Carbofuran Alters Centrosome and Spindle ... - Oxford Journals

TOXICOLOGICAL SCIENCES, 144(2), 2015, 298–306 doi: 10.1093/toxsci/kfu317 Advance Access Publication Date: January 5, 2015

Carbofuran Alters Centrosome and Spindle Organization, and Delays Cell Division in Oocytes and Mitotic Cells Ozgur Cinar1, Olcay Semiz, and Alp Can Department of Histology-Embryology, Ankara University School of Medicine, Sihhiye, 06100 Ankara, Turkey 1

To whom correspondence should be addressed. Fax: þ90 312 310 6370. E-mail: [email protected].

ABSTRACT Although many countries banned of its usage, carbofuran (CF) is still one of the most commonly used carbamate derivative insecticides against insects and nematodes in agriculture and household, threatening the human and animal health by contaminating air, water, and food. Our goal was to evaluate the potential toxic effects of CF on mammalian oocytes besides mitotic cells. Caspase-dependent apoptotic pathway was assessed by immunofluorescence and western blot techniques. Alterations in the meiotic spindle formation after CF exposure throughout the in vitro maturation of mice oocyte-cumulus complexes (COCs) were analyzed by using a 3D confocal laser microscope. Maturation efficiency and kinetics were assessed by direct observation of the COCs. Results indicated that the number of TUNEL-positive cells increased in CF-exposed groups, particularly higher doses (>250 mM) in a dose-dependent fashion. The ratio of anticleaved caspase-3 labeled cells in those groups positively correlated with TUNEL-positivity. Western blot analysis confirmed a significant increase in active caspase-3 activity. CF caused a dose-dependent accumulation of oocytes at prometaphase-I (PM-I) of meiosis. Partial loss of spindle microtubules (MTs) was noted, which consequently gave rise to a diamond shape spindle. Aberrant pericentrin foci were noted particularly in PM-I and metaphase-I (M-I) stages. Conclusively, CF (1) induces programmed cell death in a dose-dependent manner, and (2) alters spindle morphology most likely through a mechanism that interacts with MT assembly and/or disorientation of pericentriolar proteins. Overall, data suggest that CF could give rise to aneuploidy or cell death in higher doses, therefore reduce fertilization and implantation rates. Key words: apoptosis; carbofuran; cell division; mitotic cells; oocyte

Carbofuran (CF) is a widely used potent methylcarbamate derivative insecticide (CAS No. 1563-66-2; 2,3-dihydro-2,2-dimethyl-7benzofuranol methyl carbamate) in agriculture and household to control the growth of the insects and nematodes. Although CF usage has recently been banned in many developed countries including the EU countries, the United States and Canada due to environmental toxicity, it has been still in use in many African countries (Jemutai-Kimosop et al., 2014; Otieno et al., 2010). CF has systemic anticholinergic actions in cortex and striatum regions of central nervous system by forming carbofuranacetylcholinesterase complex (Gupta, 1994). Besides, many systemic side effects were also reported such as sterility (Baligar and Kaliwal, 2002; Pant et al., 1995, 1997; Yousef et al., 1995), congenital abnormalities (Gupta, 1994; Maroni et al., 2000),

clastogenicity and hepatotoxicity, and it increases the risk of a series of dysfunctions on gastrointestinal (Ram and Singh, 1988; Rizos et al., 2004), neurological (Kamboj et al., 2006; Kim et al., 2004), and endocrine systems (Ram, 1988) in animals. In humans, retinal degeneration (Kamel et al., 2000) and DNA damage in spermatozoa (Meeker et al., 2004) were reported to occur by the contamination of air, water, and food (Chauhan et al., 2000; Kamel et al., 2000; Maroni et al., 2000). These disturbances are the result of its direct action or via its metabolites on the structural and metabolic changes on various cell types. Among those, apoptosis and cell cycle arrest (Soloneski et al., 2008; Yoon et al., 2001), and chromosome aberrations (Chauhan et al., 2000; Lin et al., 2007) were demonstrated to take place when eukaryotic cell lines or isolated mice were exposed to varying

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doses of CF or its most common metabolite, N-nitrosocarbofuran. Due to the antimitotic effects of CF, the potential chemotherapeutic roles have recently become a widely discussed topic (Amanullah and Hari, 2011). It was previously shown that CF (2 mg/kg of body weight) induces mitotic inhibition and chromosome aberrations in mouse bone marrow cells (Chauhan et al., 2000). Similarly, decreased sperm motility, reduced epididymal sperm count coupled with increased morphological abnormalities in head, neck, and tail of spermatozoa were observed in rats exposed to 0.2–0.8 mg/kg CF (Pant et al., 1995). There is a growing body of evidence implying that CF interferes with chromosome integrity (Meeker et al., 2004), which consequently perturbs cell division. However, no data have been published so far, which intended to search for a link between CF and cell cycle kinetics and mitotic/meiotic spindle. In eukaryotic cells, mitosis and meiosis are regulated with a group of complex proteins that ultimately coordinate the integrity of microtubule-oriented spindle, centrosomes, and interconnected proteins. The assembly and appropriate organization of the microtubule (MT) cytoskeleton is an integral phenomenon, which is related to the expression of cellular asymmetry. Particularly in oocytes, MT display a unique paradigm as forming an eccentric meiotic spindle in a huge MT-decorated oocyte cytoplasm that consequently gives rise to asymmetric cytokinesis to form the first and the second polar bodies (Can et al., 2003). The driving machinery and the organization of the MT cytoskeleton are mainly concerned to the centrosomes or microtubule organizing centres (MTOC), which serve as sites of microtubule nucleation (Dictenberg et al., 1998). Therefore, in the present study, we aimed to test the potential toxic effects of CF on the in vitro maturation kinetics and meiotic spindle structure regarding the cytoskeletal orientation and centrosomes of mammalian oocytes. Moreover, the cell growth kinetics in somatic cells using in vitro experimental models on Vero cell lines was evaluated.

MATERIALS AND METHODS Preparation of CF. CF (purity 98%, MW 221.25) was purchased from Aldrich Chemical Co (Aldrich Chemical Co, St Louis, MO) and dissolved in dimethyl sulfoxide (DMSO) (Sigma Chemical Co, St Louis, MO) to prepare 1 M stock solution. DMSO concentration of treated and control samples never exceeded 0.1% v/v in working solution and at a given concentration, no adverse effects of DMSO on oocyte maturation and on Vero cell growth were observed. Collection, in vitro maturation, and fixation of oocytes. The study was approved by the institutional Animal Ethics Committee. COCs (n ¼ 1017) were obtained from 19 to 21 day-old Balb/C mice injected 5 IU of pregnant mare serum gonadotropin (Sigma) 48 h prior to oocytes collection. Following the sacrification of animals by cervical dislocation, ovaries were obtained, follicles were punched with a needle, and COCs were obtained. COCs were transferred into a collection medium (Eagle’s MEM with Hank’s salts and HEPES buffer supplemented with 100 IU/ml penicillin, 100 mg/ml streptomycin and 0.3% bovine serum albumin, BSA) and then incubated in CF-free or CF-containing Eagle’s MEM supplemented with Earle’s salts, 2 mM glutamine, 0.23 mM pyruvate, 100 IU/ml penicillin, 100 mg/ml streptomycin and 0.3% BSA for in vitro maturation. After removal of cumulus cells by gentle pipetting, denuded oocytes were fixed and extracted for 20 min at 37 C in a microtubule-stabilizing buffer containing 2% formaldehyde, 0.5% Triton-X 100, 1 mM

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paclitaxel, 10 U/ml aprotinin and 50% deuterium oxide (Can et al., 2003). Samples were washed three times in a blocking solution of phosphate buffered saline (PBS) containing 2% BSA, 2% powdered milk, 2% normal goat serum, 0.1 M glycine and 0.01% Triton X-100 and stored at 4 C in blocking solution until processing. CF treatment of oocytes during in vitro maturation. A series of dose experiments were initially performed in order to determine the HTD, which is the highest concentration of CF that can be tolerated by oocytes at which only minimal morphological alterations were evident. Freshly prepared 10, 50, 100, 200 or 400 mM CF in oocyte maturation medium were used to treat COCs during different stages of meiotic division, and 50 mM CF was determined as HTD; therefore, subsequent tested dose (100 mM) was used for experiments to further observe the effects of CF on meiotic spindle and progression of meiosis. COCs were treated with CF during initial stages of in vitro maturation for 8 h between germinal vesicle (GV)-stage and metaphase-I (M-I) (n ¼ 241). To test the reversibility of the effects appeared during this stage; a group of COCs (n ¼ 132) were washed from the drug and cultured for an additional 10 h in the control medium. A different set of COCs was exposed to CF (n ¼ 262) during 8–18 h interval between M-I and M-II. A total 154 COCs were used as the control group cultured in control medium (see Fig. 1 for the experiment design). Fluorescent labeling of oocytes. Oocytes were incubated with primary and then secondary antibodies for 2–5 h/antibody at either 4 C or 37 C in a rotating shaker followed by 15-min washes in a PBS-blocking solution between incubation steps. Spindle MTs were visualized using a 1:1 mixture of a þ b tubulin mouse monoclonal antibodies (1:100 dilution) (Sigma) followed by a 1:100 dilution of an affinity-purified FITC goat antimouse IgG (Jackson ImmunoResearch Laboratories, PA). Centrosomes were labeled with a rabbit polyclonal antibody directed against pericentrin protein (Abcam, MA) at a final dilution of 1:100, followed by a 1:100 dilution of an affinity-purified Cy5 conjugated donkey antirabbit IgG (Jackson ImmunoResearch Laboratories). For the evaluation of chromatin and chromosomes at specific stages in meiotic progression, oocytes were stained by 10 mM 7-aminoactinomycin-D (7-AAD) (Sigma) (Can et al., 2003). Finally, oocytes were mounted between glass coverslips and slides using spacers allowing a 170 mm space in between which was filled with a 1:1 glycerol/PBS medium containing a DNA dye Hoechst 33258 (1 mg/ml) and 25 mg/ml sodium azide as antifading reagent (Can and Albertini, 1997). 3D image reconstruction by confocal scanning microscopy and analysis of MTS loss. Labeled oocytes were initially examined by Zeiss Axiovert 100M inverted microscope (Jena, Germany) using conventional fluorescence UV filter set and mercury-arc lamp. Hoechst staining of nuclear material was used to double check for the 7-AAD staining (red emission). Then, a Zeiss LSM-510 Meta confocal laser scanning microscope (Jena) equipped with 63x plan-apo objective was used for further detection of centrosomes, MTs, and chromosomes. 3D images were constructed from consecutive optical sections using LSM software ver. 3.2. The quantitation of positive signals from tubulin staining were analyzed using ImageJ v.1.47 software by a previously described method (Burgess et al., 2010). Briefly, the formula used to calculate the corrected total cell fluorescence (CTCF) was CTCF ¼ Integrated density – (area of selected signal mean fluorescence of background readings). The CTCF results of control

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FIG. 1. Experiment design used to detect the cell cycle effects of CF on mouse oocytes. Dotted lines indicate CF-free medium, solid lines indicate CF treatments.

groups were compared with CF-treated groups at GV, M-I, or M-II stages, respectively. Cell synchronization, CF exposure, and mitotic index. Vero cells (ATCC, VA) were cultured in Dulbecco’s Minimum Essential Medium-Ham’s F-12 cell culture medium (Sigma) supplemented with 10% fetal calf serum (Gibco BRL, Rockville, MD). Cells were allowed to expand in T75 polystyrene culture flasks until 100% confluency at 37 C in a 5% CO2-95% air humidified incubator. Expanded cells were subcultured onto 12 mm diameter poly L-lysine coated glass coverslips placed in 24-well polystyrene flasks. Cells were allowed to grow until 70% confluency. To examine the effects of CF, cell cycles of Vero cells were initially synchronized by 0.3 mM nocodazole (a benzimidazole-derivative microtubule depolymerizing agent) (Sigma) for 12 h followed by triple wash by Dulbecco’s PBS, and then transferred in a nocodazole-free medium for 12 h (Cinar et al., 2008). Freshly prepared CF stock solution was diluted in the culture media at 100, 250, 500, 1000, or 2000 mM concentrations primarily based on the previous experiments by Stehrer-Schmid and Wolf (1995). Subsequently, concentrations and time of treatments would allow determining the “inhibition concentration 50 (IC50),” the concentration of a drug where the 50% of cells treated retained alive. Vero cells were incubated following the lag phase (48 h after plating) in CF-containing culture media for 12 or 24 h. To test whether CF effects are recoverable or not, CF-treated groups were incubated in CF-free culture media after cultured for 12 or 24 h in varying CF doses. Cells on coverslips were fixed and extracted for 15 min at 37 C in a microtubule-stabilizing buffer (see above) (Can et al., 2003). The samples were washed twice and stored in PBS (Sigma) containing 0.02% sodium azide until processing. Coverslips were incubated with a mouse monoclonal anti a þ b tubulin (1:1) antibody (Sigma) for 90 min at 37 C and then incubated with an affinity purified Cy3-conjugated goat antimouse IgG (Jackson ImmunoResearch Laboratories). Cells were washed with PBS after each step. Chromosomes were marked with 1.5 mM Sytox green (Molecular Probes, Eugene, OR) for 3 min. Microscopic observations were performed using Zeiss LSM 510 confocal microscope (Germany). The ratio of the total number of cells in mitosis (through prophase to telophase) to the total number of counted cells was defined as mitotic index. Cell growth assay. Cytotoxicity was tested by the MTT (Sigma) assay that determines viable cell numbers and was based on the mitochondrial conversion of the tetrazolium salt, 3-[4,5dimethylthiazol-2-yl]-2,5-diphenyltetra-zolium bromide. For this purpose, cells were initially transferred to 96-well plate. Following the adhesion and reaching the 70% confluency, different concentrations of CF (see below) in 150 ml culture medium were added and incubated for 6, 12, 24, 48, 52, 60, and 72 h. A total of 15 ml MTT from 5 mg/ml stock in PBS was added into each well, and then plates were kept in the CO2 incubator for

3 h. The converted dye was solubilized with 150 ml acidic isopropanol (0.04 M HCl in absolute isopropanol) and absorbance of the converted dye was measured at a wavelength of 570 nm with background subtraction at 650 nm in a spectrophotometer (Thermo Scientific, Wilmington, DE). Two replicates were read for each sample, and values were normalized as 0.0 (blank) to 1.0 (maximum value). Apoptosis and apoptotic index. Terminal uridine deoxynucleotidyl transferase dUTP nick end labeling (TUNEL) was used to detect apoptotic cells. A 50 ml mixture (9:1) of labeling and enzyme solutions of a TUNEL kit (Roche, Germany) was applied to 4% of paraformaldehyde-fixed cells on coverslips, followed by incubation for 60 min at 37 C in a dark humidified chamber. As a negative control, the enzyme solution (terminal deoxynucleotidyl transferase) was omitted from the reaction mixture. The ratio of the total number of TUNEL-positive cells to total number of counted cells was defined as apoptotic index. Alterations of active caspase-3, a member of apoptosis execution enzymes, were visualized by a polyclonal rabbit anticleaved caspase-3 (Asp 175) antibody (Cell Signaling, Beverly, MA) diluted in Tris-buffered saline (TBS; 100 mM Tris, 0.9% NaCl, pH: 7.5) for 2 h at 37 C in a humidified chamber. Following triple wash in TBS, slides were incubated with an affinity purified Cy3 conjugated goat antirabbit IgG (1:100 in PBS; Jackson ImmunoResearch) for 90 min at 37 C in a dark humidified chamber and finally mounted in PBS/glycerol medium containing 1 mg/ml of Hoechst 33258 (Polyscience Inc, PA) for nuclear staining and 25 mg/ml sodium azide (Sigma) as an antifading reagent (Can and Albertini, 1997). Quantification of active caspase-3. Total protein extracts were removed from cells using a mixture of cell extraction buffer (Biosource International Inc., Camarillo, CA) with a proteinase inhibitor cocktail (Sigma) and 1 mM phenyl-methyl-sulfonyl fluoride (Sigma). The protein concentration was determined by a detergent-compatible protein assay kit (BioRad Laboratories, Hercules, CA). A total of 30 mg protein was loaded into each lane of a 15% SDS-PAGE gel. The proteins were electroblotted onto nitrocellulose membrane (BioRad), washed with distilled water and then incubated in 5% (w/v) nonfat dry milk in TBS containing 0.1% Tween 20 (TBS-T) for 1 h to reduce nonspecific binding. Membranes were then incubated with a polyclonal rabbit anticleaved caspase-3 (Asp 175) antibody for 2 h at room temperature and washed 3 times with TBS-T over a total period of 20 min. Blots were then incubated in an HRP-labeled goat antirabbit IgG (Sigma) in TBS-T for 1 h at room temperature followed by 10 washes with TBS-T over a total of 1 h. A chemiluminescence-detecting reagent (Amersham-Biosciences, NJ) was used to illuminate the blots with and 25 25 cm films (Kodak, Rochester, NY) were used to show antibody-bound bands. Immunoblot analyses were quantified using a laser densitometer (Kodak). Membranes were stripped with stripping solution


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(Pierce, Rockford, IL) and remarked with a mouse monoclonal anti-b-actin (Sigma) antibody for normalization of caspase-3 bands. Statistical analysis. Statistical analyses were performed using the computer-based software package (version 15.0; SPSS Inc, Chicago, Illinois). Apoptotic index and mitotic indices were detected by counting the cells from five various microscopic area of each slide at 20 magnification. Differences in apoptotic index or mitotic indices between groups were evaluated by chisquare test. The statistical evaluation of MTT outcomes were analysed using repeated measures ANOVA with a GreenhouseGeisser correction for sphericity and post hoc Tukey-HSD test. Oocyte progression study or CTCF outcomes were performed by chi-square or student’s t test, respectively. The differences among signal intensities from active caspase-3 or b-actin bands were individually tested utilizing the one-way ANOVA and post hoc LSD tests. Level of significance was selected as p < .05.

RESULTS CF delays oocyte meiotic maturation. COCs were exposed to 100 mM of CF for either 0–8 or 8–18 h of culture, during which in vitro oocyte maturation in mice is normally completed. The first half of meiotic division (meiosis-I), 26% of cells remained in PM-I, whereas the rest reached M-I (73%) or even anaphase-I (A-I) (1%) in CF group (Fig. 2A). Compared with the controls, this group showed a significant delay in progression, and was blocked mostly around PM-I (p ¼ 0.026). When cells were exposed to CF during the transition from M-I to M-II, which normally lasts approximately 8–10 h in mice, half of them were found at M-I (52%), a small number of oocytes were found at PM-I stage (6%), and only 42% of oocytes succeeded to reach M-II stage (Fig. 2B), whereas a significant number of oocytes (92%) in control group reached M-II stage (p < .001). To determine whether the adverse effects of CF on cell cycle progression and the organization of centrosomes and spindles are reversible, oocytes were treated with CF for 8–10 h, then washed and transferred into control maturation medium for an additional 10 h (Fig. 1 and Fig. 2C). During 18 h of total incubation, 92% of control cells progressed to M-II, whereas smaller ratio of cells (8%) accumulated in PM-I to T-I. On the other hand, none of the CF-treated oocytes managed to reach M-II stage, and were found stuck at the different stages of first meiotic division (Fig. 2C). CF disrupts meiotic spindle and causes loss of MTs. Control PM-I oocytes were characterized by de novo synthesis of the first meiotic spindle in a nonpolarized form in the centre of the oocyte (Fig. 3A). Pericentrin protein was restricted to multiple, centrally located spots where microtubules were emanating. M-I spindles displayed the typical barrel shape slightly translocated to the periphery of the cell (Fig. 3B). Chromosomes were aligned around mid-plane where they were tightly bound to kinetochore microtubules. The long axis of the spindle, which transverses the spindle poles, was aligned perpendicular to the adjacent plasma membrane. Pericentrin foci were oriented at spindle poles forming two symmetrical crescents (Fig. 3B). Regarding the microtubule mass and interpolar distance, M-II spindles were found slightly smaller than M-I spindles (Fig. 3C), yet they maintained their barrel shape. They were

FIG. 2. Kinetics of oocyte maturation in control and 100 mM CF treatment groups during 0–8 h (A), 8–18 h (B), and recovery period of cell after first 8 h of CF treatment (C). A total of 26% of CF-treated oocytes remain in PM-I after 8 h CF exposure (A). More than 50% of oocytes do not reach to metaphase II (B). Oocytes could not manage to resume meiosis after CF removal from the medium and stuck at various stages of meiosis-I (C).

often found rotated compared with M-I spindles, so that the long axis became nearly parallel to the adjacent plasma membrane. Compared with M-I centrosomes, pericentrin was found similar but in smaller amounts. The spindle dynamics

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FIG. 3. Three-dimensional confocal microscopic images of spindles in control (A–C) and CF-treated (D–F) oocytes. Triple labeled meiotic spindles stained with antibodies against a þ b tubulin (1:1) (green); pericentrin (blue) and chromosomes (red). Microtubules originated from pericentrin foci are found loosely related to chromosomes at PM-I spindles (A). The spindle at M-I displays a typical barrel shape (B). Pericentrin foci are symmetrically placed at the spindle poles as two crescents. Centrally located chromosomes, symmetrical pericentrin foci at each poles, and MTs emanating from centrosomes are consistently observed at M-II spindles (C). Although CFtreated PM-I oocytes showed a slight loss of tubulin mass (D), pericentrin and chromosomes are found to be similar to that of controls. In M-I oocytes MTs lose their mass, and tend to radiate peripherally (E). Condensed pericentrin foci are displaced from both poles and scattered along the spindle microtubules. No significant displacement of chromosomes from mid-plane is detected. CF-treated M-II oocytes are associated with a few condensed spots of pericentrin located at each pole and few foci along with the spindle (F) Scale bar: 10 mM. Bar graphs represent the signal intensities obtained from tubulin fluorescence following the CTCF calculation (G). *p < .05 Control versus CF-treated group in GV or M-II stage oocytes.

summarized above are considered as normal events that are required for proper meiotic progression (Carabatsos et al., 2000). In the CF-treated group, a newly forming spindle was typically found similar to that of in a normal PM-I cell. Pericentrin, where MTs are emanating, was found to be associated with the spindle (Fig. 3D). No significant difference was observed between the control and the CF group apart from a lesser amount of MTs in CF-treated oocytes. Regarding M-I oocytes, typical barrel-shaped spindle was not seen. A decreased amount of MTs and scattered pericentrin foci between the spindle poles were consistently observed (Fig. 3E). Spindle MTs tend to displace particularly from the lateral sides of the spindle. In contrast of these alterations regarding microtubule orientation of spindle and pericentrin pattern, chromosomes were usually found to locate around the mid-plane (Fig. 3E). In M-II oocytes (Fig. 3F), partial loss of spindle MTs was frequently noted resulting in a cylindrical spindle. No misalignment of chromosomes

was noted around the spindle. Several pericentrin foci besides in polar bodies were also noted. MT integrity in GV, M-I and M-II stage oocytes was compared after calculating the CTCF values (Fig. 3G). We found a significant difference between control and CF-treated oocytes in GV (p ¼ .01) or M-II (p < .001) stages. However, the difference was not significant in M-I stage oocytes (p ¼ 0.12). Granulosa cells were not affected in 100 mM CF group (data not shown). Cytotoxic effects of CF on mitosis and cell growth. Population doubling time (PDT) of Vero cells was calculated directly from the growth curve during log phase and was found 22 6 4 h. After cells were synchronized by nocodazole for 12 h followed by a 12h wash (Cinar et al., 2008), cells were exposed to 100–2000 mM CF for 12 or 24 h. Although mitotic indices increased approximately 2.2–3.5-fold and 1.7-3.2-fold during the first 12 and 24 h-treated


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Mitotic spindle was not affected in CF; neither microtubule reorganization nor chromosome alignment was similar to that of the control group. Toxic effects of CF on Vero cell viability as a function of mitochondrial activity were measured by MTT assays at 6, 24, 48, 60, and 72 h after CF treatment. All tested doses of CF significantly inhibited the cell viability in a dose-dependent fashion compared with the control (Fig. 4B). Cell growth in 250, 500, 1000, and 2000 mM CF groups were significantly inhibited particularly after 24 h of continuous CF exposure (p < .05). The duration of IC50 for 2000, 1000, and 250 mM CF was calculated as 6, 24, and 48 h incubation periods, respectively. TUNEL and apoptotic index Apoptosis was detected by the TUNEL assay. An increase in the number of apoptotic cells was observed after CF treatments, in a dose-dependent fashion. In the control group, 0.5% of cells were apoptotic but this ratio significantly increased up to 1.5% at 100 mM and reached around 7% at 2000 mM (Fig. 4C). No significant difference was found between the control and 100 mM or 250 mM doses. On the other hand, apoptotic indices in 500, 1000, and 2000 mM doses were significantly higher than that of the control (p < .05). Moreover, apoptotic indices of 1000 and 2000 mM groups were significantly higher than the other CFtreated groups (p < .001). Taken together, mitotic indices, MTT and TUNEL results indicate that CF blocks the cell cycle at the prophase/metaphase stages of mitosis and inhibits cell growth, therefore considerably lowers the cell viability.

FIG. 4. Dose- and time-dependent analysis of apoptosis/proliferation balance in mitotic cells after CF exposure. Mitotic indices increase in a dose-dependent manner after CF exposure. *p < .05 Control group versus others after 12 h treatment. **p < .05 Control group versus others after 24 h treatment (A). MTT analyses show that the total cell number of mitotic cells gradually decreased in a dose-dependent manner during 72 h CF exposure (B). The number of apoptotic cells increase after CF treatment (C). *p < .05 Control group versus others after 12 h treatment. **p < .05 Control group versus others after 24 h treatment.

groups, respectively (Fig. 4A), there was no significant difference between 12 and 24 h groups. Mitotic index in control groups was found significantly lower than that of all the CF-treated groups (p < .05). Mitotic index in 2000 mM CF treatment during 24 h was found significantly higher than the other 24 h groups (p < .05). Considering the accumulated stage of mitotic cells after CF treatment, they were found either at prophase or metaphase. Almost no cell managed to reach further stages of mitosis.

Active caspase-3 staining and Western blot analysis of active caspase-3 Immunofluorescence analysis showed that active caspase-3 enzyme was located in the entire cytoplasm excluding the nucleus in both the control and the CF-treated cells. The number of the positive cells, and the strength of the intensity increased in a dose-dependent manner (Fig. 5A). Western blot analyses showed a significant increase in active caspase-3 protein, which were strongly consistent with the results of fluorescent signals (Fig. 5B). The amount of active caspase-3 increased after the CF incubation in a dose-dependent manner, particularly in 1000 and 2000 mM CF groups, whereas in the untreated cells it was expressed significantly lower (p < .05) (Fig. 5C). Taken together, in all mitotic cell experiments, the progression of mitosis seemed to be affected by 100 mM CF exposure; 500 mM CF significantly induced the apoptotic cell death. Thus, 250 mM of CF was considered as the critical dose for cell proliferation and apoptosis balance in mitotic cells.

DISCUSSION Carbamates, especially CF (marketed under the trade names Furadan and Curater), have been one of the most commonly used pesticides in agriculture and forestry due to their short-lasting residue persistence (Gupta, 1994). The effects of CF on organ systems in vivo and on cellular structure in vitro have been studied over the last 20 years (Baligar and Kaliwal, 2002; Chatterjee et al., 1997; Gupta et al., 1991; Kamboj et al., 2006; Kim et al., 2004). However, for the first time in the literature, here we concurrently demonstrate the toxic effects of CF on meiotic and mitotic cells by testing cell growth, cell cycle and apoptosis, oocyte maturation and meiotic spindle formation.

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FIG. 5. Alterations in the active caspase-3 enzyme after CF exposure. Immunofluorescent staining of the active caspase-3 enzyme in Vero cells (red) (A). Green signals: cell nuclei, scale bar: 40 mm. Western blot analyses of total active caspase-3 levels (around 17 kD) increase in a dose-dependent manner after CF exposure (B). Bar graphs demonstrate the densitometric illustration of blots in B. *p < .05 control versus other groups (C).

Lag phase of cell proliferation in Vero cells terminates around 48 h following seeding where log phase begins. MTT analysis showed that high doses of CF, 1000 and 2000 mM decrease cell number during lag phase indicating an increased cell death. In a study by Yoon et al, it was noted that the concentrations between 1 mM and 50 mM CF did not influence the growth rate of CHL (Chinese hamster lung fibroblast) cells, but one of its metabolite N-Nitrosocarbofuran (NOCF) significantly inhibited the cell growth (Yoon et al., 2001), the IC50 at 48 h of incubation was estimated 2.8 mM. In the current study, IC50 of 48th h was found 250 mM for CF. Naravaneni and Jamil (2005) showed that CF caused a significant increase in the frequency of chromosomal aberrations after 24 h CF exposure on shortterm lymphocyte cultures, and LD50 of CF was estimated 18 mM. Despite the fact that 100 mM CF has minimal effects on Vero cells; oocytes were substantially affected in this concentration. Although significant cell death was noticed after the CF exposure during lag phase, the cell number continued to decrease during log phase at the dose of 500, 1000, and 2000 mM. These findings imply that either cell death rate is higher than the duplication rate of cells or mitotic process is somehow blocked, and cells die subsequently. When mitotic stages of cells were

analysed, many of them were found at prophase and metaphase stages. Stehrer-Schmid and Wolf (1995) concluded that CF and three other carbamate derivatives, including benzofuran, carbosulfan, and furathiocarb, lead to a dose-dependent reduction of the polymerization degree of porcine brain tubulin. They noted that 500 mM CF results in a minor increase of the polymerization rate, where maximum reduction in polymerization is seen in a concentration of 5000 mM. Thus, mitotic arrest at prometaphase stage observed in this study may be due to the spindle tubulin disruption, and 250 mM can be considered a critical dose which gives rise to the damage of MTs in Vero cells. On the other hand, 100 mM CF significantly delays the in vitro meiotic maturation of oocytes possibly by disrupting meiotic spindle. High magnification 3D confocal analysis of the oocytes demonstrated the reduction of the spindle volume, which reveals lower signal intensity indicating the reduced MT polymerization at GV or M-II stage oocytes. On the other hand, insignificant difference between control and CF groups at M-I stages implied that the arrest in M-I stage might be owing to the disruption of spindle integrity rather than the MT loss. Dislocation and distribution of centrosomes, as the main MT organizing centers, could be the main reason of spindle disruption and the arrest of the meiotic resumption. In the present study, Vero cell apoptosis increased especially by 500 mM and above doses of CF. Yoon et al. examined the apoptotic effects of CF and NOCF in CHL cell cultures at low doses (30 mM) (Yoon et al., 2001). They noted nuclear condensation, cell shrinkage, and a loss of internal nuclear structure as well as TUNEL-positivity after NOCF exposure but no morphological alteration and TUNEL-positive staining were observed after CF treatment (Yoon et al., 2001). Lee et al. (2004) demonstrated that NOCF induces apoptosis via cytochrome-c-mediated caspase activation. Similarly, Jung et al. (2003) showed that 10–20 mM NOCF inhibits cell viability, and induces apoptosis suggesting the involvement of the ERK pathway. They also reported that CF does not affect the cell viability at the concentrations of 3–30 mM. In contrast, Kim et al. (2004) observed that CF induced DNA fragmentation, shrinkage due to protein cross-linking, membrane alterations, translocation of phosphatidyl serine from the inner leaflet of the cell membrane to the outer leaflet as characteristics for apoptosis. They also noted that CF at slight (100 mM) and severe doses (300 mM) exhibits a series of cytotoxic effects. A total of 100 mM CF affects the cells 3 days after treatment, whereas 300 mM and higher doses significantly reduced the cell viability 1–3 days after treatment. Our results were similar to that of Kim’s study, and here, we concluded that mitotic cells were slightly influenced at 100 mM and moderately affected at 250 mM during the second day of the treatment. Higher doses of CF (above 500 mM) drastically decreased the cell viability and induced apoptosis. It was previously demonstrated that CF inhibited the fertilization capacity of mature oocytes (Chatterjee and Ghosh, 1995) and significantly decreased ovarian weight (Chatterjee et al., 1997) compared with the controls in catfish. In two independent studies (Baligar and Kaliwal, 2003, 2004), CF caused a considerable reduction in the number of estrous cycle and the duration of proestrus, significantly decreased the number of healthy follicles, and concomitantly increased the atretic follicles. Histologic observation of the mouse ovaries treated with CF showed the presence of few developing follicles, few small corpora lutea, and many atretic follicles. Although meiotic aneuploidy was reported relatively low in mice compared with humans, and the rates of hyperploidy are approximately 0.5%–1% (Bond and Chandley, 1983), an increasing amount of


data emerge from experiments in mice suggest that certain environmental agents possess potent meiosis-disrupting capability that directly interferes with the cell division machinery both in meiosis-I and early meiosis-II (Can and Albertini, 1997; Can et al., 2005). Chauhan et al. (2000) reported that CF caused an increase not only in the chromosome aberrations and micronucleus formation in mouse bone marrow cells but also induced the sperm shape abnormalities, such as amorphous, balloon or microcephaly in head region and bent tail. Similarly, CF-treated rabbits showed a decline in body weight, libido, ejaculate volume, sperm concentration, semen initial fructose, and semen osmolality (Yousef et al., 1995). Sperm abnormalities and the number of dead sperm increased after CF exposure as well as during the recovery period in a dose-dependent fashion. Abnormal sperm shapes due to the anomalies in head, neck, and/or tail region decreased sperm motility, reduced epididymal weight, damaged Sertoli cells and germ cells were demonstrated after CF treatment in rats (Chauhan et al., 2000; Pant et al., 1995, 1997). Mouse oocytes, COCs and meiotic spindles in particular, increasingly gain importance serving as sensitive cellular assay systems for the study of reproductive toxins. We now can carefully extend our results to humans. Nonetheless, meiosis is a highly conserved process among mammalian organisms. The other part of the present study was designed to evaluate the effects of CF on the process of meiotic cell division machinery in cultured mouse oocytes. Given the multistep nature of meiosis and the stage-specific involvement of the cytoskeleton in general, and the centrosomal proteins and microtubules in particular (Carabatsos et al., 2000), the present study was undertaken, for the first time in the literature, to evaluate the actions of CF during oocyte maturation. On the other hand, further studies are still required to address the effects of CF on intracellular microtubule dynamics and apoptotic pathway.

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