM essential amino acids (B6766)) supplemented with 3 mg/mL BSA (A8806; Sigma) and 5 mM ionomycin (I0634; Sigma) for 7 min, and then in CR1-aa supplemented with 2 mM 6-dimethylaminopurine (D2629; Sigma) and 7.5 mg/mL cytochalasin B (C6762; Sigma) for 3 h. The oocytes were then washed with IVC-1 (CR1-aa supplemented with 10% BSA, 0.31 mg/mL glutathione (G6013; Sigma), 100 IU/mL penicillin/streptomycin, 44 μg/mL sodium pyruvate, and 0.146 mg/mL glutamine), cultured in IVC-1 for 4 d and then in IVC-2 (IVC-1 in which 10% BSA was replaced with 10% fetal calf serum) for 3 d in a humidified atmosphere of 5% CO2 and 95% air. 2.3. Immunofluorescence staining and confocal microscopy Oocytes were fixed in 3.7% paraformaldehyde for 1 h at room temperature, washed three times with phosphate-buffered saline/ polyvinyl alcohol (PBS/PVA), permeabilized with PBS/PVA containing 1.0% triton X-100 at room temperature for 1 h, and washed three times with PBS/PVA. The oocytes were then blocked in PVA-PBS containing 1.0% bovine serum albumin at room temperature for 1 h and subsequently incubated overnight at 4 °C with an anti-α-tubulin-FITC antibody (1:100; Sigma; USA), anti-γH2AX antibody (1:100, pS139; Cell Signaling Technology, Danvers, MA, USA), anti-CDC25C antibody (1:100, sc-5620; Santa Cruz Biotechnology, Dallas, TX, USA), anti-CDC2 antibody (1:100, pT161; Cell Signaling; USA), anti-actin antibody (1:100, sc-1616; Santa Cruz Biotechnology), or an anti-Bub3 antibody (1:50, sc-28,258; Santa Cruz Biotechnology). After incubation with the primary antibody (except for anti-α-tubulin-FITC antibody), the oocytes were washed three times with PBS/PVA, incubated at 37 °C for 1 h with a secondary IgG antibody, stained with Hoechst 33342 (bisBenzimide H33342 trihydrochloride, 1:2000; Sigma Life Science) for 15 min, washed three times in PBS/PVA, mounted on slides, and examined under a confocal microscope (Zeiss LSM 710 META; Jena, Germany). Images were processed using Zen software (version 8.0, Zeiss).
2. Materials and methods 2.1. Oocyte collection and in vitro maturation Ovaries from female Hanwoo cattle were obtained from a local slaughterhouse, maintained in saline at 38.5 °C, and transported to the laboratory. Cumulus oocyte complexes were extracted from follicles using an injection syringe, washed three times in Tyrode's lactate-4-(2hydroxyethyl)-1-piperazineethanesulfonic acid, and cultured in tissue culture medium 199 (TCM 199) supplemented with 10% fetal calf serum (10082–147; Gibco, USA), 1 μg/mL β-estradiol (E2758; Sigma, USA), 10 μg/mL follicle-stimulating hormone (F2293; Sigma), 10 ng/
2.4. ROS staining The ROS contents of the treated and untreated oocytes were measured after oocytes had been cultured for 24 h. ROS content was quantified using the dichlorodihydrofluorescein diacetate (DCHFDA, Molecular Probes, Invitrogen, Carlsbad, CA, USA) method, as described previously (Yi et al., 1990). Briefly, the cumulus cells were removed from cumulus oocyte complexes with hyaluronidase. Denuded oocytes 3
Toxicology in Vitro 61 (2019) 104635
Z.-W. Nie, et al.
Fig. 2. Molecular regulation from germinal vesicle breakdown (GVBD) to metaphase I (MI) stages. Immunofluorescence staining for CDC25 at the GV and GVBD stages (A) and relative intensity of CDC25 fluorescence (B), showing higher CDC25 activity in the thiamethoxam (TMX) group. Immunofluorescence staining of CDC2 at the GV, pre-MI, and MI stages (C), relative intensity of CDC2 fluorescence (D), and relative activity of CDC2 extracted from oocytes (E), showing lower CDC2 activity in the TMX group between 8 and 12 h. The scale bar in the merged figures = 30 μm. The scale bar in enlarged figures = 8 μm. “n” represents the total number of oocytes in each stage.
2.5. Analysis of mitochondrial membrane potential
were exposed to DCHFDA diluted in PBS/PVA (1:1000) for 15 min and then washed three times with PBS/PVA. Live imaging and quantitation were conducted using a fluorescence microscope (Nikon, Tokyo, Japan) and Photoshop (CS2; Adobe, San Jose, CA, USA). Then Image J software was used to filter out the contaminated spots with strong intensity and calculate the ROS signal of oocytes.
To measure mitochondrial membrane potential, denuded oocytes were washed three times with PBS/PVA and incubated with 5,5′,6,6′tetrachloro-1,1′,3,3′-tetraethyl-imidacarbocyanine iodide (JC-1) (Invitrogen) diluted in culture medium (1:200) for 30 min. The membrane potential was calculated as the ratio of red florescence, 4
Toxicology in Vitro 61 (2019) 104635
Z.-W. Nie, et al.
Fig. 3. Molecular regulation from metaphase I (MI) to anaphase I (AI) stages. Time-lapse of H2B-Cherry and cyclin B-eGFP (A) and relative intensity of cyclin B-eGFP fluorescence (C), showing the maintenance of cyclin B1 from 12 to 18 h (Scale bar = 60 μm). The extrusion rate of polar bodies in the thiamethoxam (TMX) group at 18 h (B) was decreased significantly. Immunofluorescence staining of actin (D) and percentage of oocytes with actin caps (F) at 18 h, showing that actin caps could not be formed in the TMX group (Scale bar = 30 μm). Immunofluorescent staining for BUB3 (E) and percentage of oocytes with BUB3 puncta at 18 h (G), showing the maintenance of puncta in the TMX group (Scale bar = 4 μm). “n” represents the total number of oocytes in each stage.
(50 mM tris-HCl (pH 7.5), 0.5 M NaCl, 5 mM EDTA, 2 mM EGTA, 0.01% Brij-35 (v/v), 1 mM PMSF, 0.05 mg/mL leupeptin, 50 mM 2-mercaptoethanol, 25 mM β-glycerophosphate, and 1 mM sodium orthovanadate) and stored at −80 °C. Sample extracts were quantified by ELISA using a MESACUP cdc2 Kinase Assay Kit (MBL International, Nagoya, Japan), and the optical density of each well was read at 492 nm using a microplate reader.
corresponding to activated mitochondria (J-aggregates), to green fluorescence, corresponding to less-activated mitochondria (J-monomers). Fluorescence was visualized using a Zeiss inverted confocal microscope equipped with a × 40 oil immersion objective (Zeiss, Jena, Germany). Images were processed using ZEN software (Zen Software, Manchester, UK). 2.6. Detection of early apoptosis
2.8. Statistical analysis
Apoptosis was detected using an Annexin V-FITC-PI Apoptosis Detection kit (Vazyme, A211-01) according to the manufacturer's instructions. Briefly, 20 oocytes were washed in PBS/PVA and then incubated in 100 mL binding buffer containing 5 mL of Annexin V-FITC for 10 min in the dark and washed twice in PBS/PVA. The green FITC signal was detected using a fluorescent microscope with 450–490 nm (excitation) and 520 nm (emission) filters.
All results were subjected to a one-way analysis of variance, and differences between the treatment groups were assessed using a least significant difference test in the Statistical Package for Social Sciences (SPSS) software. Each experiment was conducted at least three times, and differences were considered significant when P < .05 or highly significant when P < .01. The relative value of each repeat in the control group was set to 1, and the values of the treatment group were standardized to those of the control.
2.7. In vitro p34cdc2 kinase assay Thirty oocytes, as one sample, were lysed with 5 μL of sample buffer 5
Toxicology in Vitro 61 (2019) 104635
Z.-W. Nie, et al.
Fig. 4. Reactive oxygen species (ROS), mitochondrial membrane potential (MMP), and apoptosis induced by thiamethoxam (TMX) at 24 h. ROS staining (A) and relative levels of ROS (B), showing increased ROS in the TMX group (Scale bar = 120 μm). JC-1 staining (C) and relative intensity of red to green staining (D), showing decreased MMP in the TMX group (Scale bar = 120 μm). Immunofluorescence staining for γH2AX (E) and relative intensity of fluorescence (F), showing no significant difference in γH2AX between the TMX group and control groups (Scale bar = 30 μm). AnnexinV staining (G) and relative intensity (H), showing early apoptosis in the TMX group (Scale bar = 120 μm). “n” represents the total number of oocytes in each stage. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)
6
Toxicology in Vitro 61 (2019) 104635
Z.-W. Nie, et al.
Fig. 5. Development of embryos derived from oocytes treated with or without thiamethoxam (TMX). Embryos derived from oocytes treated with or without TMX at day 7 (A) and percentages at each stage (B), showing arrest before the 8-cell stage. Scale bar = 240 μm. “MO” and “BL”, respectively, represent morula and blastocyst stage. “n” represents the total number of oocytes in each stage.
3. Results 3.1. TMX affects bovine oocyte maturation To determine the toxicity of TMX during bovine oocyte maturation, we examined the effects of a concentration gradient of this agent according to its lowest effective dose in previous study (Green et al., 2005). GV oocytes were cultured and exposed to various concentrations of TMX for 24 h to determine the appropriate concentration of TMX for subsequent analyses. Compared to 0 mM TMX, 1.6 mM TMX significantly reduced the maturation rate of oocytes (65.27% ± 3.57% vs. 51.68% ± 4.70%; P < .05) and that 3.2 mM TMX resulted in a highly significant reduction (65.27% ± 3.57% vs. 39.67% ± 4.74%; P < .01; Fig. 1A). Therefore, we established 1.6 mM TMX as an appropriate dose to study the effect of TMX on oocyte maturation. To further study the process of bovine oocyte maturation, oocytes cultured for 0, 8, 12, and 24 h, which corresponds to the GV, germinal vesicle breakdown (GVBD), MI, and MII stages of maturation, respectively (Tosti et al., 2000), were exposed to 1.6 mM TMX and subsequently analyzed by immunofluorescence staining. The characteristics of oocytes at each stage were determined by tracking spindles and chromosomes, as shown in Fig. 1B. Oocytes contained a nucleus at the GV stage, a ball of microtubes and scattered chromosomes at the beginning of the GVBD stage, a dipolar spindle and aligned chromosomes at the MI stage, and two sets of spindles and chromosomes at the MII stage. The pattern of maturation was altered when oocytes were exposed to TMX (Fig. 1C). For the convenience of analysis, oocytes at each stage were individually tracked during the course of culture (Fig. 1D and Table S1). At 8 h, the proportion of GVBD oocytes was highly significantly decreased compared to that in the control group (84.00% ± 2.40% vs. 24.90% ± 3.02%; P < .01) but recovered to
Fig. 6. Model of the mechanism to explain how thiamethoxam (TMX) affects bovine oocyte maturation. During bovine oocyte maturation, TMX induces excess accumulation of reactive oxygen species (ROS), a decrease in mitochondrial membrane potential (MMP), and apoptosis. These changes affect the maturation and developmental potential of bovine oocytes, reducing the level of active CDC25, and subsequently retarding the phosphorylation of CDC2. Consequently, there is a delay in the progression of oocytes to the germinal vesicle breakdown stage. After this stage, the TMX-induced effects cause damage including inhibition of the inactivation of the spindle assembly checkpoint (SAC), suppression of cyclin B degradation, and reduced formation of actin caps, which are essential for oocytes to successfully enter the AI stage. Exposure to TMX also diminishes the quality of metaphase II oocytes due to chromosomal abnormalities, which reduce the ability of oocytes to develop into embryos at all stages.
7
Toxicology in Vitro 61 (2019) 104635
Z.-W. Nie, et al.
3.4. TMX induces increases in ROS and apoptosis
control levels at 12 h. The proportion of MI oocytes in the TMX group was significantly lower than that in the control group only at 12 h (63.51% ± 6.71% vs. 39.50 ± 9.28%; P < .01), whereas the percentage of MII oocytes exposed to TMX decreased only at 24 h. At the long pre-MI stage, it was difficult to quantify the proportions based on two-dimensional images because of the specific three-dimensional structure of chromosomes in the cricoid metaphase band at this long stage (Kitajima et al., 2011). However, the oocytes at both GVBD (including all stages except the GV stage) and the MII stage showed no significant changes. Based on these observations, we concluded that TMX blocked oocytes before the MI stage. Similarly, at 24 h, TMX blocked oocytes prior to the AI/TI stage. We subsequently selected all MII oocytes and examined them for abnormal spindle and chromosome development. Compared to that in the control group, the percentage of abnormal MII oocytes, in which the spindle and chromosomes were disordered, increased in the TMX group during the maturation process (14.27 ± 10.39% vs. 31.89 ± 11.75%; P < .01); those MII oocytes with cohesive chromosomes (6.50 ± 1.80%) only appeared in the TMX group, but the rate was not significantly different from that in the control group (Fig. 1B and E).
We initially detected ROS using DCHFDA at 24 h. The average relative intensity of the ROS signal in the TMX group was significantly higher than that in the control group (1 vs. 1.33 ± 0.07; P < .05; Fig. 4A and B). A cellular imbalance of redox reactions is one factor contributing to the production of ROS, and this process is related to mitochondrial dysfunction. We therefore examined mitochondrial membrane potential (MMP), which revealed that the MMP of oocytes exposed to TMX was significantly decreased (1% vs. 0.81% ± 0.04%; P < .05; Fig. 4C and D). We subsequently evaluated the damage induced by ROS. Previous studies (Perez et al., 2007; Yuh et al., 2010) have indicated the possibility that ROS production induced by TMX triggers DNA damage. Therefore, to assess potential DNA damage, we selected γH2AX as a marker to stain for (Mayer et al., 2016; Omata and Kato, 2019). However, we observed no significant difference in γH2AX between the control and TMX groups (1 vs. 1.20 ± 0.21; P > .05; Fig. 4E and F), indicating that exposure to TMX does not result in DNA damage. Nevertheless, the higher intensity of the Annexin-V signals in the membrane of TMX-treated oocytes than that in the control indicated that TMX does induce early apoptosis at 24 h (2.33 ± 1.45 vs. 24.88 ± 4.99; P < .05; Fig. 4G and H).
3.2. TMX delays the activation of maturation promoting factor before the MI stage
3.5. TMX results in poor early embryonic development To assess the potential for embryonic development, we evaluated parthenogenetic embryos at each stage. We accordingly found that the percentages of TMX-treated embryos at 1-cell and 2–8-cell stages were significantly increased compared to those in respective controls (Fig. 5A and B, Table S2). In contrast, we observed significant decreases at the morula and blastocyst stage.
To determine how oocytes are arrested, we performed immunofluorescence staining for active CDC25 and CDK2. We accordingly found that oocytes in the TMX group had higher CDC25 activity compared to that in the control group at 12 h (1 vs. 1.97 ± 0.21; P < .05; Fig. 2A and B). Moreover, as described in previous studies, we found that the phosphorylation of CDC2 decreased from 8 to 12 h in the control group; however, the phosphorylation showed a marginal increase in the TMX group and was invariably lower (8 h: 1 vs. 0.71 ± 0.05; P < .05; 12 h: 0.90 ± 0.13 vs. 0.74 ± 0.10; P < .05; Fig. 2C and D). This suggested increased activity of CDC2 in the control group and decreased activity in the TMX group. Here, the specificity of the P-CDC2 antibody was tested by western blotting and a single band for P-CDC2 indicated the accuracy of assessing P-CDC2 in Fig. 2C and D (Fig. 2E). Additionally, we assessed substrate catalysis and the results showed that enzymatic activities decreased significantly compared to those in the control group at 8 h (1 vs. 0.71 ± 0.03; P < .05; Fig. 2F), but were not significantly changed at 12 h (1 vs. 0.99 ± 0.10; P > .05; Fig. 2F).
4. Discussion Neonicotinoid including TMX has anti-reproductive cytotoxic effects on animals and increases ROS or RNS formation in vivo (Wang et al., 2018). Similarly, it has previously been shown that TMX induces oxidative stress and an antioxidant response in the zebrafish liver (Yan et al., 2016). Consistent with these previous studies, we found that TMX induces high levels of ROS, as determined by DCHFDA levels. Only TMX at a concentration of > 1.6 mM could disturb bovine oocyte maturation, which indicates that oocytes have an effective antioxidant control system. The inhibition of GVBD induced by TMX could be overcome at 12 h; however, thereafter, the arrest between the MI and MII stages could not be recovered. In normal in vitro mature cultures, bovine oocytes show a rapid reduction in ROS, which are subsequently maintained at low levels until 12 h, after which the levels gradually increase (Morado et al., 2009). This indicates that bovine oocytes have effective antioxidant capacity prior to the MI stage, which subsequently wanes. ROS and mitochondrial dysfunction generally occur concomitantly. Peroxide can perturb mitochondrial function and trigger a transition in mitochondrial permeability (Vercesi et al., 1997). In addition, although ROS generated by mitochondria can be removed by antioxidative enzymes, mitochondrial dysfunction such as the inhibition of complex I and a high NADH:NAD+ ratio results in the accumulation of excess ROS, which is damaging to cells (Adam-Vizi and Chinopoulos, 2006). An important change observed in dysfunctional mitochondria is a decrease in MMP. The mechanism underlying the adverse effects of neonicotinoids has been evaluated and it was suggested that these chemicals can disrupt mitochondrial function and that impaired mitochondria consequently produce increased amounts of ROS (Wang et al., 2018). Indeed, in the present study, we also demonstrated the association of low MMP and excess ROS. These changes can promote cell apoptosis, which has been observed in many previous studies (Girish et al., 2013; Li et al., 2016; Yang et al., 2017). During the early phase of apoptosis, no apoptotic bodies form; however, cells in this phase exhibit exposed
3.3. TMX arrests oocytes at the MI stage To determine whether some oocytes were arrested between the MI and MII stages when exposed to TMX, we tracked chromosomes and cyclin B by time-lapse from 12 to 18 h. As shown in Fig. 3A–C, whereas oocytes in the control group developed a polar body, those exposed to TMX did not, as determined by the pole body listing percentages at 18 h (58.12% ± 5.06% vs. 52.98% ± 6.10%; P < .05; Fig. 3B). Consistent with this observation, in the control group, the cyclin B1-EGFP signal gradually diminished during pole body listing, whereas in the TMX group, the signal was maintained (Fig. 3A–C, Movies S1 and S2). The progression of oocytes from the AI/TI stage not only requires degradation of cyclin B1 but also the coordination of cytoplasmic and nuclear division. To examine these processes, staining for actin and Bub3 was performed at 18 h, and the results showed that compared to those in the control group, actin caps did not form in oocytes exposed to TMX (23.74% ± 2.58% vs. 15.22% ± 3.30%; P < .05; Fig. 3D and E). In addition, in contrast to that observed in the control group, the mottled signal for Bub3 in the TMX group indicated that the spindle assembly checkpoint was still active (Fig. 3F and G).
8
Toxicology in Vitro 61 (2019) 104635
Z.-W. Nie, et al.
Given the marked decrease in the proportion of morulae observed in the TMX group, these materials are assumed to be vital for major genome activation.
phosphatidylserine on the outer surface of the plasma membrane, to which Annexin-V can bind (Wuest et al., 2019). ROS can trigger apoptosis by disrupting mitochondria (Lee and Lee, 2018). Consistently, in the present study, we detected positive Annexin-V signals only in the membranes of oocytes exposed to TMX. However, although it has previously been found that increased oxidative stress can result in DNA damage (Driessens et al., 2009; Stone and Yang, 2006), our results indicate that ROS induced by TMX does not trigger DNA damage in bovine oocytes. Given that there have been no previous reports of the detection of γH2AX (a marker of DNA damage) in bovine oocytes, DNA damage is probably an infrequent event in these oocytes. Events during oocyte maturation are controlled by a series of proteins, among which the maturation promoting factor (MPF), consisting of CDC2 and cyclin B, regulates meiosis. Although the timing of events during bovine oocyte maturation might differ depending upon the medium in which they are cultured, the activity of MPF shows a significant increase at the GVBD stage, peaks at the MI stage, decreases at the AI stage, and subsequently increases at the MII stage (Wehrend and Meinecke, 2001; Wu et al., 1997). When MPF activity is inhibited, bovine oocytes become arrested at the GV stage (Beker-van Woudenberg et al., 2006). Cdc25 dephosphorylates and activates cdc2 kinase at Thr14 and Tyr15 (Nurse, 1997). In addition, it has been found that Cdc25C, in conjunction with Zn2+, dephosphorylates its substrate MPF/cdk1 in Xenopus oocytes (Lincoln et al., 2002). Accordingly, it can be reasoned that the delay in GVBD observed in the present study is due to the inactivation of CDC25C and the resulting inactivation of cdc2 kinase. In addition, degradation of cyclin B is required for the transition from the MI to the MII stage. It is not cyclin B2 but cyclin B1 that is responsible for progression from the MI stage and entry into the AI stage in mouse oocytes (Ledan et al., 2001). Consistently, we found that cyclin B1 was degraded in normal bovine oocytes. However, this process was inhibited by a significant degree in the TMX group. This suggests that TMX results in failed events related to the AI stage. At the MI stage, the spindle assembly checkpoint (SAC) in the active state can ensure the correct alignment of chromosomes and all the requisite connections of kinetochores with microtubules. Inactive SAC triggers the anaphase-promoting complex/cyclosome (APC/C)–CDC20-dependent degradation of cyclin B1. In this context, it has previously been shown that disturbances in Bub3, a member of the SAC, result in the incorrect attachment of microtubules to kinetochores, misaligned chromosomes, abnormal polar bodies, and aneuploidy (Li et al., 2009). Furthermore, it has been shown that the actin cap, which is under the control of actin-capping proteins, plays a role in extrusion of the first polar body during the maturation of mouse oocytes (Jo et al., 2015). Therefore, exit from the MI stage is controlled by multiple molecular mechanisms. Consistent with the aforementioned findings, the results obtained in the present study indicate that TMX-induced ROS blocks the inactivation of SAC and formation of an actin cap. Oocytes store mRNAs and proteins such as cyclin A2 and histone deacetylases for early embryonic development (Kanka et al., 2012). During bovine preimplantation development, embryos gradually lose the control exerted by maternal materials and show increasing expression of zygotic genes (Kanka et al., 2009). The commencement of major genome activation in bovine embryos is assumed to occur at the 8-cell stage because the expression pattern of genes at this stage is significantly different from that at the previous stage (Kanka et al., 2009; Memili and First, 2000). Consequently, the development of bovine embryos prior to the 8-cell stage mainly depends on the influence of maternal materials from oocytes. In the present study, we observed that embryos at all stages sustained damage resulting from the exposure of oocytes to TMX. It is noteworthy that blocks in embryo development gradually increased prior to the morula stage, whereas thereafter the differences between the TMX and control groups declined. Considering aforementioned points, we can conclude that TMX-induced ROS might destroy materials stored in oocytes that are required for further embryonic development, which results in arrest prior to the 8-cell stage.
5. Conclusion TMX exerts multiple effects on bovine oocyte maturation and quality. It reduces the level of active CDC25 and subsequently blocks the phosphorylation of CDC2. As a result, the development of bovine oocytes is arrested at the GVBD stage. However, progression of the maturation process can be recovered by certain repair mechanisms. In addition, this results in the maintenance of SAC activity at the MI stage or the inhibition of cyclin B degradation and the formation of actin caps. Consequently, oocytes are arrested at the MI stage or fail to enter the AI stage. However, even those oocytes that do progress from the MI stage and develop to the MII stage show evidence of TMX-induced damage, including abnormal chromosomes, apoptosis, the accumulation of ROS, decreased MMP, and poor embryonic developmental potential (Fig. 6). Supplementary data to this article can be found online at https:// doi.org/10.1016/j.tiv.2019.104635. Declaration of Competing Interest The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper. Acknowledgments This research was supported by the Basic Science Research Program through the National Research Foundation of Korea (NRF) funded by the Ministry of Education (No. 2018R1A2B6001173), Republic of Korea. References Adam-Vizi, V., Chinopoulos, C., 2006. Bioenergetics and the formation of mitochondrial reactive oxygen species. Trends Pharmacol. Sci. 27, 639–645. Beker-van Woudenberg, A.R., Zeinstra, E.C., Roelen, B.A., Colenbrander, B., Bevers, M.M., 2006. Developmental competence of bovine oocytes after specific inhibition of MPF kinase activity: effect of estradiol supplementation and follicle size. Anim. Reprod. Sci. 92, 231–240. Caron-Beaudoin, E., Denison, M.S., Sanderson, J.T., 2016. Effects of neonicotinoids on promoter-specific expression and activity of aromatase (CYP19) in human adrenocortical carcinoma (H295R) and primary umbilical vein endothelial (HUVEC) cells. Toxicol. Sci. 149, 134–144. Caron-Beaudoin, E., Viau, R., Hudon-Thibeault, A.A., Vaillancourt, C., Sanderson, J.T., 2017. The use of a unique co-culture model of fetoplacental steroidogenesis as a screening tool for endocrine disruptors: the effects of neonicotinoids on aromatase activity and hormone production. Toxicol. Appl. Pharmacol. 332, 15–24. Cecchino, G.N., Seli, E., Alves da Motta, E.L., Garcia-Velasco, J.A., 2018. The role of mitochondrial activity in female fertility and assisted reproductive technologies: overview and current insights. Reprod. BioMed. Online 36, 686–697. Dalton, C.M., Carroll, J., 2013. Biased inheritance of mitochondria during asymmetric cell division in the mouse oocyte. J. Cell Sci. 126, 2955–2964. Dalton, C.M., Szabadkai, G., Carroll, J., 2014. Measurement of ATP in single oocytes: impact of maturation and cumulus cells on levels and consumption. J. Cell. Physiol. 229, 353–361. Driessens, N., Versteyhe, S., Ghaddhab, C., Burniat, A., De Deken, X., Van Sande, J., Dumont, J.E., Miot, F., Corvilain, B., 2009. Hydrogen peroxide induces DNA singleand double-strand breaks in thyroid cells and is therefore a potential mutagen for this organ. Endocr. Relat. Cancer 16, 845–856. El Okle, O.S., El Euony, O.I., Khafaga, A.F., Lebda, M.A., 2018. Thiamethoxam induced hepatotoxicity and pro-carcinogenicity in rabbits via motivation of oxidative stress, inflammation, and anti-apoptotic pathway. Environ. Sci. Pollut. Res. Int. 25, 4678–4689. Fedrizzi, G., Altafini, A., Armorini, S., Al-Qudah, K.M., Roncada, P., 2019. LC-MS/MS analysis of five neonicotinoid pesticides in sheep and cow Milk samples collected in Jordan Valley. Bull. Environ. Contam. Toxicol. 102, 347–352. Garcia-Ruiz, C., Fernandez-Checa, J.C., 2018. Mitochondrial oxidative stress and antioxidants balance in fatty liver disease. Hepatol Commun 2, 1425–1439. Girish, K.S., Paul, M., Thushara, R.M., Hemshekhar, M., Shanmuga Sundaram, M., Rangappa, K.S., Kemparaju, K., 2013. Melatonin elevates apoptosis in human
9
Toxicology in Vitro 61 (2019) 104635
Z.-W. Nie, et al.
Simon-Delso, N., Amaral-Rogers, V., Belzunces, L.P., Bonmatin, J.M., Chagnon, M., Downs, C., Furlan, L., Gibbons, D.W., Giorio, C., Girolami, V., Goulson, D., Kreutzweiser, D.P., Krupke, C.H., Liess, M., Long, E., McField, M., Mineau, P., Mitchell, E.A., Morrissey, C.A., Noome, D.A., Pisa, L., Settele, J., Stark, J.D., Tapparo, A., Van Dyck, H., Van Praagh, J., Van der Sluijs, J.P., Whitehorn, P.R., Wiemers, M., 2015. Systemic insecticides (neonicotinoids and fipronil): trends, uses, mode of action and metabolites. Environ. Sci. Pollut. Res. Int. 22, 5–34. St John, J., 2014. The control of mtDNA replication during differentiation and development. Biochim. Biophys. Acta 1840, 1345–1354. Stone, J.R., Yang, S., 2006. Hydrogen peroxide: a signaling messenger. Antioxid. Redox Signal. 8, 243–270. Swenson, T.L., Casida, J.E., 2013. Neonicotinoid formaldehyde generators: possible mechanism of mouse-specific hepatotoxicity/hepatocarcinogenicity of thiamethoxam. Toxicol. Lett. 216, 139–145. Taillebois, E., Cartereau, A., Jones, A.K., Thany, S.H., 2018. Neonicotinoid insecticides mode of action on insect nicotinic acetylcholine receptors using binding studies. Pestic. Biochem. Physiol. 151, 59–66. Telo, G.M., Senseman, S.A., Marchesan, E., Camargo, E.R., Jones, T., McCauley, G., 2015. Residues of thiamethoxam and chlorantraniliprole in rice grain. J. Agric. Food Chem. 63, 2119–2126. Tosti, E., Boni, R., Cuomo, A., 2000. Ca(2+) current activity decreases during meiotic progression in bovine oocytes. Am J Physiol Cell Physiol 279, C1795–C1800. Vercesi, A.E., Kowaltowski, A.J., Grijalba, M.T., Meinicke, A.R., Castilho, R.F., 1997. The role of reactive oxygen species in mitochondrial permeability transition. Biosci. Rep. 17, 43–52. Veysset, P., Lherm, M., Bebin, D., Roulenc, M., 2014. Mixed crop-livestock farming systems: a sustainable way to produce beef? Commercial farms results, questions and perspectives. Animal 8, 1218–1228. Wang, X., Anadon, A., Wu, Q., Qiao, F., Ares, I., Martinez-Larranaga, M.R., Yuan, Z., Martinez, M.A., 2018. Mechanism of neonicotinoid toxicity: impact on oxidative stress and metabolism. Annu. Rev. Pharmacol. Toxicol. 58, 471–507. Wehrend, A., Meinecke, B., 2001. Kinetics of meiotic progression, M-phase promoting factor (MPF) and mitogen-activated protein kinase (MAP kinase) activities during in vitro maturation of porcine and bovine oocytes: species specific differences in the length of the meiotic stages. Anim. Reprod. Sci. 66, 175–184. Wu, B., Ignotz, G., Currie, W.B., Yang, X., 1997. Dynamics of maturation-promoting factor and its constituent proteins during in vitro maturation of bovine oocytes. Biol. Reprod. 56, 253–259. Wuest, M., Perreault, A., Richter, S., Knight, J.C., Wuest, F., 2019. Targeting phosphatidylserine for radionuclide-based molecular imaging of apoptosis. Apoptosis 24, 221–244. Yan, S.H., Wang, J.H., Zhu, L.S., Chen, A.M., Wang, J., 2016. Thiamethoxam induces oxidative stress and antioxidant response in zebrafish (Danio Rerio) livers. Environ. Toxicol. 31, 2006–2015. Yang, H., Xie, Y., Yang, D., Ren, D., 2017. Oxidative stress-induced apoptosis in granulosa cells involves JNK, p53 and Puma. Oncotarget 8, 25310–25322. Yi, M., Rosin, M.P., Anderson, C.K., 1990. Response of fibroblast cultures from ataxiatelangiectasia patients to oxidative stress. Cancer Lett. 54, 43–50. Yuh, H.S., Yu, D.H., Shin, M.J., Kim, H.J., Bae, K.B., Lee, D.S., Lee, H.C., Chang, W.K., Park, S.B., Lee, S.G., Park, H.D., Ha, J.H., Hyun, B.H., Ryoo, Z.Y., 2010. The effects of various antioxidants on the development of parthenogenetic porcine embryos. In Vitro Cellular and Developmental Biology: Animal 46, 148–154.
platelets via ROS mediated mitochondrial damage. Biochem. Biophys. Res. Commun. 438, 198–204. Green, T., Toghill, A., Lee, R., Waechter, F., Weber, E., Noakes, J., 2005. Thiamethoxam induced mouse liver tumors and their relevance to humans. Part 1: mode of action studies in the mouse. Toxicol. Sci. 86, 36–47. Jeschke, P., Nauen, R., 2008. Neonicotinoids-from zero to hero in insecticide chemistry. Pest Manag. Sci. 64, 1084–1098. Jo, Y.J., Jang, W.I., Namgoong, S., Kim, N.H., 2015. Actin-capping proteins play essential roles in the asymmetric division of maturing mouse oocytes. J. Cell Sci. 128, 160–170. Kanka, J., Kepkova, K., Nemcova, L., 2009. Gene expression during minor genome activation in preimplantation bovine development. Theriogenology 72, 572–583. Kanka, J., Nemcova, L., Toralova, T., Vodickova-Kepkova, K., Vodicka, P., Jeseta, M., Machatkova, M., 2012. Association of the transcription profile of bovine oocytes and embryos with developmental potential. Anim. Reprod. Sci. 134, 29–35. Kitajima, T.S., Ohsugi, M., Ellenberg, J., 2011. Complete kinetochore tracking reveals error-prone homologous chromosome biorientation in mammalian oocytes. Cell 146, 568–581. Ledan, E., Polanski, Z., Terret, M.E., Maro, B., 2001. Meiotic maturation of the mouse oocyte requires an equilibrium between cyclin B synthesis and degradation. Dev. Biol. 232, 400–413. Lee, W., Lee, D.G., 2018. Reactive oxygen species modulate itraconazole-induced apoptosis via mitochondrial disruption in Candida albicans. Free Radic. Res. 52, 39–50. Li, M., Li, S., Yuan, J., Wang, Z.B., Sun, S.C., Schatten, H., Sun, Q.Y., 2009. Bub3 is a spindle assembly checkpoint protein regulating chromosome segregation during mouse oocyte meiosis. PLoS One 4, e7701. Li, Y., Liu, X., Jiang, D., Lin, Y., Wang, Y., Li, Q., Liu, L., Jin, Y.H., 2016. Betulin induces reactive oxygen species-dependent apoptosis in human gastric cancer SGC7901 cells. Arch. Pharm. Res. 39, 1257–1265. Lincoln, A.J., Wickramasinghe, D., Stein, P., Schultz, R.M., Palko, M.E., De Miguel, M.P., Tessarollo, L., Donovan, P.J., 2002. Cdc25b phosphatase is required for resumption of meiosis during oocyte maturation. Nat. Genet. 30, 446–449. Mayer, A., Baran, V., Sakakibara, Y., Brzakova, A., Ferencova, I., Motlik, J., Kitajima, T.S., Schultz, R.M., Solc, P., 2016. DNA damage response during mouse oocyte maturation. Cell Cycle 15, 546–558. Memili, E., First, N.L., 2000. Zygotic and embryonic gene expression in cow: a review of timing and mechanisms of early gene expression as compared with other species. Zygote 8, 87–96. Morado, S.A., Cetica, P.D., Beconi, M.T., Dalvit, G.C., 2009. Reactive oxygen species in bovine oocyte maturation in vitro. Reprod. Fertil. Dev. 21, 608–614. Nurse, P., 1997. Checkpoint pathways come of age. Cell 91, 865–867. Omata, C., Kato, T.A., 2019. In situ DNA damaging foci analysis on metaphase chromosomes. Methods Mol. Biol. 1984, 87–93. Perez, G.I., Acton, B.M., Jurisicova, A., Perkins, G.A., White, A., Brown, J., Trbovich, A.M., Kim, M.R., Fissore, R., Xu, J., Ahmady, A., D'Estaing, S.G., Li, H., Kagawa, W., Kurumizaka, H., Yokoyama, S., Okada, H., Mak, T.W., Ellisman, M.H., Casper, R.F., Tilly, J.L., 2007. Genetic variance modifies apoptosis susceptibility in mature oocytes via alterations in DNA repair capacity and mitochondrial ultrastructure. Cell Death Differ. 14, 524–533. Senyildiz, M., Kilinc, A., Ozden, S., 2018. Investigation of the genotoxic and cytotoxic effects of widely used neonicotinoid insecticides in HepG2 and SH-SY5Y cells. Toxicol. Ind. Health 34, 375–383.
10