Cell Cycle Inhibition by Sodium Arsenite in Primary Embryonic Rat Midb
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《毒物学科学杂志》
Department of Environmental and Occupational Health Sciences, University of Washington, Seattle, Washington 98195
Center for Ecogenetics and Environmental Health and Institute for Risk Analysis and Risk Communication, Seattle, Washington 98105
Center on Human Development and Disability, Seattle, Washington 98195
ABSTRACT
Arsenite (As3+) exposure during development has been associated with neural tube defects and other structural malformations, and with behavioral alterations including altered locomotor activity and operant learning. The molecular mechanisms underlying these effects are uncertain. Because arsenic can cross the placenta and accumulate in the developing neuroepithelium, we examined cell cycling effects of sodium arsenite (As3+ 0, 0.5, 1, 2, and 4 μM) on embryonic primary rat midbrain (gestational day [GD] 12) neuroepithelial cells over 48 h. There was a concentration- and time-dependent As3+-induced reduction in cell viability assessed by neutral red dye uptake assay but minimal apoptosis at concentrations below 4 μM. Morphologically, apoptosis was not apparent until 4 μM at 24 h, which was demonstrated by a marginal but statistically significant increase in cleaved caspase-3/7 activity. Cell cycling effects over several rounds of replication were determined by continuous 5-bromo-2'-deoxyuridine (BrdU) labeling and bivariate flow cytometric Hoechst-Propidium Iodide analysis. We observed a time- and concentration-dependent inhibition of cell cycle progression as early as 12 h after exposure (0.5 μM). In addition, data demonstrated a concentration-dependent increase in cytostasis within all cell cycle phases, a decreased proportion of cells able to reach the second cell cycle, and a reduced cell cycle entry from gap 1 phase (G1). The proportion of affected cells and the severity of the cell cycle perturbation, which ranged from a decreased transition probability to complete cytostasis in all cell cycle phases, were also found to be concentration-dependent. Together, these data support a role for perturbed cell cycle progression in As3+ mediated neurodevelopmental toxicity.
Key Words: sodium arsenite; cell cycle; embryonic neuroepithelial cells; development; neurotoxicity.
INTRODUCTION
Arsenic (As) is a naturally occurring element that exists in several oxidative states but it is the pentavalent (arsenate, As5+) and trivalent (arsenite, As3+) forms that are most prevalent in the environment and have toxicological significance. Of the two forms, arsenite is considered a more potent developmental toxicant (Willhite and Ferm, 1984). Trivalent arsenicals readily react with thiol-containing molecules interfering with various biochemical processes resulting in toxicity (Scott et al., 1993). The U.S. Food and Drug Administration has calculated the mean daily intake of inorganic arsenic (arsenite and arsenate) to be approximately 0.5 g/kg body weight for adults including sources from air, water, and food. Because of the extensive distribution and associated toxicity of As, contamination of drinking waters represents a worldwide problem. In response, the Environmental Protection Agency (EPA) has established a drinking water standard of 10 ppb or a maximum contaminant level of 0.010 mg/l (EPA, 2001).
Inorganic arsenic has been shown to freely cross the placenta and accumulate in the embryonic neuroepithelium (Lindgren et al., 1984) and has been linked to various embryotoxic outcomes, including neural tube defects, reduced organ and body weight, malformations, and fetal death (Willhite and Ferm 1984; Wlodarczyk et al., 1996). Acute exposure has been associated with overt malformation in the offspring and nearly fatal maternal toxicity, implying that environmentally relevant levels of As3+ are not toxicologically significant for developmental structural defects (DeSesso et al., 1998). However, chronic exposures of As3+ (maternal dose estimated at 2.9–4.2 mg/kg/day) demonstrated a significant impact on neurological function in developing and postnatal animals, suggesting neurodevelopmental risk with low concentrations (Rodriguez et al., 2002). In addition, studies in neonatal (P0) neuronal cells exposed to less than 5 μM of As3+, found decreased cell viability, altered cell growth, and apoptosis (Chattopadhyay et al., 2002a,b) reflecting mechanisms by which low doses of As may lead to neurodevelopmental toxicity. Developmental impact of As is also apparent in studies that have shown transplacental carcinogenicity with in utero exposure (Waalkes et al., 2004).
Both neurons (Namgung and Xia, 2001) and astroglia (Jin et al., 2004) are more sensitive to As3+ as compared to As5+, with exposure resulting in morphological changes and decreased cell viability. In addition, a significantly lower dose of As3+ is sufficient to induce apoptosis in neuronal cells as compared to non-neuronal cells illustrating the sensitivity of the nervous system to this metal (Wang et al., 1996). Mechanistically, As3+ has been shown to interfere with cell regulatory control via the induction of p53 and bcl-2 (Wlodarczyk et al., 1996). P53 is a critical regulator of the cell cycle and has been shown to undergo a dose- and time-dependent induction in response to both As3+ (Filippova and Duerksen-Hughes, 2003) and arsenic trioxide (As2O3) (Yeh et al., 2003). Induction of p53 in response to As2O3 exposure has been associated with subsequent cell cycle arrest at gap 2 phase (G2)/mitosis (M) (Zhao et al., 2002). Both As2O3 (Li et al., 2003) and As3+ (Huang et al., 2002) have been shown to induce p21, a critical cell cycle regulatory protein. As with p53, arsenic induced p21 has also been associated with blocking cell cycle progression at G2/M (Chou and Huang, 2002) as well as G1/S (synthesis phase) (Park et al., 2000). Other thiol reactive metals, such as methylmercury (MeHg) (Ou et al., 1999), similarly induce p21 (Faustman et al., 2002) and lead to G2/M arrest (Gribble et al., 2003), suggesting a common cellular response between the metals involving cell cycle perturbation.
Cell cycle arrest at G2/M induced by As3+ (Ma et al., 1998) and As203 (Li et al., 2003) has been associated with subsequent apoptosis indicating the latter may occur due to As-mediated cell cycle inhibition. Namgung and Xia (2001) have reported As induced apoptosis in studies with primary cerebellar and cortical neurons exposed to environmentally relevant doses of As3+ (Namgung and Xia, 2001). These studies highlight the impact of As on cell cycle progression and apoptosis and their potential link to As associated neurodevelopmental effects.
Here we present results of recent experiments examining the effect of low-dose concentrations of As3+ on cell cycle kinetics and dynamics in primary embryonic rat neuroepithelial cells during the critical "window of susceptibility" when rapid neurogenesis is occurring (Copp et al., 1990). Using continuous bromodeoxyuridine (BrdU) labeling in vitro, Hoechst-Propidium Iodide DNA staining, and bivariate flow cytometry, we were able to monitor individual cell cycle progression through several cell division rounds in primary rat midbrain cell cultures with continuous As3+ exposure. Our findings demonstrate a dose-responsive delay in cell cycle progression in all cell cycle phases induced by As3+ in primary embryonic neuroepithelial cells and suggest that cell cycle arrest may contribute to the neurodevelopmental toxicity observed with this metal.
MATERIALS AND METHODS
Cell culture.
Embryonic rat midbrain neuroepithelial cell cultures were prepared according to our previously established protocol (Flint, 1983; Ponce et al., 1994b; Ribeiro and Faustman, 1990; Whittaker and Faustman, 1991, 1992; Whittaker et al., 1993) with slight modifications. Gravid uteri were removed from pregnant (GD 12) Sprague-Dawley albino rats (Charles River, Hollester, CA) and placed in Earl's Balanced Salt Solution (EBSS, Gibco BRL, Grand Island, NY). Midbrain sections were dissected from individual embryos and incubated in calcium- and magnesium-free EBSS (CMF-EBSS, Gibco BRL, 37°C, 20 min). Midbrain sections were treated with trypsin (1% w/v, Gibco BRL) in CMF-EBSS (pH 7.2–7.4, 37°C, 20 min). Enzymatic digestion was stopped with addition of culture media containing 10% fetal bovine serum (Qualified, Gibco BRL), 2 mM L-glutamine (200 mM stock, Gibco BRL), 100 units/ml penicillin, 0.1 mg/ml streptomycin (Gibco BRL), and 88% Ham's F-12 nutrient mixture (Gibco BRL, stored at 4°C and kept from light). A single cell suspension of the neuroepithelial cells was prepared by mechanical dissociation of the cells with a fine bore (0.7 mm) glass Pasteur pipette followed by filtration (10 mm nylon mesh) to remove debris and clumps. The cell concentration was adjusted to 5 x 106 cells/ml with addition of culture media. Five 10-μl aliquots of cell suspension were plated per 35-mm Primaria-coated culture dish (Falcon). The cultures were incubated (5% CO2/95% air, 37°C, 100% humidity) for 2 h to allow cell attachment. After the attachment period, 2 ml of culture media containing 5-bromo-2'-deoxyuridine (80 μM, BrdU, Sigma) and sodium arsenite (0–4 μM) was added. Under these conditions, these cultures have been demonstrated through immunocytochemistry staining to consist of a nearly pure population of proliferating, immature neuronal precursor cells that differentiate into mature neuron over time (Whittaker et al., 1993). Sodium arsenite (As3+ NaAs02, Fischer Chemical Corporation) was prepared as a working stock (100 mM) solution in sterile water and stored at –20°C. Treatment was conducted in a Class II Type B2 hood (The Baker Company Inc., Sanford, ME).
Cell harvest.
Cell harvest was conducted at serial time points post-treatment. Culture media containing BrdU and arsenic was removed, cells were rinsed with CMF-PBS (37°C), and then incubated with 500 μl CMF-PBS containing 0.05% w/v trypsin and 0.02% EDTA (Fischer Scientific, 5 min, 37°C). Trypsinization was stopped by the addition of 500 μl culture media and the resulting cell suspension was transferred to 2 ml polyethylene tubes (Falcon) for centrifugation (1500 x g, 15 min, RT). After centrifugation, the supernatant was removed and 450 μl of Hoechst buffer [1.2 μg/ml Hoechst 33258 (Molecular Probes, Eugene OR), 0.146 M NaCl, 0.1 M TRIS Base, 0.5 mM MgCl2, 0.2% BSA, 0.1% NP-40 (Sigma, St. Louis, MO) in dd H20 and 50 μl of 10% dimethyl sulfoxide I were added to each sample. Cells were transferred to labeled amber 1.5 ml microfuge tubes (VWR Scientific Products, West Chester, PA) and frozen at a rate of –1°C/min to –20°C until analysis. Cell treatment and harvest procedures were conducted under sodium lamp illumination to minimize BrdU-induced photosensitization.
Flow cytometry.
BrdU-Hoechst analysis was performed according to established procedures (Mendoza et al., 2002; Ponce, 1994). Cells were thawed in a water bath (37°C), and 10 μg/ml Propidium Iodide (PI, Sigma) was added to each tube. Cells were stored on ice, protected from light for a minimum of 30 min prior to analysis. Flow cytometric cell cycle analysis was performed on a Coulter Epics Elite flow cytometer using a UV laser emitting 335 nm ultraviolet light. A 450/35 nm band pass filter was used to collect the Hoechst 33258 fluorescence. Hoechst 33258 binds to AT-rich regions in DNA and its fluorescence is proportionally quenched as cycling cells incorporate BrdU. An air-cooled 15 mW argon ion laser (Cyonics, San Jose, CA) emitting 488 nm light was used with a 590 nm long pass filter to excite and collect PI fluorescence, respectively. The flow rate was maintained at less than 200 cells per second. A minimum of 20,000 events per treatment was analyzed. Samples were protected from light throughout the experiment. Cell cycle analysis was quantified using MultiPlus Software (Phoenix Flow Systems, San Diego CA). Clumps and doublets were excluded from the analysis by electronic gating. Analysis was conducted to determine the proportion of cells in each cell cycle phase (G0/G1, S, G2/M) from successive rounds of cell division.
Assessment of cell morphology.
Following chemical treatment, cellular morphology was examined at 24 and 48 h, with a Nikon inverted microscope equipped with phase-contrast optics (Nikon, Tokyo, Japan). All images were captured and digitized using a Coolsnap Camera (Roper Scientific, Inc) and processed using Photoshop software (v. 5.5, Adobe, Seattle, WA).
Cytotoxicity assay.
Cytotoxicity assessments were conducted using the neutral red (NR) dye uptake assay (Borenfreund and Puerner, 1985), which determines cell viability by assessing lysosomal accumulation of NR dye. Cytotoxicity assessments were done with continuous exposure of the primary neuroepithelial cells to As3+ over 48 h (Komissarova et al., 2005). Briefly, cell culture plates were removed from the incubator at each sampling time and the media was aspirated. Cells were washed once with CMF-PBS. Fresh media containing 50 μg/ml NR was added to each 35 mm dish. After incubation (3 h, 37°C, 5% CO2), the cells were washed with CMF-PBS and the NR was eluted with 2 ml of 0.5% acetic acid/50% ethanol solution for 1 h at room temperature. Four 200-μl aliquots of NR solution were removed from each dish and placed into replicate wells of a micro titer plate. After rapid agitation of the plate reader, light absorbance was measured at 540 nm. A correction for background absorbance was conducted by subtracting media background absorbance from each well. Using these background corrected values, the relative cytotoxicity was estimated as the ratio of average absorbance between As3+ treated cultures relative to untreated control cultures.
Preparation of cell extracts.
At the stated time points, cells were washed twice with 2.5 ml ice cold CMF-PBS followed by the addition of 50 μl of cell lysis buffer (Sidhu and Omiecinski, 1998) containing additional phosphatase and protease inhibitor cocktails (Calbiochem). Cells were harvested by scraping in cell lysis buffer and then placed on ice. After storage at –80°C, all extracts were homogenized by sonication and then centrifuged (16,000 x g, 10 min, 4°C) to remove insoluble material. The resulting supernatant (cell extract) was gently removed, and total protein was determined using the Bio-Rad kit (Pierce, IL).
Apoptosis assessment.
Apoptosis was determined in cell extracts by measuring the functional activity of caspase-3 using a caspase-specific fluorogenic substrate. The activity of cleaved-caspase 3 was measured in a micro titer plate by a fluorometric assay, using Ac-DEVD-AMC as the caspase-specific substrate. Briefly 10 μg of cell extract was added in duplicate in 96 well plate format. Reaction buffer was added and incubated at 37°C for 2 h. The enzyme-catalyzed release of 7-amino-4-methyl coumarin (AMC) was measured by a fluorescence microplate reader at excitation 360 nm and emission 460 nm. Fluorescent units were converted to pmol of AMC released per μg of protein and incubation time (2 h) using a standard curve generated with known serial dilutions of AMC. The absolute activities due to As3+ treatments relative to untreated controls were converted by expressing the former as a percentage of control.
Statistical analysis.
ANOVA was performed using the General Linear Model (GLM) General Factorial procedure. Repeated measures of ANOVA were conducted to evaluate differences between As3+-treated and control cell proliferation. Post-hoc analysis of pair-wise differences between treatment groups was conducted using Fisher's PLSD. A p-value 0.05 was considered statistically significant. All statistical analyses were performed using JMP for Windows Release 5.0 (SAS Institute, Cary, NC).
RESULTS
We examined As3+-induced cytotoxicity and cell cycle inhibition in primary cultures of gestational day (GD) 12 rat midbrain neuroepithelial cells over 12–48 h of continuous exposure with PBS or sodium arsenite (0.5–4 μM). See Figure 1. Cells cultured for flow cytometric analysis of cell cycle progression were simultaneously treated with bromodeoxyuridine (BrdU). Use of continuous BrdU labeling, Hoechst-ethidium bromide DNA staining, and bivariate flow cytometry allowed analysis of individual cell cycle progression through several rounds of cell division.
Concentration-Dependent As3+-Induced Cell Loss
A time- and concentration-dependent decrease in cell number was observed with continuous exposure of the primary neuroepithelial cells to As3+ over 48 h. For example, whereas there was no appreciable cell loss upon continuous exposure to As3+ 2 μM after 24 h of exposure, dye-uptake was reduced to approximately 60% of control cells at 48 h. In contrast, exposure to 4 μM reduced dye-uptake to 40% of control cells after 24 h and approximately 20% of control cells after 48 h of exposure to As3+. Because the NR dye-uptake assay is sensitive to both the cell loss associated with cell death and reduced cell proliferation when compared against untreated control cultures, these results reflect the aggregate effects of As3+ on cell production and viability.
As3+ Inhibited Cell Cycle Progression
Asynchronous primary rat midbrain neuroepithelial cells were incubated continuously with As3+ for up to 48 h and analyzed for cell cycle progression using BrdU-Hoechst bivariate flow cytometry. Representative BrdU-Hoechst flow cytograms identifying the distribution of cells in the first and second round of cell division at 24 h (Panel A) and 48 h (Panel B) are presented in Figure 2. Because Hoechst staining is inversely proportional to BrdU incorporation, cells in G2/M of the first cell division round will have the highest Hoechst and propidium iodide fluorescence; these cells are labeled G2/M0 in Figure 2A. Cells remaining in the first round G0/G1 phase during the BrdU labeling period are labeled as and cells that were able to complete one cell cycle and reach a new G0/G1 during the BrdU labeling period are labeled as in Figure 2.
Inspection of the cytograms from treated cells revealed a concentration- and time-dependent inhibition of cell proliferation in response to As3+ exposure (Fig. 3). Whereas 55 ± 5% of control cells successfully completed at least one round of cell division over 48 h of cell culture, only 46 ± 5% of cells treated with 0.5 μM As3+ reached the second round of cell division over this time period (Fig. 4). Among cells treated with 4 μM As3+, only 8 ± 4% reached a second round of cell division over 48 h of culture.
The cell cycle inhibition occurred in all cell cycle phases, and there was an increased probability of cytostasis with increasing As3+ concentration regardless of cell cycle phase at which As3+ exposure occurred in the asynchronous culture (Fig. 5). This conclusion is based on an evaluation of the proportion of cells that failed to take up BrdU during the culture period. Because BrdU incorporation during DNA synthesis results in a diminished Hoechst fluorescence (without affecting PI fluorescence), it is possible to discriminate the cells that have remained in their original cell cycle stage (that approximate lie on a 45° line on a bivariate Hoechst-PI scatter gram) from those that have progressed through the cell cycle in the presence of BrdU (see Fig. 2). Evaluation of the relative proportion of cytostatic cells as a function of As3+ concentration and time revealed a concentration-dependent increase in the proportion of cytostatic cells in Fig. 5. For example, whereas approximately 43 ± 4% of control cells were in their original cell cycle position (relative to the start of dosing), approximately 60 ± 2% of cells exposed to 1 μM As3+ and 91 ± 2% of cells exposed to 4 μM As3+ remained in their original cell cycle position after 48 h of culture. Evaluation of the untreated control (0 μM As3+) cells demonstrate approximately 43% of cells were in the G0-phase of the cell cycle at 48 h of culture (Fig. 6), consistent with previous observations regarding the proportion of non-cycling, presumably differentiating cells, in this cell culture system (Ponce et al., 1994). The concentration-dependent increase in the proportion of noncycling cells exposed to 0.5–2 μM As3+ can be explained by a predominant increase in the number of cells that do not enter the cell cycle and remain in G0/G1 (see Fig. 6). In contrast, not all of the cytostasis observed upon exposure to 4 μM As3+ is attributable to G0/G1 inhibition, but also includes inhibition in S- and G2/M (see Fig. 6). Specifically, whereas 91 ± 2% of cells exposed to 4 μM As3+ remained in their original cell cycle position after 48 h of culture (Fig. 5), only 76 ± 6% of these cells were in (Fig. 6).
A complex relationship between As3+ exposure, relative to concentration and duration, was observed with respect to the proportion of cytostatic cells relative to the total population of cells. For example, while it is possible to leave the original G0/G1 population (identified as Fig. 2) during 48-h of culture, thereby reducing the relative abundance of this cell type in the culture, it is not possible to repopulate this cell population. This ability to discriminate "original" G0-phase cells from "new" G0/G1-phase cells ( Fig. 2) arises from the incorporation of BrdU during the DNA synthesis, which results in a distinct population with reduced Hoechst fluorescence. As a result, any apparent increase in the relative proportion of cells in phase cells must be explained by a differential loss in the cycling cell population relative to the population. We observed an apparent increase between 24 and 48 h in the relative abundance of cytostatic cells upon exposure to 1 μM As3+ as reflected in the increased proportion of cytostatic cells over time (Fig. 5), suggesting a greater proportional loss in the cycling population relative to arrested cells. Evaluation of the cell cycle phase distribution demonstrated that the majority of cytostatic cells were in (see Fig. 6).
Apoptosis
To determine whether the observed loss in cell viability and cell number was associated with the induction of apoptosis, we measured cleaved caspase 3/7 activity using a fluorogenic and caspase-specific substrate assay (Fig. 7). The results demonstrate that a marginal concentration-dependent induction of apoptosis was indeed observed after 12 and 24 h post-treatment with As3+. However, a statistically significant induction of cleaved caspase-3/7 activity was only observed at the highest As3+ concentration examined (4 μM) at both time points with a marginal but statistically insignificant induction at 2 μM. After 48 h no statistically significant evidence of apoptosis was observed at As3+ 2 μM; this is presumed to result from the detachment/loss of cells that had undergone apoptosis at earlier time points. These results are consistent with the morphological evidence (Fig. 8) supporting significant cell loss and overt toxicity. Together these findings demonstrate that As3+-induced growth arrest occurs independent of apoptosis and may be more related to a generalized cytostasis.
DISCUSSION
The toxicity of inorganic arsenic (As) is well established with known human impacts including skin lesions, peripheral vascular disorder, peripheral neuropathy, liver injury, and cancer (Gebel, 2000). Arsenic has also been implicated as a possible teratogen (Beaudoin, 1974). The degree to which gestational As exposure affects neurodevelopment, however, remains controversial and poorly understood (DeSesso et al., 1998). Furthermore, the impact of environmentally relevant doses of As remains uncertain and requires further investigation to elucidate the mechanisms involved and their subsequent role in developmental toxicity.
Mechanistically, As-mediated effects on neurodevelopment have been linked to cell cycle perturbation through the up-regulation of key checkpoint proteins. A significant up-regulation of p53 and bcl-2 was demonstrated in neural tubes on gestational day (GD) 9.0 following maternal exposure to arsenate at 40 mg/kg (Wlodarczyk et al., 1996). The tumor suppressor protein, p53, is a key regulator of the cell cycle and is responsible for initiating cell cycle arrest in response to DNA damage (Bates and Vousden, 1996). Arsenite (As3+) has been shown to induce p53 in a dose- and time-dependent manner in several cell types (Filippova and Duerksen-Hughes, 2003; Vogt and Rossman, 2001), with induction levels and toxicity dependent on the cell system (Salazar et al., 1997). Previous studies have demonstrated that both As2O3 and As3+-mediated p53 induction is followed by cell cycle arrest at G2/M (Yih and Lee, 2000; Zhao et al., 2002). Cell cycle inhibition by As has also been associated with a subsequent apoptotic response (Chou and Huang, 2002; Park et al., 2003). Between GD 8 and GD 10 in the rat, the brain and spinal cord are developing via rapid proliferation and subsequent differentiation of neuroepithelial cells (Copp et al., 1990). Arsenic-induced cell cycle inhibition and potential apoptosis at this stage of development, therefore, could have profound impact on the development of these systems.
The present study examined the impact of concentrations (0.5–4 μM) of As3+ on cell cycle kinetics of primary embryonic rat midbrain (GD 12) neuroepithelial cells. The embryonic midbrain neuroepithelial cell culture system has been used extensively by our group and others to examine the effects of toxicants on proliferation and differentiation of neuronal precursor cells on GD 12 (Flint, 1983; Ponce et al., 1994; Ribeiro and Faustman, 1990; Whittaker and Faustman, 1991, 1992; Whittaker et al., 1993). This culture system has been used to evaluate the effects of chemicals on cell viability, [3H] -aminobutyric acid uptake (George and Gabay, 1968), and hematoxylin-stained neurites (Flint and Orton, 1984). Whittaker et al. characterized neuronal cells in these cultures that specifically stained for neuronal markers A2B5 (GQ ganglioside), -aminobutyric acid (GABA), microtubule-associated protein (MAP2, MAP5), neuron-specific enolase (NSE), neural cell adhesion molecule (N-CAM), and tau after 5 days in culture (Whittaker et al., 1993). The cells were non-immunoreactive for glial fibrillary acidic protein (GFAP) (Whittaker et al., 1993). This system has also been successfully utilized to characterize changes in cell cycling in response to exogenous factors such as alkylating agents, albendazole, methylmercury, chlorpyrifos, and serotonin (Cosenza and Bidanset, 1995; Menegola et al., 2004; Ponce et al., 1994; Ribeiro and Faustman, 1990; Seeley and Faustman, 1995; Whittaker and Faustman, 1991).
We observed a concentration- and time-dependent As3+-induced cell cycle inhibition. This response was seen as early as 12 h after exposure and was manifested as decreased entry into the cell cycle from the G0-phase (Fig. 4), an increased proportion of cytostatic cells (Fig. 5), and decreased proportion of cells reaching the second cell cycle (Fig. 4). At 24 h there was a delay in progression from S-phase to G2 and at 36 h cell cycle entry from G0/G1 was reduced. Analysis of cell cycle-phase distributions suggests a possible accumulation in G2/M (see Fig. 6). Results from the present study indicate that the As3+-associated reduction in the number of cells reaching the second cell cycle after 48 h, relative to the control, is associated with both cell cycle inhibition and resultant cytostasis. Whether these cells subsequently underwent apoptosis is still inconclusive since only a marginal increase in leaved caspase-3 activity was observed. Morphological evaluations, however, demonstrated cellular stress occurring at concentrations 4.0 μM consistent with cells exhibiting characteristics of both apoptosis and necrosis. An apoptotic response does not mandate the correlation between morphology and caspase-3 activity as previously demonstrated in cerebellar granule cells (Dare et al., 2001). Although exhibiting apoptotic morphology, the cells failed to show an increase in cleaved caspase activity upon treatment with either MeHg or H2O2. An alternate study found that intermediate concentrations of As3+ (5–10 μM) induced cytostasis while concentrations >10 μM lead to caspase-3-dependent apoptotic cell death irrespective of cell cycle phase (McCabe et al., 2000). Our results corroborate that cell cycle arrest is a more sensitive endpoint than apoptosis to As3+ exposure as we saw changes in cell cycle kinetics following treatment with 1.0 μM (12 h–48 h) in the absence of caspase activation or apoptotic morphology.
Our previous studies with another neurodevelopmental toxicant, methylmercury (MeHg), support a role for metal induced cell cycle perturbations as demonstrated by the changes to key cell cycle proteins such as p21 (Ou et al., 1999) and/or GADD genes (Ou et al., 1997), and accumulation of cells in G2/M phase that was dependent on p53 and p21 genotype (Gribble et al., 2005; Mendoza et al., 2002). Similar associations have been made with As2O3 exposure where the up-regulation of GADD45, GADD153 (Li et al., 2003), and p21 (Park et al., 2000) was associated with arrest and subsequent apoptosis. The evident involvement of cell cycle proteins speaks to the critical role that cell cycle regulation plays in the overall mechanism of As-mediated neurodevelopmental toxicity.
As demonstrated by the present study, low-dose As3+ exposure within primary embryonic rat neuroepithelial cells resulted in a decrease in cell viability. Specifically, there was a concentration- and time-dependent decrease in cell viability upon exposure to As3+ concentrations >2 μM for 24 and 48 h. Studies in cortical neurons (isolated P0, cultured 6 days prior to treatment) (Namgung and Xia, 2000) and in cerebellar neurons (isolated P6-8, cultured 7 days) (Namgung and Xia, 2001), observed a similar reduced cell viability in response to As3+ but at somewhat higher concentrations (5–10 μM). In addition, these studies demonstrated a dose-dependent, stress-induced apoptotic response involving the activation of the stress proteins, JNK and P38 MAPK, suggesting a role for the stress response in neuronal apoptosis. The As-induced stress response may also play into the cell cycle effects demonstrated with As exposure. Increased nitric oxide generation with 10 μM As3+ has been observed specifically in the G2/M phase of the cell cycle (Yuan et al., 2003) suggesting a causal relationship.
Our investigation demonstrates a direct effect of As3+ on cell cycle entry and cell cycle progression of primary midbrain neuroepithelial cells. The research presented here provides evidence that developmental As3+ exposure could be associated with subtle effects to neurodevelopment that may not manifest as overt teratogenesis. Given the well-described windows of sensitivity of the developing nervous system to environmental insult, and restricted compensatory capacity to overcome such insult, the results presented here suggest possible impacts on cell cycle progression which in turn may lead to deleterious induction of cytostasis and/or apoptosis. In other studies, we are using p53 wildtype and null fibroblasts to investigate specific cell cycle targets and their role in As3+ mediated growth arrest and toxicity.
ACKNOWLEDGMENTS
This work was supported in part by the USEPA-NIEHS UW Center for Child Environmental Health Risks Research (EPA R826886-01 and NIEHS PO1ES09601), the NIEHS grant R01-ES10613, UW NIEHS Center for Ecogenetics and Environmental Health (P30 ES07033), NIEHS Toxicogenomic Research Consortium NIEHS (U10 ES 11387), and the Institute for Evaluating Health Risks.
REFERENCES
Bates, S., and Vousden, K. H. (1996). p53 in signaling checkpoint arrest or apoptosis. Curr. Opin. Genet. Dev. 6, 12–18.
Beaudoin, A. R. (1974). Teratogenicity of sodium arsenate in rats. Teratology 10, 153–157.
Borenfreund, E., and Puerner, J. (1985). Toxicity determined in vitro by morphological alterations and neurtral red absorption. Toxicol. Lett. 24, 119–124.
Chattopadhyay, S., Bhaumik, S., Nag Chaudhury, A., and Das Gupta, S. (2002a). Arsenic induced changes in growth development and apoptosis in neonatal and adult brain cells in vivo and in tissue culture. Toxicol. Lett. 128, 73–84.
Chattopadhyay, S., Bhaumik, S., Purkayastha, M., Basu, S., Nag Chaudhuri, A., and Das Gupta, S. (2002b). Apoptosis and necrosis in developing brain cells due to arsenic toxicity and protection with antioxidants. Toxicol. Lett. 136, 65–76.
Chou, R., and Huang, H. (2002). Restoration of p53 tumor suppressor pathway in human cervical carcinoma cells by sodium arsenite. Biochem. Biohphys. Res. Comm. 293, 298–306.
Copp, A., Brook, F., Estibeiro, J., Shum, A., and Cockroft, D. (1990). The embryonic development of mammalian neural tube defects. Prog. Neurobiol. 35, 363–403.
Cosenza, M. E., and Bidanset, J. (1995). Effects of chlorpyrifos on neuronal development in rat embryo midbrain micromass cultures. Vet. Hum. Toxicol. 37, 118–21.
Dare, E., Gorman, A., Ahlbom, E., Gotz, M., Momoi, T., and Ceccatelli, S. (2001). Apoptotic morphology does not always require caspase activity in rat cerebellar granule neurons. Neurotox. Res. 3, 501–514.
DeSesso, J. M., Jacobson, C. F., Scialli, A. R., Farr, C. H., and Holson, J. F. (1998). An assessment of the developmental toxicity of inorganic arsenic. Reprod. Toxicol. 12, 385–433.
EPA (2001). www.epa.gov/safewater/ars/ars_rule_factsheet.html.
Faustman, E., Ponce, R., Ou, Y., Mendoza, A., Lewandowski, T., and Kavanagh, T. (2002). Investigations of methylmercury-induced alterations in neurogenesis. Environ. Health Perspect. 110, 859–864.
Filippova, M., and Duerksen-Hughes, P. J. (2003). Inorganic and dimethylated arsenic species induce cellular p53. Chem. Res. Toxicol. 16, 423–431.
Flint, O. P. (1983). A micromass culture method for rat embryonic neural cells. J. Cell Sci. 61, 247–262.
Flint, O. P., and Orton, T. C. (1984). An in vitro assay for teratogens with cultures of rat embryo midbrain and limb bud cells. Toxicol. Appl. Pharmacol. 76, 383–395.
Gebel, T. (2000). Confounding variables in the environmental toxicology of arsenic. Toxicology 144, 155–162.
George, H., and Gabay, S. (1968). Brain aromatic aminotransferase. I. Purification and some properties of pig brain L-phenylalanine-2-oxoglutarate aminotransferase. Biochim. Biophys. Acta 167, 555–566.
Gribble, E. J., Hong, S. W., and Faustman, E. M. (2005). The magnitude of methylmercury-induced cytotoxicity and cell cycle arrest is p53-dependent. Birth Defects Res. A Clin. Mol. Teratol. 73, 29–38.
Gribble, E. J., Mendoza, A., Hong, S., Sidhu, J., and Faustman, E. M. (2003). Evaluation of cell cycle kinetics in p53 mouse embryonal fibroblasts: Effects of meythyl mercury. Toxicol. Sci. 72, 67–67.
Huang, H., Chang, W., and Chen, C. (2002). Involvement of reactive oxygen species in arsenite-induced downregulation of phospholipid hydroperoxide glutathione peroxidase in human epidermoid carcinoma A431 cells. Free-Radical Biol. Med. 33, 864–873.
Jin, Y., Sun, G., Li, X., Li, G., Lu, C., and Qu, L. (2004). Study on the toxic effects induced by different arsenicals in primary cultured rat astroglia. Toxicol. Appl. Pharmacol. 196, 396–403.
Komissarova, E. V., Saha, S. K., and Rossman, T. G. (2005). Dead or dying: The importance of time in cytotoxicity assays using arsenite as an example. Toxicol. Appl. Pharmacol. 202, 99–107.
Li, X., Ding, X., and Adrian, T. E. (2003). Arsenic trioxide induces apoptosis in pancreatic cancer cells via changes in cell cycle, caspase activation, and GADD expression. Pancreas 27, 174–179.
Lindgren, A., Danielsson, B., Dencker, L., and Vahter, M. (1984). Embryotoxicity of arsenite and arsenate: distribution in pregnant mice and monkeys and effects on embryonic cells in vitro. Acta Pharmacol. Toxicol. 54, 311–320.
Ma, D., Sun, Y., Chang, K., Ma, X., Huang, S., Bai, Y., Kang, J., Liu, Y., and Chu, J. (1998). Selective induction of apoptosis of NB4 cells from G2/M phase by sodium arsenite at lower doses. Eur. J. Haematol. 61, 27–35.
McCabe, M. J., Jr., Singh, K. P., Reddy, S. A., Chelladurai, B., Pounds, J. G., Reiners, J. J., Jr., and States, J. C. (2000). Sensitivity of myelomonocytic leukemia cells to arsenite-induced cell cycle disruption, apoptosis, and enhanced differentiation is dependent on the inter-relationship between arsenic concentration, duration of treatment, and cell cycle phase. J. Pharmacol. Exp. Ther. 295, 724–733.
Mendoza, M. A., Ponce, R. A., Ou, Y. C., and Faustman, E. M. (2002). p21(WAF1/CIP1) inhibits cell cycle progression but not G2/M-phase transition following methylmercury exposure. Toxicol. Appl. Pharmacol. 178, 117–125.
Menegola, E., Broccia, M. L., Di Renzo, F., Massa, V., and Giavini, E. (2004). Effects of excess and deprivation of serotonin on in vitro neuronal differentiation. In Vitro Cell Dev. Biol. Anim. 40, 52–56.
Namgung, U., and Xia, Z. (2000). Arsenite-induced apoptosis in cortical neurons is mediated by c-Jun N-terminal protein kinase 3 and p38 mitogen-activated protein kinase. J. Neurosci. 20, 6442–6451.
Namgung, U., and Xia, Z. (2001). Arsenic induces apoptosis in rat cerebellar neurons via activation of JNK3 and p38 MAP kinases. Toxicol. Appl. Pharmacol. 174, 130–138.
Ou, Y., Thompson, S., Kirchner, S., Kavanagh, T., and Faustman, E. (1997). Induction of growth arrest and DNA damage-inducible genes Gadd45 and Gadd153 in primary rodent embryonic cells following exposure to methylmercury. Toxicol. Appl. Pharmacol. 147, 31–38.
Ou, Y., Thompson, S., Ponce, R., Schroeder, J., Kavanagh, T., and Faustman, E. (1999). Induction of the cell cycle regulatory gene p21 (Waf1, Cip1) following methylmercury exposure in vitro and in vivo. Toxicol. Appl. Pharmacol. 157, 203–212.
Park, W., Cho, Y., Jung, C., Park, J., Kim, K., Im, Y., Lee, M., Kang, W., and Park, K. (2003). Arsenic trioxide inhibits the growth of A498 renal cell carcinoma cells via cell cycle arrest or apoptosis. Biochem. Biohphys. Res. Comm. 300, 230–235.
Park, W. H., Seol, J. G., Kim, E. S., Hyun, J. M., Jung, C. W., Lee, C. C., Kim, B. K., and Lee, Y. Y. (2000). Arsenic trioxide-mediated growth inhibition in MC/CAR myeloma cells via cell cycle arrest in association with induction of cyclin-dependent kinase inhibitor, p21, and apoptosis. Cancer Res. 60, 3065–3071.
Ponce, R., Kavanagh, T., Mottet, N., Whittaker, S., and Faustman, E. (1994). Effects of methyl mercury on the cell cycle of primary rat CNS cells in vitro. Toxicol. Appl. Pharmacol. 127, 83–90.
Ribeiro, P. L., and Faustman, E. M. (1990). Chemically induced growth inhibition and cell cycle perturbations in cultures of differentiating rodent embryonic cells. Toxicol. Appl. Pharmacol. 104, 200–211.
Rodriguez, V. M., Carrizales, L., Mendoza, M. S., Fajardo, O. R., and Giordano, M. (2002). Effects of sodium arsenite exposure on development and behavior in the rat. Neurotoxicol. Teratol. 24, 743–750.
Salazar, A. M., Ostrosky-Wegman, P., Menendez, D., Miranda, E., Garcia-Carranca, A., and Rojas, E. (1997). Induction of p53 protein expression by sodium arsenite. Mutat. Res. 381, 259–265.
Scott, N., Hatlelid, K. M., MacKenzie, N. E., and Carter, D. E. (1993). Reactions of arsenic(III) and arsenic(V) species with glutathione. Chem. Res. Toxicol. 6, 102–106.
Seeley, M. R., and Faustman, E. M. (1995). Toxicity of four alkylating agents on in vitro rat embryo differentiation and development. Fundam. Appl. Toxicol. 26, 136–142.
Sidhu, J., and Omiecinski, C. (1998). Protein synthesis inhibitors exhibit a nonspecific effect on phenobarbital-inducible cytochrome P450 gene expression in primary rat hepatocytes. J. Biol. Chem. 273, 4769–4775.
Vogt, B. L., and Rossman, T. G. (2001). Effects of arsenite on p53, p21 and cyclin D expression in normal human fibroblasts – a possible mechanism for arsenite's comutagenicity. Mutat. Res. 478, 159–168.
Waalkes, M. P., Liu, J., Ward, J. M., and Diwan, B. A. (2004). Animal models for arsenic carcinogenesis: Inorganic arsenic is a transplacental carcinogen in mice. Toxicol. Appl. Pharmacol. 198, 377–384.
Wang, T. S., Kuo, C. F., Jan, K. Y., and Huang, H. (1996). Arsenite induces apoptosis in Chinese hamster ovary cells by generation of reactive oxygen species. J. Cell. Physiol. 169, 256–268.
Whittaker, S. G., and Faustman, E. M. (1991). Effects of albendazole and albendazole sulfoxide on cultures of differentiating rodent embryonic cells. Toxicol. Appl. Pharmacol. 109, 73–84.
Whittaker, S. G., and Faustman, E. M. (1992). Effects of benzimidazole analogs on cultures of differentiating rodent embryonic cells. Toxicol. Appl. Pharmacol. 113, 144–151.
Whittaker, S. G., Wroble, J. T., Silbernagel, S. M., and Faustman, E. M. (1993). Characterization of cytoskeletal and neuronal markers in micromass cultures of rat embryonic midbrain cells. Cell Biol. Toxicol. 9, 359–375.
Willhite, C. C., and Ferm, V. H. (1984). Prenatal and developmental toxicology of arsenicals. Adv. Exp. Med. Biol. 177, 205–228.
Wlodarczyk, B. J., Bennett, G. D., Calvin, J. A., and Finnell, R. H. (1996). Arsenic-induced neural tube defects in mice: Alterations in cell cycle gene expression. Reprod. Toxicol. 10, 447–454.
Yeh, J. Y., Cheng, L. C., Liang, Y. C., and Ou, B. R. (2003). Modulation of the arsenic effects on cytotoxicity, viability, and cell cycle in porcine endothelial cells by selenium. Endothelium 10, 127–139.
Yih, L. H., and Lee, T. C. (2000). Arsenite induces p53 accumulation through an ATM-dependent pathway in human fibroblasts. Cancer Res. 60, 6346–6352.
Yuan, S. S., Hou, M. F., Chang, H. L., Chan, T. F., Wu, Y. H., Wu, Y. C., and Su, J. H. (2003). Arsenite-induced nitric oxide generation is cell cycle-dependent and aberrant in NBS cells. Toxicol. In Vitro 17, 139–143.
Zhao, S., Tsuchida, T., Kawakami, K., Shi, C., and Kawamoto, K. (2002). Effect of As2O3 on cell cycle progression and cyclins D1 and B1 expression in two glioblastoma cell lines differing in p53 status. Int. J. Oncol. 21, 49–55.(Jaspreet S. Sidhu, Rafael A. Ponce, Meli)
Center for Ecogenetics and Environmental Health and Institute for Risk Analysis and Risk Communication, Seattle, Washington 98105
Center on Human Development and Disability, Seattle, Washington 98195
ABSTRACT
Arsenite (As3+) exposure during development has been associated with neural tube defects and other structural malformations, and with behavioral alterations including altered locomotor activity and operant learning. The molecular mechanisms underlying these effects are uncertain. Because arsenic can cross the placenta and accumulate in the developing neuroepithelium, we examined cell cycling effects of sodium arsenite (As3+ 0, 0.5, 1, 2, and 4 μM) on embryonic primary rat midbrain (gestational day [GD] 12) neuroepithelial cells over 48 h. There was a concentration- and time-dependent As3+-induced reduction in cell viability assessed by neutral red dye uptake assay but minimal apoptosis at concentrations below 4 μM. Morphologically, apoptosis was not apparent until 4 μM at 24 h, which was demonstrated by a marginal but statistically significant increase in cleaved caspase-3/7 activity. Cell cycling effects over several rounds of replication were determined by continuous 5-bromo-2'-deoxyuridine (BrdU) labeling and bivariate flow cytometric Hoechst-Propidium Iodide analysis. We observed a time- and concentration-dependent inhibition of cell cycle progression as early as 12 h after exposure (0.5 μM). In addition, data demonstrated a concentration-dependent increase in cytostasis within all cell cycle phases, a decreased proportion of cells able to reach the second cell cycle, and a reduced cell cycle entry from gap 1 phase (G1). The proportion of affected cells and the severity of the cell cycle perturbation, which ranged from a decreased transition probability to complete cytostasis in all cell cycle phases, were also found to be concentration-dependent. Together, these data support a role for perturbed cell cycle progression in As3+ mediated neurodevelopmental toxicity.
Key Words: sodium arsenite; cell cycle; embryonic neuroepithelial cells; development; neurotoxicity.
INTRODUCTION
Arsenic (As) is a naturally occurring element that exists in several oxidative states but it is the pentavalent (arsenate, As5+) and trivalent (arsenite, As3+) forms that are most prevalent in the environment and have toxicological significance. Of the two forms, arsenite is considered a more potent developmental toxicant (Willhite and Ferm, 1984). Trivalent arsenicals readily react with thiol-containing molecules interfering with various biochemical processes resulting in toxicity (Scott et al., 1993). The U.S. Food and Drug Administration has calculated the mean daily intake of inorganic arsenic (arsenite and arsenate) to be approximately 0.5 g/kg body weight for adults including sources from air, water, and food. Because of the extensive distribution and associated toxicity of As, contamination of drinking waters represents a worldwide problem. In response, the Environmental Protection Agency (EPA) has established a drinking water standard of 10 ppb or a maximum contaminant level of 0.010 mg/l (EPA, 2001).
Inorganic arsenic has been shown to freely cross the placenta and accumulate in the embryonic neuroepithelium (Lindgren et al., 1984) and has been linked to various embryotoxic outcomes, including neural tube defects, reduced organ and body weight, malformations, and fetal death (Willhite and Ferm 1984; Wlodarczyk et al., 1996). Acute exposure has been associated with overt malformation in the offspring and nearly fatal maternal toxicity, implying that environmentally relevant levels of As3+ are not toxicologically significant for developmental structural defects (DeSesso et al., 1998). However, chronic exposures of As3+ (maternal dose estimated at 2.9–4.2 mg/kg/day) demonstrated a significant impact on neurological function in developing and postnatal animals, suggesting neurodevelopmental risk with low concentrations (Rodriguez et al., 2002). In addition, studies in neonatal (P0) neuronal cells exposed to less than 5 μM of As3+, found decreased cell viability, altered cell growth, and apoptosis (Chattopadhyay et al., 2002a,b) reflecting mechanisms by which low doses of As may lead to neurodevelopmental toxicity. Developmental impact of As is also apparent in studies that have shown transplacental carcinogenicity with in utero exposure (Waalkes et al., 2004).
Both neurons (Namgung and Xia, 2001) and astroglia (Jin et al., 2004) are more sensitive to As3+ as compared to As5+, with exposure resulting in morphological changes and decreased cell viability. In addition, a significantly lower dose of As3+ is sufficient to induce apoptosis in neuronal cells as compared to non-neuronal cells illustrating the sensitivity of the nervous system to this metal (Wang et al., 1996). Mechanistically, As3+ has been shown to interfere with cell regulatory control via the induction of p53 and bcl-2 (Wlodarczyk et al., 1996). P53 is a critical regulator of the cell cycle and has been shown to undergo a dose- and time-dependent induction in response to both As3+ (Filippova and Duerksen-Hughes, 2003) and arsenic trioxide (As2O3) (Yeh et al., 2003). Induction of p53 in response to As2O3 exposure has been associated with subsequent cell cycle arrest at gap 2 phase (G2)/mitosis (M) (Zhao et al., 2002). Both As2O3 (Li et al., 2003) and As3+ (Huang et al., 2002) have been shown to induce p21, a critical cell cycle regulatory protein. As with p53, arsenic induced p21 has also been associated with blocking cell cycle progression at G2/M (Chou and Huang, 2002) as well as G1/S (synthesis phase) (Park et al., 2000). Other thiol reactive metals, such as methylmercury (MeHg) (Ou et al., 1999), similarly induce p21 (Faustman et al., 2002) and lead to G2/M arrest (Gribble et al., 2003), suggesting a common cellular response between the metals involving cell cycle perturbation.
Cell cycle arrest at G2/M induced by As3+ (Ma et al., 1998) and As203 (Li et al., 2003) has been associated with subsequent apoptosis indicating the latter may occur due to As-mediated cell cycle inhibition. Namgung and Xia (2001) have reported As induced apoptosis in studies with primary cerebellar and cortical neurons exposed to environmentally relevant doses of As3+ (Namgung and Xia, 2001). These studies highlight the impact of As on cell cycle progression and apoptosis and their potential link to As associated neurodevelopmental effects.
Here we present results of recent experiments examining the effect of low-dose concentrations of As3+ on cell cycle kinetics and dynamics in primary embryonic rat neuroepithelial cells during the critical "window of susceptibility" when rapid neurogenesis is occurring (Copp et al., 1990). Using continuous bromodeoxyuridine (BrdU) labeling in vitro, Hoechst-Propidium Iodide DNA staining, and bivariate flow cytometry, we were able to monitor individual cell cycle progression through several cell division rounds in primary rat midbrain cell cultures with continuous As3+ exposure. Our findings demonstrate a dose-responsive delay in cell cycle progression in all cell cycle phases induced by As3+ in primary embryonic neuroepithelial cells and suggest that cell cycle arrest may contribute to the neurodevelopmental toxicity observed with this metal.
MATERIALS AND METHODS
Cell culture.
Embryonic rat midbrain neuroepithelial cell cultures were prepared according to our previously established protocol (Flint, 1983; Ponce et al., 1994b; Ribeiro and Faustman, 1990; Whittaker and Faustman, 1991, 1992; Whittaker et al., 1993) with slight modifications. Gravid uteri were removed from pregnant (GD 12) Sprague-Dawley albino rats (Charles River, Hollester, CA) and placed in Earl's Balanced Salt Solution (EBSS, Gibco BRL, Grand Island, NY). Midbrain sections were dissected from individual embryos and incubated in calcium- and magnesium-free EBSS (CMF-EBSS, Gibco BRL, 37°C, 20 min). Midbrain sections were treated with trypsin (1% w/v, Gibco BRL) in CMF-EBSS (pH 7.2–7.4, 37°C, 20 min). Enzymatic digestion was stopped with addition of culture media containing 10% fetal bovine serum (Qualified, Gibco BRL), 2 mM L-glutamine (200 mM stock, Gibco BRL), 100 units/ml penicillin, 0.1 mg/ml streptomycin (Gibco BRL), and 88% Ham's F-12 nutrient mixture (Gibco BRL, stored at 4°C and kept from light). A single cell suspension of the neuroepithelial cells was prepared by mechanical dissociation of the cells with a fine bore (0.7 mm) glass Pasteur pipette followed by filtration (10 mm nylon mesh) to remove debris and clumps. The cell concentration was adjusted to 5 x 106 cells/ml with addition of culture media. Five 10-μl aliquots of cell suspension were plated per 35-mm Primaria-coated culture dish (Falcon). The cultures were incubated (5% CO2/95% air, 37°C, 100% humidity) for 2 h to allow cell attachment. After the attachment period, 2 ml of culture media containing 5-bromo-2'-deoxyuridine (80 μM, BrdU, Sigma) and sodium arsenite (0–4 μM) was added. Under these conditions, these cultures have been demonstrated through immunocytochemistry staining to consist of a nearly pure population of proliferating, immature neuronal precursor cells that differentiate into mature neuron over time (Whittaker et al., 1993). Sodium arsenite (As3+ NaAs02, Fischer Chemical Corporation) was prepared as a working stock (100 mM) solution in sterile water and stored at –20°C. Treatment was conducted in a Class II Type B2 hood (The Baker Company Inc., Sanford, ME).
Cell harvest.
Cell harvest was conducted at serial time points post-treatment. Culture media containing BrdU and arsenic was removed, cells were rinsed with CMF-PBS (37°C), and then incubated with 500 μl CMF-PBS containing 0.05% w/v trypsin and 0.02% EDTA (Fischer Scientific, 5 min, 37°C). Trypsinization was stopped by the addition of 500 μl culture media and the resulting cell suspension was transferred to 2 ml polyethylene tubes (Falcon) for centrifugation (1500 x g, 15 min, RT). After centrifugation, the supernatant was removed and 450 μl of Hoechst buffer [1.2 μg/ml Hoechst 33258 (Molecular Probes, Eugene OR), 0.146 M NaCl, 0.1 M TRIS Base, 0.5 mM MgCl2, 0.2% BSA, 0.1% NP-40 (Sigma, St. Louis, MO) in dd H20 and 50 μl of 10% dimethyl sulfoxide I were added to each sample. Cells were transferred to labeled amber 1.5 ml microfuge tubes (VWR Scientific Products, West Chester, PA) and frozen at a rate of –1°C/min to –20°C until analysis. Cell treatment and harvest procedures were conducted under sodium lamp illumination to minimize BrdU-induced photosensitization.
Flow cytometry.
BrdU-Hoechst analysis was performed according to established procedures (Mendoza et al., 2002; Ponce, 1994). Cells were thawed in a water bath (37°C), and 10 μg/ml Propidium Iodide (PI, Sigma) was added to each tube. Cells were stored on ice, protected from light for a minimum of 30 min prior to analysis. Flow cytometric cell cycle analysis was performed on a Coulter Epics Elite flow cytometer using a UV laser emitting 335 nm ultraviolet light. A 450/35 nm band pass filter was used to collect the Hoechst 33258 fluorescence. Hoechst 33258 binds to AT-rich regions in DNA and its fluorescence is proportionally quenched as cycling cells incorporate BrdU. An air-cooled 15 mW argon ion laser (Cyonics, San Jose, CA) emitting 488 nm light was used with a 590 nm long pass filter to excite and collect PI fluorescence, respectively. The flow rate was maintained at less than 200 cells per second. A minimum of 20,000 events per treatment was analyzed. Samples were protected from light throughout the experiment. Cell cycle analysis was quantified using MultiPlus Software (Phoenix Flow Systems, San Diego CA). Clumps and doublets were excluded from the analysis by electronic gating. Analysis was conducted to determine the proportion of cells in each cell cycle phase (G0/G1, S, G2/M) from successive rounds of cell division.
Assessment of cell morphology.
Following chemical treatment, cellular morphology was examined at 24 and 48 h, with a Nikon inverted microscope equipped with phase-contrast optics (Nikon, Tokyo, Japan). All images were captured and digitized using a Coolsnap Camera (Roper Scientific, Inc) and processed using Photoshop software (v. 5.5, Adobe, Seattle, WA).
Cytotoxicity assay.
Cytotoxicity assessments were conducted using the neutral red (NR) dye uptake assay (Borenfreund and Puerner, 1985), which determines cell viability by assessing lysosomal accumulation of NR dye. Cytotoxicity assessments were done with continuous exposure of the primary neuroepithelial cells to As3+ over 48 h (Komissarova et al., 2005). Briefly, cell culture plates were removed from the incubator at each sampling time and the media was aspirated. Cells were washed once with CMF-PBS. Fresh media containing 50 μg/ml NR was added to each 35 mm dish. After incubation (3 h, 37°C, 5% CO2), the cells were washed with CMF-PBS and the NR was eluted with 2 ml of 0.5% acetic acid/50% ethanol solution for 1 h at room temperature. Four 200-μl aliquots of NR solution were removed from each dish and placed into replicate wells of a micro titer plate. After rapid agitation of the plate reader, light absorbance was measured at 540 nm. A correction for background absorbance was conducted by subtracting media background absorbance from each well. Using these background corrected values, the relative cytotoxicity was estimated as the ratio of average absorbance between As3+ treated cultures relative to untreated control cultures.
Preparation of cell extracts.
At the stated time points, cells were washed twice with 2.5 ml ice cold CMF-PBS followed by the addition of 50 μl of cell lysis buffer (Sidhu and Omiecinski, 1998) containing additional phosphatase and protease inhibitor cocktails (Calbiochem). Cells were harvested by scraping in cell lysis buffer and then placed on ice. After storage at –80°C, all extracts were homogenized by sonication and then centrifuged (16,000 x g, 10 min, 4°C) to remove insoluble material. The resulting supernatant (cell extract) was gently removed, and total protein was determined using the Bio-Rad kit (Pierce, IL).
Apoptosis assessment.
Apoptosis was determined in cell extracts by measuring the functional activity of caspase-3 using a caspase-specific fluorogenic substrate. The activity of cleaved-caspase 3 was measured in a micro titer plate by a fluorometric assay, using Ac-DEVD-AMC as the caspase-specific substrate. Briefly 10 μg of cell extract was added in duplicate in 96 well plate format. Reaction buffer was added and incubated at 37°C for 2 h. The enzyme-catalyzed release of 7-amino-4-methyl coumarin (AMC) was measured by a fluorescence microplate reader at excitation 360 nm and emission 460 nm. Fluorescent units were converted to pmol of AMC released per μg of protein and incubation time (2 h) using a standard curve generated with known serial dilutions of AMC. The absolute activities due to As3+ treatments relative to untreated controls were converted by expressing the former as a percentage of control.
Statistical analysis.
ANOVA was performed using the General Linear Model (GLM) General Factorial procedure. Repeated measures of ANOVA were conducted to evaluate differences between As3+-treated and control cell proliferation. Post-hoc analysis of pair-wise differences between treatment groups was conducted using Fisher's PLSD. A p-value 0.05 was considered statistically significant. All statistical analyses were performed using JMP for Windows Release 5.0 (SAS Institute, Cary, NC).
RESULTS
We examined As3+-induced cytotoxicity and cell cycle inhibition in primary cultures of gestational day (GD) 12 rat midbrain neuroepithelial cells over 12–48 h of continuous exposure with PBS or sodium arsenite (0.5–4 μM). See Figure 1. Cells cultured for flow cytometric analysis of cell cycle progression were simultaneously treated with bromodeoxyuridine (BrdU). Use of continuous BrdU labeling, Hoechst-ethidium bromide DNA staining, and bivariate flow cytometry allowed analysis of individual cell cycle progression through several rounds of cell division.
Concentration-Dependent As3+-Induced Cell Loss
A time- and concentration-dependent decrease in cell number was observed with continuous exposure of the primary neuroepithelial cells to As3+ over 48 h. For example, whereas there was no appreciable cell loss upon continuous exposure to As3+ 2 μM after 24 h of exposure, dye-uptake was reduced to approximately 60% of control cells at 48 h. In contrast, exposure to 4 μM reduced dye-uptake to 40% of control cells after 24 h and approximately 20% of control cells after 48 h of exposure to As3+. Because the NR dye-uptake assay is sensitive to both the cell loss associated with cell death and reduced cell proliferation when compared against untreated control cultures, these results reflect the aggregate effects of As3+ on cell production and viability.
As3+ Inhibited Cell Cycle Progression
Asynchronous primary rat midbrain neuroepithelial cells were incubated continuously with As3+ for up to 48 h and analyzed for cell cycle progression using BrdU-Hoechst bivariate flow cytometry. Representative BrdU-Hoechst flow cytograms identifying the distribution of cells in the first and second round of cell division at 24 h (Panel A) and 48 h (Panel B) are presented in Figure 2. Because Hoechst staining is inversely proportional to BrdU incorporation, cells in G2/M of the first cell division round will have the highest Hoechst and propidium iodide fluorescence; these cells are labeled G2/M0 in Figure 2A. Cells remaining in the first round G0/G1 phase during the BrdU labeling period are labeled as and cells that were able to complete one cell cycle and reach a new G0/G1 during the BrdU labeling period are labeled as in Figure 2.
Inspection of the cytograms from treated cells revealed a concentration- and time-dependent inhibition of cell proliferation in response to As3+ exposure (Fig. 3). Whereas 55 ± 5% of control cells successfully completed at least one round of cell division over 48 h of cell culture, only 46 ± 5% of cells treated with 0.5 μM As3+ reached the second round of cell division over this time period (Fig. 4). Among cells treated with 4 μM As3+, only 8 ± 4% reached a second round of cell division over 48 h of culture.
The cell cycle inhibition occurred in all cell cycle phases, and there was an increased probability of cytostasis with increasing As3+ concentration regardless of cell cycle phase at which As3+ exposure occurred in the asynchronous culture (Fig. 5). This conclusion is based on an evaluation of the proportion of cells that failed to take up BrdU during the culture period. Because BrdU incorporation during DNA synthesis results in a diminished Hoechst fluorescence (without affecting PI fluorescence), it is possible to discriminate the cells that have remained in their original cell cycle stage (that approximate lie on a 45° line on a bivariate Hoechst-PI scatter gram) from those that have progressed through the cell cycle in the presence of BrdU (see Fig. 2). Evaluation of the relative proportion of cytostatic cells as a function of As3+ concentration and time revealed a concentration-dependent increase in the proportion of cytostatic cells in Fig. 5. For example, whereas approximately 43 ± 4% of control cells were in their original cell cycle position (relative to the start of dosing), approximately 60 ± 2% of cells exposed to 1 μM As3+ and 91 ± 2% of cells exposed to 4 μM As3+ remained in their original cell cycle position after 48 h of culture. Evaluation of the untreated control (0 μM As3+) cells demonstrate approximately 43% of cells were in the G0-phase of the cell cycle at 48 h of culture (Fig. 6), consistent with previous observations regarding the proportion of non-cycling, presumably differentiating cells, in this cell culture system (Ponce et al., 1994). The concentration-dependent increase in the proportion of noncycling cells exposed to 0.5–2 μM As3+ can be explained by a predominant increase in the number of cells that do not enter the cell cycle and remain in G0/G1 (see Fig. 6). In contrast, not all of the cytostasis observed upon exposure to 4 μM As3+ is attributable to G0/G1 inhibition, but also includes inhibition in S- and G2/M (see Fig. 6). Specifically, whereas 91 ± 2% of cells exposed to 4 μM As3+ remained in their original cell cycle position after 48 h of culture (Fig. 5), only 76 ± 6% of these cells were in (Fig. 6).
A complex relationship between As3+ exposure, relative to concentration and duration, was observed with respect to the proportion of cytostatic cells relative to the total population of cells. For example, while it is possible to leave the original G0/G1 population (identified as Fig. 2) during 48-h of culture, thereby reducing the relative abundance of this cell type in the culture, it is not possible to repopulate this cell population. This ability to discriminate "original" G0-phase cells from "new" G0/G1-phase cells ( Fig. 2) arises from the incorporation of BrdU during the DNA synthesis, which results in a distinct population with reduced Hoechst fluorescence. As a result, any apparent increase in the relative proportion of cells in phase cells must be explained by a differential loss in the cycling cell population relative to the population. We observed an apparent increase between 24 and 48 h in the relative abundance of cytostatic cells upon exposure to 1 μM As3+ as reflected in the increased proportion of cytostatic cells over time (Fig. 5), suggesting a greater proportional loss in the cycling population relative to arrested cells. Evaluation of the cell cycle phase distribution demonstrated that the majority of cytostatic cells were in (see Fig. 6).
Apoptosis
To determine whether the observed loss in cell viability and cell number was associated with the induction of apoptosis, we measured cleaved caspase 3/7 activity using a fluorogenic and caspase-specific substrate assay (Fig. 7). The results demonstrate that a marginal concentration-dependent induction of apoptosis was indeed observed after 12 and 24 h post-treatment with As3+. However, a statistically significant induction of cleaved caspase-3/7 activity was only observed at the highest As3+ concentration examined (4 μM) at both time points with a marginal but statistically insignificant induction at 2 μM. After 48 h no statistically significant evidence of apoptosis was observed at As3+ 2 μM; this is presumed to result from the detachment/loss of cells that had undergone apoptosis at earlier time points. These results are consistent with the morphological evidence (Fig. 8) supporting significant cell loss and overt toxicity. Together these findings demonstrate that As3+-induced growth arrest occurs independent of apoptosis and may be more related to a generalized cytostasis.
DISCUSSION
The toxicity of inorganic arsenic (As) is well established with known human impacts including skin lesions, peripheral vascular disorder, peripheral neuropathy, liver injury, and cancer (Gebel, 2000). Arsenic has also been implicated as a possible teratogen (Beaudoin, 1974). The degree to which gestational As exposure affects neurodevelopment, however, remains controversial and poorly understood (DeSesso et al., 1998). Furthermore, the impact of environmentally relevant doses of As remains uncertain and requires further investigation to elucidate the mechanisms involved and their subsequent role in developmental toxicity.
Mechanistically, As-mediated effects on neurodevelopment have been linked to cell cycle perturbation through the up-regulation of key checkpoint proteins. A significant up-regulation of p53 and bcl-2 was demonstrated in neural tubes on gestational day (GD) 9.0 following maternal exposure to arsenate at 40 mg/kg (Wlodarczyk et al., 1996). The tumor suppressor protein, p53, is a key regulator of the cell cycle and is responsible for initiating cell cycle arrest in response to DNA damage (Bates and Vousden, 1996). Arsenite (As3+) has been shown to induce p53 in a dose- and time-dependent manner in several cell types (Filippova and Duerksen-Hughes, 2003; Vogt and Rossman, 2001), with induction levels and toxicity dependent on the cell system (Salazar et al., 1997). Previous studies have demonstrated that both As2O3 and As3+-mediated p53 induction is followed by cell cycle arrest at G2/M (Yih and Lee, 2000; Zhao et al., 2002). Cell cycle inhibition by As has also been associated with a subsequent apoptotic response (Chou and Huang, 2002; Park et al., 2003). Between GD 8 and GD 10 in the rat, the brain and spinal cord are developing via rapid proliferation and subsequent differentiation of neuroepithelial cells (Copp et al., 1990). Arsenic-induced cell cycle inhibition and potential apoptosis at this stage of development, therefore, could have profound impact on the development of these systems.
The present study examined the impact of concentrations (0.5–4 μM) of As3+ on cell cycle kinetics of primary embryonic rat midbrain (GD 12) neuroepithelial cells. The embryonic midbrain neuroepithelial cell culture system has been used extensively by our group and others to examine the effects of toxicants on proliferation and differentiation of neuronal precursor cells on GD 12 (Flint, 1983; Ponce et al., 1994; Ribeiro and Faustman, 1990; Whittaker and Faustman, 1991, 1992; Whittaker et al., 1993). This culture system has been used to evaluate the effects of chemicals on cell viability, [3H] -aminobutyric acid uptake (George and Gabay, 1968), and hematoxylin-stained neurites (Flint and Orton, 1984). Whittaker et al. characterized neuronal cells in these cultures that specifically stained for neuronal markers A2B5 (GQ ganglioside), -aminobutyric acid (GABA), microtubule-associated protein (MAP2, MAP5), neuron-specific enolase (NSE), neural cell adhesion molecule (N-CAM), and tau after 5 days in culture (Whittaker et al., 1993). The cells were non-immunoreactive for glial fibrillary acidic protein (GFAP) (Whittaker et al., 1993). This system has also been successfully utilized to characterize changes in cell cycling in response to exogenous factors such as alkylating agents, albendazole, methylmercury, chlorpyrifos, and serotonin (Cosenza and Bidanset, 1995; Menegola et al., 2004; Ponce et al., 1994; Ribeiro and Faustman, 1990; Seeley and Faustman, 1995; Whittaker and Faustman, 1991).
We observed a concentration- and time-dependent As3+-induced cell cycle inhibition. This response was seen as early as 12 h after exposure and was manifested as decreased entry into the cell cycle from the G0-phase (Fig. 4), an increased proportion of cytostatic cells (Fig. 5), and decreased proportion of cells reaching the second cell cycle (Fig. 4). At 24 h there was a delay in progression from S-phase to G2 and at 36 h cell cycle entry from G0/G1 was reduced. Analysis of cell cycle-phase distributions suggests a possible accumulation in G2/M (see Fig. 6). Results from the present study indicate that the As3+-associated reduction in the number of cells reaching the second cell cycle after 48 h, relative to the control, is associated with both cell cycle inhibition and resultant cytostasis. Whether these cells subsequently underwent apoptosis is still inconclusive since only a marginal increase in leaved caspase-3 activity was observed. Morphological evaluations, however, demonstrated cellular stress occurring at concentrations 4.0 μM consistent with cells exhibiting characteristics of both apoptosis and necrosis. An apoptotic response does not mandate the correlation between morphology and caspase-3 activity as previously demonstrated in cerebellar granule cells (Dare et al., 2001). Although exhibiting apoptotic morphology, the cells failed to show an increase in cleaved caspase activity upon treatment with either MeHg or H2O2. An alternate study found that intermediate concentrations of As3+ (5–10 μM) induced cytostasis while concentrations >10 μM lead to caspase-3-dependent apoptotic cell death irrespective of cell cycle phase (McCabe et al., 2000). Our results corroborate that cell cycle arrest is a more sensitive endpoint than apoptosis to As3+ exposure as we saw changes in cell cycle kinetics following treatment with 1.0 μM (12 h–48 h) in the absence of caspase activation or apoptotic morphology.
Our previous studies with another neurodevelopmental toxicant, methylmercury (MeHg), support a role for metal induced cell cycle perturbations as demonstrated by the changes to key cell cycle proteins such as p21 (Ou et al., 1999) and/or GADD genes (Ou et al., 1997), and accumulation of cells in G2/M phase that was dependent on p53 and p21 genotype (Gribble et al., 2005; Mendoza et al., 2002). Similar associations have been made with As2O3 exposure where the up-regulation of GADD45, GADD153 (Li et al., 2003), and p21 (Park et al., 2000) was associated with arrest and subsequent apoptosis. The evident involvement of cell cycle proteins speaks to the critical role that cell cycle regulation plays in the overall mechanism of As-mediated neurodevelopmental toxicity.
As demonstrated by the present study, low-dose As3+ exposure within primary embryonic rat neuroepithelial cells resulted in a decrease in cell viability. Specifically, there was a concentration- and time-dependent decrease in cell viability upon exposure to As3+ concentrations >2 μM for 24 and 48 h. Studies in cortical neurons (isolated P0, cultured 6 days prior to treatment) (Namgung and Xia, 2000) and in cerebellar neurons (isolated P6-8, cultured 7 days) (Namgung and Xia, 2001), observed a similar reduced cell viability in response to As3+ but at somewhat higher concentrations (5–10 μM). In addition, these studies demonstrated a dose-dependent, stress-induced apoptotic response involving the activation of the stress proteins, JNK and P38 MAPK, suggesting a role for the stress response in neuronal apoptosis. The As-induced stress response may also play into the cell cycle effects demonstrated with As exposure. Increased nitric oxide generation with 10 μM As3+ has been observed specifically in the G2/M phase of the cell cycle (Yuan et al., 2003) suggesting a causal relationship.
Our investigation demonstrates a direct effect of As3+ on cell cycle entry and cell cycle progression of primary midbrain neuroepithelial cells. The research presented here provides evidence that developmental As3+ exposure could be associated with subtle effects to neurodevelopment that may not manifest as overt teratogenesis. Given the well-described windows of sensitivity of the developing nervous system to environmental insult, and restricted compensatory capacity to overcome such insult, the results presented here suggest possible impacts on cell cycle progression which in turn may lead to deleterious induction of cytostasis and/or apoptosis. In other studies, we are using p53 wildtype and null fibroblasts to investigate specific cell cycle targets and their role in As3+ mediated growth arrest and toxicity.
ACKNOWLEDGMENTS
This work was supported in part by the USEPA-NIEHS UW Center for Child Environmental Health Risks Research (EPA R826886-01 and NIEHS PO1ES09601), the NIEHS grant R01-ES10613, UW NIEHS Center for Ecogenetics and Environmental Health (P30 ES07033), NIEHS Toxicogenomic Research Consortium NIEHS (U10 ES 11387), and the Institute for Evaluating Health Risks.
REFERENCES
Bates, S., and Vousden, K. H. (1996). p53 in signaling checkpoint arrest or apoptosis. Curr. Opin. Genet. Dev. 6, 12–18.
Beaudoin, A. R. (1974). Teratogenicity of sodium arsenate in rats. Teratology 10, 153–157.
Borenfreund, E., and Puerner, J. (1985). Toxicity determined in vitro by morphological alterations and neurtral red absorption. Toxicol. Lett. 24, 119–124.
Chattopadhyay, S., Bhaumik, S., Nag Chaudhury, A., and Das Gupta, S. (2002a). Arsenic induced changes in growth development and apoptosis in neonatal and adult brain cells in vivo and in tissue culture. Toxicol. Lett. 128, 73–84.
Chattopadhyay, S., Bhaumik, S., Purkayastha, M., Basu, S., Nag Chaudhuri, A., and Das Gupta, S. (2002b). Apoptosis and necrosis in developing brain cells due to arsenic toxicity and protection with antioxidants. Toxicol. Lett. 136, 65–76.
Chou, R., and Huang, H. (2002). Restoration of p53 tumor suppressor pathway in human cervical carcinoma cells by sodium arsenite. Biochem. Biohphys. Res. Comm. 293, 298–306.
Copp, A., Brook, F., Estibeiro, J., Shum, A., and Cockroft, D. (1990). The embryonic development of mammalian neural tube defects. Prog. Neurobiol. 35, 363–403.
Cosenza, M. E., and Bidanset, J. (1995). Effects of chlorpyrifos on neuronal development in rat embryo midbrain micromass cultures. Vet. Hum. Toxicol. 37, 118–21.
Dare, E., Gorman, A., Ahlbom, E., Gotz, M., Momoi, T., and Ceccatelli, S. (2001). Apoptotic morphology does not always require caspase activity in rat cerebellar granule neurons. Neurotox. Res. 3, 501–514.
DeSesso, J. M., Jacobson, C. F., Scialli, A. R., Farr, C. H., and Holson, J. F. (1998). An assessment of the developmental toxicity of inorganic arsenic. Reprod. Toxicol. 12, 385–433.
EPA (2001). www.epa.gov/safewater/ars/ars_rule_factsheet.html.
Faustman, E., Ponce, R., Ou, Y., Mendoza, A., Lewandowski, T., and Kavanagh, T. (2002). Investigations of methylmercury-induced alterations in neurogenesis. Environ. Health Perspect. 110, 859–864.
Filippova, M., and Duerksen-Hughes, P. J. (2003). Inorganic and dimethylated arsenic species induce cellular p53. Chem. Res. Toxicol. 16, 423–431.
Flint, O. P. (1983). A micromass culture method for rat embryonic neural cells. J. Cell Sci. 61, 247–262.
Flint, O. P., and Orton, T. C. (1984). An in vitro assay for teratogens with cultures of rat embryo midbrain and limb bud cells. Toxicol. Appl. Pharmacol. 76, 383–395.
Gebel, T. (2000). Confounding variables in the environmental toxicology of arsenic. Toxicology 144, 155–162.
George, H., and Gabay, S. (1968). Brain aromatic aminotransferase. I. Purification and some properties of pig brain L-phenylalanine-2-oxoglutarate aminotransferase. Biochim. Biophys. Acta 167, 555–566.
Gribble, E. J., Hong, S. W., and Faustman, E. M. (2005). The magnitude of methylmercury-induced cytotoxicity and cell cycle arrest is p53-dependent. Birth Defects Res. A Clin. Mol. Teratol. 73, 29–38.
Gribble, E. J., Mendoza, A., Hong, S., Sidhu, J., and Faustman, E. M. (2003). Evaluation of cell cycle kinetics in p53 mouse embryonal fibroblasts: Effects of meythyl mercury. Toxicol. Sci. 72, 67–67.
Huang, H., Chang, W., and Chen, C. (2002). Involvement of reactive oxygen species in arsenite-induced downregulation of phospholipid hydroperoxide glutathione peroxidase in human epidermoid carcinoma A431 cells. Free-Radical Biol. Med. 33, 864–873.
Jin, Y., Sun, G., Li, X., Li, G., Lu, C., and Qu, L. (2004). Study on the toxic effects induced by different arsenicals in primary cultured rat astroglia. Toxicol. Appl. Pharmacol. 196, 396–403.
Komissarova, E. V., Saha, S. K., and Rossman, T. G. (2005). Dead or dying: The importance of time in cytotoxicity assays using arsenite as an example. Toxicol. Appl. Pharmacol. 202, 99–107.
Li, X., Ding, X., and Adrian, T. E. (2003). Arsenic trioxide induces apoptosis in pancreatic cancer cells via changes in cell cycle, caspase activation, and GADD expression. Pancreas 27, 174–179.
Lindgren, A., Danielsson, B., Dencker, L., and Vahter, M. (1984). Embryotoxicity of arsenite and arsenate: distribution in pregnant mice and monkeys and effects on embryonic cells in vitro. Acta Pharmacol. Toxicol. 54, 311–320.
Ma, D., Sun, Y., Chang, K., Ma, X., Huang, S., Bai, Y., Kang, J., Liu, Y., and Chu, J. (1998). Selective induction of apoptosis of NB4 cells from G2/M phase by sodium arsenite at lower doses. Eur. J. Haematol. 61, 27–35.
McCabe, M. J., Jr., Singh, K. P., Reddy, S. A., Chelladurai, B., Pounds, J. G., Reiners, J. J., Jr., and States, J. C. (2000). Sensitivity of myelomonocytic leukemia cells to arsenite-induced cell cycle disruption, apoptosis, and enhanced differentiation is dependent on the inter-relationship between arsenic concentration, duration of treatment, and cell cycle phase. J. Pharmacol. Exp. Ther. 295, 724–733.
Mendoza, M. A., Ponce, R. A., Ou, Y. C., and Faustman, E. M. (2002). p21(WAF1/CIP1) inhibits cell cycle progression but not G2/M-phase transition following methylmercury exposure. Toxicol. Appl. Pharmacol. 178, 117–125.
Menegola, E., Broccia, M. L., Di Renzo, F., Massa, V., and Giavini, E. (2004). Effects of excess and deprivation of serotonin on in vitro neuronal differentiation. In Vitro Cell Dev. Biol. Anim. 40, 52–56.
Namgung, U., and Xia, Z. (2000). Arsenite-induced apoptosis in cortical neurons is mediated by c-Jun N-terminal protein kinase 3 and p38 mitogen-activated protein kinase. J. Neurosci. 20, 6442–6451.
Namgung, U., and Xia, Z. (2001). Arsenic induces apoptosis in rat cerebellar neurons via activation of JNK3 and p38 MAP kinases. Toxicol. Appl. Pharmacol. 174, 130–138.
Ou, Y., Thompson, S., Kirchner, S., Kavanagh, T., and Faustman, E. (1997). Induction of growth arrest and DNA damage-inducible genes Gadd45 and Gadd153 in primary rodent embryonic cells following exposure to methylmercury. Toxicol. Appl. Pharmacol. 147, 31–38.
Ou, Y., Thompson, S., Ponce, R., Schroeder, J., Kavanagh, T., and Faustman, E. (1999). Induction of the cell cycle regulatory gene p21 (Waf1, Cip1) following methylmercury exposure in vitro and in vivo. Toxicol. Appl. Pharmacol. 157, 203–212.
Park, W., Cho, Y., Jung, C., Park, J., Kim, K., Im, Y., Lee, M., Kang, W., and Park, K. (2003). Arsenic trioxide inhibits the growth of A498 renal cell carcinoma cells via cell cycle arrest or apoptosis. Biochem. Biohphys. Res. Comm. 300, 230–235.
Park, W. H., Seol, J. G., Kim, E. S., Hyun, J. M., Jung, C. W., Lee, C. C., Kim, B. K., and Lee, Y. Y. (2000). Arsenic trioxide-mediated growth inhibition in MC/CAR myeloma cells via cell cycle arrest in association with induction of cyclin-dependent kinase inhibitor, p21, and apoptosis. Cancer Res. 60, 3065–3071.
Ponce, R., Kavanagh, T., Mottet, N., Whittaker, S., and Faustman, E. (1994). Effects of methyl mercury on the cell cycle of primary rat CNS cells in vitro. Toxicol. Appl. Pharmacol. 127, 83–90.
Ribeiro, P. L., and Faustman, E. M. (1990). Chemically induced growth inhibition and cell cycle perturbations in cultures of differentiating rodent embryonic cells. Toxicol. Appl. Pharmacol. 104, 200–211.
Rodriguez, V. M., Carrizales, L., Mendoza, M. S., Fajardo, O. R., and Giordano, M. (2002). Effects of sodium arsenite exposure on development and behavior in the rat. Neurotoxicol. Teratol. 24, 743–750.
Salazar, A. M., Ostrosky-Wegman, P., Menendez, D., Miranda, E., Garcia-Carranca, A., and Rojas, E. (1997). Induction of p53 protein expression by sodium arsenite. Mutat. Res. 381, 259–265.
Scott, N., Hatlelid, K. M., MacKenzie, N. E., and Carter, D. E. (1993). Reactions of arsenic(III) and arsenic(V) species with glutathione. Chem. Res. Toxicol. 6, 102–106.
Seeley, M. R., and Faustman, E. M. (1995). Toxicity of four alkylating agents on in vitro rat embryo differentiation and development. Fundam. Appl. Toxicol. 26, 136–142.
Sidhu, J., and Omiecinski, C. (1998). Protein synthesis inhibitors exhibit a nonspecific effect on phenobarbital-inducible cytochrome P450 gene expression in primary rat hepatocytes. J. Biol. Chem. 273, 4769–4775.
Vogt, B. L., and Rossman, T. G. (2001). Effects of arsenite on p53, p21 and cyclin D expression in normal human fibroblasts – a possible mechanism for arsenite's comutagenicity. Mutat. Res. 478, 159–168.
Waalkes, M. P., Liu, J., Ward, J. M., and Diwan, B. A. (2004). Animal models for arsenic carcinogenesis: Inorganic arsenic is a transplacental carcinogen in mice. Toxicol. Appl. Pharmacol. 198, 377–384.
Wang, T. S., Kuo, C. F., Jan, K. Y., and Huang, H. (1996). Arsenite induces apoptosis in Chinese hamster ovary cells by generation of reactive oxygen species. J. Cell. Physiol. 169, 256–268.
Whittaker, S. G., and Faustman, E. M. (1991). Effects of albendazole and albendazole sulfoxide on cultures of differentiating rodent embryonic cells. Toxicol. Appl. Pharmacol. 109, 73–84.
Whittaker, S. G., and Faustman, E. M. (1992). Effects of benzimidazole analogs on cultures of differentiating rodent embryonic cells. Toxicol. Appl. Pharmacol. 113, 144–151.
Whittaker, S. G., Wroble, J. T., Silbernagel, S. M., and Faustman, E. M. (1993). Characterization of cytoskeletal and neuronal markers in micromass cultures of rat embryonic midbrain cells. Cell Biol. Toxicol. 9, 359–375.
Willhite, C. C., and Ferm, V. H. (1984). Prenatal and developmental toxicology of arsenicals. Adv. Exp. Med. Biol. 177, 205–228.
Wlodarczyk, B. J., Bennett, G. D., Calvin, J. A., and Finnell, R. H. (1996). Arsenic-induced neural tube defects in mice: Alterations in cell cycle gene expression. Reprod. Toxicol. 10, 447–454.
Yeh, J. Y., Cheng, L. C., Liang, Y. C., and Ou, B. R. (2003). Modulation of the arsenic effects on cytotoxicity, viability, and cell cycle in porcine endothelial cells by selenium. Endothelium 10, 127–139.
Yih, L. H., and Lee, T. C. (2000). Arsenite induces p53 accumulation through an ATM-dependent pathway in human fibroblasts. Cancer Res. 60, 6346–6352.
Yuan, S. S., Hou, M. F., Chang, H. L., Chan, T. F., Wu, Y. H., Wu, Y. C., and Su, J. H. (2003). Arsenite-induced nitric oxide generation is cell cycle-dependent and aberrant in NBS cells. Toxicol. In Vitro 17, 139–143.
Zhao, S., Tsuchida, T., Kawakami, K., Shi, C., and Kawamoto, K. (2002). Effect of As2O3 on cell cycle progression and cyclins D1 and B1 expression in two glioblastoma cell lines differing in p53 status. Int. J. Oncol. 21, 49–55.(Jaspreet S. Sidhu, Rafael A. Ponce, Meli)