Styrene-7,8-oxide activates a complex apoptotic response in neuronal PC12 cell line

Mariarosaria Boccellino1, Franca Cuccovillo2, Maria Napolitano3, Nicola Sannolo4, Ciro Balestrieri1, Antonio Acampora2, Alfonso Giovane1 and Lucio Quagliuolo1

1 Dipartimento di Biochimica e Biofisica, Seconda Università degli Studi di Napoli, Via Costantinopoli 16, I-80138 Napoli, Italy
2 Dipartimento di Medicina Pubblica e Sicurezza Sociale, Università degli Studi di Napoli ‘Federico II’, Via S. Pansini 5, I-80131 Napoli, Italy
3 Istituto Nazionale per la Cura e lo Studio dei Tumori, Fondazione ‘Sen. G. Pascale’, Via M. Semmola, I-80131 Napoli, Italy
4 Dipartimento di Medicina Sperimentale, Sezione di Medicina del Lavoro, Igiene e Tossicologia Professionale, Seconda Università degli Studi di Napoli, Piazza Miraglia 2, I-80134 Napoli, Italy


    Abstract
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Styrene-7,8-oxide (SO), the major in vivo metabolite of styrene, one of the most important plastic monomers worldwide, is classified as carcinogenic in humans and animals. Although the toxic effects of SO have been extensively documented in human lymphocytes, the molecular mechanisms responsible for SO-induced cell damage are still unknown. In the present study, we evaluated the effect of SO on growth and apoptosis, assessed by FACS and gel ladder analysis, in neuronal PC12 cell line. Our results demonstrate that SO triggered PC12 cell apoptosis in a dose- and time-dependent manner. PC12 apoptosis was associated with caspase-3 activation and modulation of the Bcl-2 family proteins. In addition, examination of the cytoskeleton showed that SO induced F-actin depolymerization and a rapid cell rounding before caspase-3 activation, suggesting that the changes in cell shape involving cytoskeletal structure are an early step in the apoptotic pathway. Therefore, SO triggers a complex apoptotic response consisting of a loss of cytoskeletal organization that precedes caspase-3 activation. These mechanisms may represent the molecular basis of the different SO sensitivity to tumor promotion among species and organs.

Abbreviations: AFC, 7-amino-4-trifluoromethyl-coumarin; FITC, fluorescein isothiocyanate; PI, propidium iodide; SO, styrene-7,8-oxide.


    Introduction
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Styrene (vinylbenzene) used as an organic solvent, is a synthetic chemical widely used in the production of plastics and resins such as polystyrene resins, styrene-butadiene rubber and unsaturated polyester resins. Therefore, it represents a potential hazard for humans, particularly in the work place. However, apart from occupational setting, the exposure to very low levels of styrene can occur in ambient air (mainly via automobile exhausts), food (natural levels or through packaging made of styrene polymers or co-polymers) and cigarette smoke (1). Exposure occurs mainly via inhalation of styrene vapour; following inhalation, styrene is absorbed into the blood and metabolized in liver and lung to styrene-7,8-oxide (SO) by cytochrome P450-dependent monoxygenases (2,3). Further in vivo metabolism of SO involves two major pathways: hydrolysis catalyzed by epoxide hydrolase forming styrene glycol, a non-genotoxic metabolite, and conjugation of SO with glutathione by the glutathione S-transferase (4). In addition, SO is also produced commercially and used as a chemical intermediate, e.g. the preparation of fragrances and as a reactive diluent in manufacturing of epoxy resins (5).

Styrene and SO have been shown to be genotoxic in in vitro tests, with SO being more toxic than styrene (6). Moreover, styrene by inhalation had been shown to increase the lung tumor incidence in mice at 20 p.p.m. and higher, but was not carcinogenic in rats up to 1000 p.p.m. (7). SO binds covalently to peptides (8), human plasma proteins and haemoglobin; in vitro incubation of SO with blood from humans, mice and rats, demonstrated that SO reacts with a variety of nucleophilic sites in hemoglobin to form SO–Hb adducts (9–11). Furthermore, it has been demonstrated that SO induces DNA strand breaks (12,13). The genotoxic effects of SO are also documented in humans; in lymphocytes of SO-exposed workers there are increased frequencies of chromosome aberration, micronuclei and sister-chromatid exchanges (14). Lastly, there is substantial evidence that exposure to SO in humans brings about several changes in the central and the peripheral nervous systems, including depressive effects, neuroendocrine disturbances, disruption of dopaminergic neurones (15); and, in laboratory animals, tremor, convulsions and loss of equilibrium (16).

Although cytotoxic and genotoxic effects of SO are documented in peripheral blood lymphocytes (6,9,10,14), the mechanisms for producing these effects are still unknown. As DNA damage appears as an early event following SO exposure, it is expected that it underlies the cellular changes associated with SO.

Apoptosis normally eliminates cells with damaged DNA or an aberrant cell cycle, that is, those most likely to engender a neoplastic clone. Apoptosis is a genetically regulated cell death process characterized by a unique pattern of morphological changes in both the nucleus (chromatin condensation, internucleosomal DNA cleavage) and the cytoplasm (cytoskeletal disruption, cell shrinkage and membrane blebbing). The whole process leads to fragmentation of dying cells into apoptotic bodies that are phagocytosed by neighboring cells or macrophages (17–19). Biochemical changes such as the activation of members of the interleukine-1ß converting enzyme protease family, recently named ‘caspases’, also characterized apoptosis (20,21). Caspases are an evolutionarily conserved family of aspartic acid-specific cysteine proteases (22,23) that are synthesized as inactive precursor molecules and are converted to active heterodimers by proteolytic cleavage (24). The active form is composed of two subunits, large and small, both of which are derived from the precursor by cleavage at the C-terminal side of specific Asp residues. There is evidence that the caspases constitute a protease cascade that plays a prominent role in the apoptotic process (25).

Apoptosis is a complex phenomenon regulated by a finely tuned balance of inducer and repressor factors, e.g. the Bcl-2 gene family, heavily involved in the regulation of apoptosis, consists of both inducers and repressor of apoptosis, and their ratio determines, in part, the response to a death signal (26). In fact, the expression of anti-apoptotic factors, such as Bcl-2 and Bcl-xL, suppresses whilst the activation of pro-apoptotic factors, such as Bax and Bak, promotes apoptosis by countering Bcl-2 activity and thus accelerating apoptotic cell death. This highly regulated pathway plays a central role in the maintenance of cellular machinery as under physiological conditions, the Bcl-2 excess in the cell protects it from death. Alternatively, excess Bax leads to cell death and susceptibility to apoptosis (27).

The aim of the present study was to evaluate the effect of SO on growth and apoptosis in neuronal PC12 cells. In particular, we have investigated the apoptotic pathways in PC12 cells following treatment with SO. Our results demonstrate that SO induces typical apoptosis through activation of caspase-3 and modulation of the Bcl-2 family proteins in PC12 cells. Furthermore, we found that SO induced F-actin depolymerization and a rapid cell rounding before caspase-3 activation, suggesting that the changes in cell shape involving cytoskeletal structure are an early step in the apoptotic pathway.


    Materials and methods
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Reagents
Culture media and supplements were purchased from Life Technologies (Paisley, UK), and fetal calf serum was from Hyclone Lab (Logan, UT). SO, 98% (v/v), was obtained from Sigma-Aldrich Chemie GmbH, Germany. Rabbit polyclonal anti-procaspase-3, rabbit polyclonal anti-Bcl-2, rabbit polyclonal anti-Bax, and caspase-3 apoptosis detection kit were purchased from Santa Cruz Biotechnology (Santa Cruz, 2160 Delaware Avenue, CA). Etoposide and fluorescein isothiocyanate (FITC)-conjugated phalloidin (F-PHD) were obtained from Sigma Chemical Company (St Louis, MO). Annexin V-FITC was purchased from ImmunoKontact (AMS Biotechnology Europe, UK).

Cell culture and treatments
PC12 cells were grown at 37°C (5% CO2 atmosphere) in RPMI 1640 medium supplemented with 10% heat-inactivated horse serum, 5% heat-inactivated fetal bovine serum, 2 mM L-glutamine, 1 mM sodium pyruvate, 100 U/ml of penicillin and 100 mg/ml of streptomycin. Culture medium was changed three times per week and, when confluent, cells were split 1:6. For apoptosis evaluation subconfluent cells were treated with various concentrations of SO (0.1–1 mM), or 50 µM etoposide (VP-16) used as a positive control. SO stocks were prepared by dilution with an ethanol/water (60:40) mixture. Control cells were treated with 0.05% ethanol, which was the maximal final concentration of the vehicle in SO treated cultures. Apoptotic cells, which appeared as ‘floaters’, were harvested by centrifugation of culture medium at 1000 g for 5 min, whereas adherent cells remaining on culture dish and control cells were trypsinized, and subsequently collected by centrifugation.

Evaluation of apoptosis by flow cytometry
Apoptosis was evaluated by propidium iodide (PI) staining of cells followed by flow cytometry analysis, as described (28). Briefly, PC12 cells (106) were incubated for 4 h at 4°C in 2 ml hypotonic solution containing 50 µg/ml PI, 0.1% sodium citrate, 0.1% Triton X-100 and 20 µg/ml DNase-free RNase A. The stained cells were analyzed, for relative DNA content, on a FACScan flow cytometer (Becton Dickinson, San Jose, CA) with a 15 mV air-cooled argon laser tuned to single-line emission at 488 nm. Histograms of cell number versus logarithm integrated FL3 fluorescence were recorded for 20 000 nuclei at flow rates no greater than 50–100 events/s. Cells with subdiploid DNA content (subG0/G1 peak) were considered apoptotic cells. All cultures were done in triplicate.

In order to discriminate between apoptotic and necrotic cells we used the annexin V-FITC/PI staining assay (29). Cells were washed twice with annexin V-binding buffer (140 mM NaCl, 10 mM HEPES, 2.5 mM CaCI2, pH 7.4), resuspended in 1 ml of the same buffer and incubated in ice for 30 min with 2 µl of 140 nM annexin V-FITC. Five minutes before flow cytometry analysis, 5 µl of PI (50 µg/ml H2O stock solution) were added to each sample and then analyzed by flow cytometry (Becton Dickinson, San Jose, CA).

Internucleosomal DNA fragmentation (Ladder)
DNA fragmentation was measured after extraction of low molecular weight DNA. Briefly, 5x106 cells were resuspended in 500 µl Tris–EDTA buffer and lysed with 0.2% Triton X-100. DNA was precipitated in ethanol for 6 h in the presence of 0.5 M NaCl. The high molecular weight fraction was sedimented by high-speed centrifugation, and the fragmented DNA was extracted from the aqueous phase with phenol and chloroform and then precipitated with isopropanol. After resuspension in Tris–EDTA buffer, the samples were incubated with Rnase A (0.1 mg/ml) for 30 min at 37°C. Finally, DNA was electrophoresed using 1% agarose gel and visualized by ultraviolet light following ethidium bromide staining. This method can be used to semi-quantify the degree of DNA fragmentation and apoptosis, when started with the same cell number.

Detection of caspase-3 activity
Caspase-3 activity was measured by using a caspase-3 assay kit according to manufacturer’s instructions. Briefly, attached and detached PC12 cells were harvested and collected by centrifugation. The cell pellet was taken up in lysis buffer. Lysates were incubated for 1 h at 37°C with the caspase-3-specific fluorescent substrate DEVD-AFC. Fluorescence derived from release of the 7-amino-4-trifluoromethyl-coumarin (AFC) moiety was followed using a Perkin-Elmer L955 fluorimeter at 400 nm excitation and 505 nm emission.

Western blot analysis
For western blot analysis, PC12 cells treated with SO as described above were lysed at 4°C for 1 h in a lysis buffer (50 mM Tris–HCl pH 7.5, 150 mM NaCl, 1% Triton X-100), supplemented with a cocktail of phosphatase and proteinase inhibitors (1 mM sodium ortho-vanadate, 1 mM phenylmethylsulfonyl fluoride (PMSF), 10 mg/ml leupeptin, 10 mg/ml pepstatine and 10 mg/ml aprotinin). After centrifugation of the lysates at 13 000 g for 10 min, the supernatants were quantified for protein content by the Bradford method. Aliquots containing 60 µg of protein per lane were subjected to SDS–10% PAGE under reducing (5% ß-mercaptoethanol) conditions and electroblotted onto nitrocellulose membrane filters. The blots were blocked with 5% non-fat milk in 20 mM Tris–HCl, pH 7.5, 500 mM NaCl plus 0,1% Tween (TBS-T). The membranes were subsequently incubated overnight at 4°C in agitation with appropriate primary antibodies as follows: polyclonal rabbit anti-procaspase-3, polyclonal rabbit anti-Bcl-2, polyclonal rabbit anti-Bax, at a concentration of 500 ng/ml. After four times washing with TBS-T the blots were incubated for 1 h at room temperature with peroxidase-conjugated protein A (200 ng/ml; Amersham), washed four times with TBS-T, developed with ECL detection reagents (Amersham) for 1 min and exposed to X-Omat film (Eastman Kodak Co., Rochester, NY). The densitometric measurements were performed using the gel image system Fluor-S equipped with the analysis software Quantity One (Bio-Rad).

Cell shape and cytoskeletal changes studies
PC12 cells were plated on 35x10 mm tissue culture dishes with appropriate media for 12 h, then washed three times with PBS and incubated with RPMI medium and SO. Cell shape was studied over a 2-h period under a Nikon Eclipse TE 300 inverted microscope with a 40x phase-contrast objective in an attached, hermetically sealed Plexiglas Nikon NP-2 incubator at 37°C. Cell image was recorded using a COHU HP-CCD camera and image analysis was performed with a Arkon Fluo analysis system (Nikon Instrument S.p.A., Florence, Italy) and an IBM-compatible system equipped with a video card (Intel Corporation 740-854 Win 9X PV2.X).

Cytoskeletal alterations were studied on fixed PC12 cells after permeabilization with 0.1% Triton X-100 in PBS and stained for F-actin with 2 µg/ml FITC-conjugated phalloidin (F-PHD).


    Results
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 Abstract
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 Materials and methods
 Results
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 References
 
Styrene-7,8-oxide induces apoptosis of PC12 cells
To determine whether SO does induce apoptosis of PC12 cells, PC12 were exposed to various concentrations of SO (0.1–1 mM), harvested at different time points, and examined for DNA content. Figure 1Go shows the results of experiments performed to evaluate the apoptotic effect of SO. Exposure of the PC12 cells to 50 µM etoposide was carried out as a positive control; etoposide (VP-16) is a topoisomerase inhibitor commonly used as apoptosis inducer. Apoptosis was investigated by DNA distribution as revealed by flow cytometry demonstrating hypodiploid DNA (see Materials and methods). As shown in Figure 1Go, when PC12 unstimulated cells were cultured for 48 h, <10% were found to be apoptotic while exposure of cells to SO caused a dose- and time-dependent increase in apoptosis. A dose level of 0.1 mM SO had no effect on cell apoptosis, 0.8 mM SO induced apoptotic death in ~60% of cells after 48 h incubation, whereas at the same time, cells treated with 1 mM SO were found detached from the plate with an apoptotic death of 70%. Further evidence that the subdiploid population are truly apoptotic cells come from annexin V-FITC/PI assay. As can be seen in Figure 2Go, cells treated with 0.5 mM SO for 24 h were found to be 40.4% viable (region R1), 57.7% in early apoptosis stage (region R2) and 1.3% in late apoptosis stage (region R3), increasing SO concentration results in a shift from viable to early/late apoptosis stage.



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Fig. 1. Time-course effect of different SO concentrations on apoptosis of PC12 cells. Cells were incubated for the indicated concentrations and times with SO and the percent of apoptotic cells was determined by flow cytometry after PI labeling, as reported in the Materials and methods. Each value is the average of triplicate experiments ± SD. Untreated cells were used as control. In the lower panel the data obtained after 24 h incubation are reported as a representative experiment. Apoptotic cells are characterized by low DNA stainability and appear below the G1 peak in the distribution. Control, untreated cells; VP-16, etoposide 50 µM, was used as positive control.

 


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Fig. 2. Annexin V-FITC apoptosis assay on PC12 cells treated with SO. Cells were treated with 0.5, 0.8 and 1 mM SO for 24 h, labeled with annexin V-FITC and PI as described in the Materials and methods and analyzed by flow cytometry. Dot plots are representation of annexin V log fluorescence versus PI log fluorescence. Cells in region Rl (annexin Vneg-PIneg) represent living cells, cells in R2 (annexin Vpos-PIneg) early apoptotic cells, cells in R3 (annexin Vpos-PIpos) late apoptotic cells and cells in R4 (annexin Vneg-PIpos) those with a damaged membrane only. Untreated cells and cells treated with VP-16 (50 µM) were used as negative and positive control respectively. The data shown are representative of three different experiments.

 
The data derived from FACScan analysis were confirmed by determination of apoptotic cells by DNA fragmentation observed in gel electrophoresis. In the presence of SO a typical laddering was seen in PC12 cells but not in the control (Figure 3Go). Time-course studies revealed the presence of early changes indicative of apoptosis development in the cell population 8 h after SO treatment.



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Fig. 3. Detection of DNA fragmentation by agarose gel electrophoresis. PC12 cells were exposed to SO at the indicated concentrations for 8–48 h, and then cells, including floating and adherent ones, were collected. Fragmented DNA was extracted from cells as described in the Materials and methods and electrophoresed on 1% agarose gels, stained with ethidium bromide, and photographed. C, untreated cells; VP-16 (12 h 50 µM), positive control; M, DNA molecular weight marker. Three experiments were performed with similar results.

 
Styrene-7,8-oxide-induced apoptosis is mediated by caspase-3 activation
We investigated the molecular mechanisms of apoptosis induced by SO in PC12 cells. The activation of caspases is a critical event in the proteolytic cascade elicited by apoptotic stimuli; in particular caspase-3 is an effector caspase that plays a role in cell death induced by a variety of stimuli (30). Consistent with an induction of apoptosis, we found that caspase-3 was activated in SO-treated PC12 cells (Figures 4 and 5GoGo). The activity of caspase-3, during SO-induced apoptosis, was examined as a decrease in pro-enzyme level using western blotting analysis. Time-course experiments established that SO 0.8 mM induced a rapid decrease of proenzyme level starting 6 h after stimulation (Figure 4AGo). Maximal activation was observed 12 h after incubation and remained steady for the whole period of observation. In fact treatment for 24 or 48 h, with SO 0.5–0.8 mM markedly reduced the amount of detectable proenzyme levels of caspase-3 (Figure 4BGo).



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Fig. 4. Western blot analysis of procaspase-3 expression in PC12 cells. Cells were treated with SO at the indicated concentrations and times and subjected to western blot analysis using procaspase-3-specific antibody. The activation of caspase-3 was detected as a decrease in the proenzyme amount. (A) Time-course of 0.8 mM SO on caspase-3 activation; VP-16 treated cells (12 h, 50 µM) was used as positive control. (B) Effect of SO (0.5 and 0.8 mM) and of VP-16 (50 µM) for 24 and 48 h on procaspase-3 expression in PC12 cells. Untreated cells (C) were used as negative control. The data shown are representative of three different experiments.

 


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Fig. 5. Detection of caspase-3 activity in apoptotic PC12 cells. Cells were treated with various concentrations of SO for 12 h and harvested in lysis buffer. Enzymatic activity of caspase-3 was determined by protease assay by using the specific DEVD-AFC chromogenic substrate as described in the Materials and methods. The release of the chromophore AFC was monitored spectrophotometrically (505 nm). Control, untreated cells; VP-16 (50 µM), positive control. Values are represented as mean ± SD from three independent experiments.

 
To further investigate and quantify the proteolytic activity of caspase-3, we performed an in vitro assay based on the proteolytic cleavage of the peptide substrate DEVD-AFC by caspase-3 into the fluorophore AFC. Treatment with SO for 12 h caused an increase in caspase-3 activity in a dose-dependent manner (Figure 5Go). No effect on caspase-3 activity was seen with PC12 cells alone. These results suggest that apoptosis, induced by SO, is associated with caspase-3 activation.

Expression of Bcl family proteins in SO-treated PC12 cells
We next examined whether SO-induced apoptosis involves modulation of Bcl-2 family proteins. Time-course experiments were carried out to detect the protein level of apoptosis-related genes in 0.5 and 0.8 mM SO-treated PC12 cells. We found that Bcl-2 protein level was significantly reduced from 8 to 48 h after exposure to 0.5 mM SO, concomitantly Bax protein levels were increased (Figure 6Go). Consistent with the time-course of caspase-3 activation, treatment with 0.8 mM SO induced a more rapid modulation of Bcl-2 and Bax from 6 to 12 h (Figure 6Go). The ratio of anti-apoptotic factor to pro-apoptotic factor in the cell is assumed to be a critical mechanism in maintaining normal homeostasis. The net result following SO treatment was a time-dependent decrease in the ratio Bcl-2/Bax, thus shifting the homeostasis toward a preference for apoptosis (Figure 6Go). These results indicate that some of Bcl-2 family proteins are modulated in SO-induced apoptosis.



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Fig. 6. Protein levels of Bcl-2 and Bax in SO-treated PC12 cells. Cells were exposed to 0.5 or 0.8 mM SO for different time intervals. Protein levels were determined by western blot analysis. Band intensity of Bcl-2 and Bax were determined by densitometric analysis using the gel image software Quantity One (Bio-Rad). The Bcl-2/Bax ratio is an average calculated on the results from three separate experiments.

 
Cell shape and cytoskeletal changes
Cytoskeleton plays a critical role in the regulation of various cellular processes such as cell shape, cell division and apoptosis. Therefore, we evaluated whether SO induces shape changes and modifies the normal distribution of actin containing stress fiber. PC12 cells normally spread within 1–3 h after plating, extend over 24 h of culture, appear largely polygonal in form with visible dendrites and attach to each other (Figure 7BGo). In the presence of 0.5 mM SO some of the cells cultured for 1 h at 37°C remained rounded, whereas other cells took on a partially spread, polygonal shape (Figure 7DGo); exposure of cells to 0.8 mM SO for 1 h led to mostly rounded cells with loss of dendrites and separated from each other (Figure 7FGo). On the other hand, cell treated with SO showed a good viability, measured by trypan blue exclusion assay, as shown in Table IGo.



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Fig. 7. Micrographs representative of shape change of PC12 cells. SO induces cell shape as observed by comparison of the time 0 (A, C and E) and 1-h frames (B, D and F). (A and B) Unstimulated PC12 cells; (C and D) PC12 cells stimulated with 0.5 mM SO; (E and F) PC12 cells stimulated with 0.8 mM SO. Three experiments were performed with similar results. Original magnification, x400.

 

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Table I. Cell viability estimated by trypan blue analysis
 
To determine whether the cytoskeleton plays a role in this SO-dependent change in cell shape, we studied, by fluorescence microscopy, actin cytoplasmatic distribution in SO-treated PC12 cells after labeling with FITC-conjugated phalloidin. The normal actin fibers distribution in untreated cells shows that F-actin forms fine-meshed networks (Figure 8AGo). After 1 h stimulation with 0.5 mM SO at 37°C cells lose actin cytoskeleton organization, which is characterized by F-actin depolymerization, as evident by breaks along fibers (Figure 8BGo), treatment with 0.8 mM SO led to a complete loss of fibers (Figure 8CGo).



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Fig. 8. SO induces stress fiber dissolution-disassembly. Micrographs representative of F-actin distribution in fixed and permeabilized PC12 cells after incubation for 1 h in the following experimental conditions: (A) unstimulated PC12 cells; (B) PC12 cells stimulated with 0.5 mM SO; (C) PC12 cells stimulated with 0.8 mM SO. Three experiments were performed with similar results. Original magnification, x400.

 

    Discussion
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 Abstract
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 Materials and methods
 Results
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 References
 
In the present study the ability of SO to induce inhibition of cell growth and apoptosis in the neuronal PC12 cell line was evaluated; furthermore, a set of experiments was carried out to examine the molecular and cellular mechanisms occurring during the apoptotic process triggered by SO. Although one must be aware that the cellular response to drug exposure may depend on the cell type, the PC12 cell line seems a suitable cell model as the primary target for the SO toxicity is the nervous system.

Styrene is one of the most important plastic monomers worldwide. SO, the major metabolite of styrene in vivo, is classified as probably carcinogenic in human and animals (2,6). Nevertheless, although the toxic effects of SO have been extensively documented in human lymphocytes (9,10,14), the molecular mechanisms responsible for SO-induced cell death are still unknown.

We found that SO induced growth inhibition and massive apoptosis, as revealed by FACS analysis using DNA fragmentation and hypodiploidism as an indirect marker of apoptosis and confirmed by annexin V-FITC/PI assay (Figures 1–3GoGoGo). Increases in the proportion of apoptotic cells occurred in a dose- and time-dependent manner and at a relatively low range of concentrations (0.1–1 mM).

It has been demonstrated previously (31) that in PC12 cells SO stimulated the formation of alkali-sensitive sites and DNA single-strand breaks at doses of 0.03 and 0.1 mM. These DNA lesions were fully reversible following an exposure to 0.03 mM SO, whereas strand breaks induced by 0.1 mM were only partially repaired. Interestingly, in our study it was found that SO at a concentration of 0.03 mM did not induce growth inhibition, apparent cytotoxicity and apoptosis. Under this point of view, apoptosis, an intrinsic suicide program of the cell, represents a mechanism to limit tissue injury.

Furthermore, we explored the molecular mechanism associated with SO-induced apoptosis. Apoptosis is mediated by multiple pathways that involve a complex array of biochemical regulators and molecular interactions. Although the upstream signaling of apoptosis is uncertain, the family of cysteine proteases, caspases, seems to play a key role in the apoptotic process. In particular, caspase-3 is the major effector caspase responsible for cellular proteolysis associated with the apoptotic cascade (23).

Our results (Figures 4 and 5GoGo) indicate that SO induced a significant increase of the caspase-3 activity in PC12 cells compared with the control cells. In addition, and consistently, as shown in Figure 6Go, SO reduced Bcl-2 and stimulated Bax. Moreover, the observation that SO induced a loss of cytoskeletal organization and a rapid cell rounding starting 1 h after stimulation (Figure 7Go), suggests the possibility that the chemical trigger is a more complex apoptotic response. In particular, we found that F-actin was depolymerized in >90% of the PC12 SO-treated cells not yet detached from substrate (Figure 8Go) and that the disruption of the F-actin cytoskeletal network occurred long before caspase-3 activation. However, it is largely recognized that actin disassembly and cytoskeleton rearrangement may activate apoptosis (32,33). In addition, HIV-1 vpr protein protects from anoikis by promoting actin microfilament assembly (34) and microtubule disassembly similarly induces apoptosis in fibroblasts (35). More recently, a possible causal link between depolymerization of F-actin and cell death has been established by Korichneva and Hammerling (36): in murine EL4 T lymphoma cells, disorganization of the actin cytoskeleton by cytochalasin B renders the cells more sensitive to anhydroretinol-induced cell death; conversely, stabilization of the actin filaments by jasplakinolide promotes cell resistance to apoptosis (36).

Taken together, these results suggest that during SO-induced apoptosis in PC12 cells, the loss of actin cytoskeletal organization occurs independently of caspase-3 activation, and represents an early step in the apoptotic pathway. However, it remains to be elucidated whether this pathway is common to other cell lines, in fact a different apoptotic pathway could be responsible for the differing sensitivity found among species and organs toward tumor promotion elicited by SO.


    Notes
 
4 To whom correspondence should be addressed Email: nicola.sannolo{at}unina2.it Back


    References
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 

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Received May 31, 2002; revised November 19, 2002; accepted November 26, 2002.