Disruption of cell cycle kinetics and cyclin-dependent kinase system by ethanol in cultured cerebellar granule progenitors

Developmental Brain Research 132 (2001) 47–58 www.elsevier.com / locate / bres

Research report

Disruption of cell cycle kinetics and cyclin-dependent kinase system by ethanol in cultured cerebellar granule progenitors Zheng Li, Hong Lin, Yunfeng Zhu, Mei Wang, Jia Luo* Department of Microbiology, Immunology and Cell Biology, West Virginia University School of Medicine, Robert C. Byrd Health Science Center, P.O. Box 9177, Morgantown, WV 26506 -9177, USA Accepted 9 October 2001

Abstract An in vitro model of neuronal precursors, primary culture of cerebellar granule progenitors (CGPs), was used to investigate the mechanisms underlying ethanol-induced cell cycle damage. The CGP cultures were generated from 3-day-old rats. Ethanol significantly inhibited the proliferation of the CGPs in culture. Analysis of cell cycle kinetics by a cumulative 5-bromo-29-deoxyuridine (BrdU) labeling technique demonstrated that ethanol exposure increased the duration of the cell cycle and decreased the growth fraction (the cycling population). The duration of the S-phase and total cell cycle was significantly prolonged by ethanol exposure by 220% and 135%, respectively, while the growth fraction was decreased from 44% in the control groups to 22% in the ethanol-exposed cultures. Cyclin-dependent kinase 2 (Cdk2) is a key protein that regulates both the passage from G1 into S, and the S phase progression. The results from in vitro phosphorylation assay and Western blot demonstrated that ethanol dramatically down-regulated both the activity and the expression of Cdk2. In addition, ethanol significantly decreased the expression of Cyclin A and Cyclin D 2 . Further studies using in situ TUNEL assay and DNA fragmentation ELISA showed that ethanol caused a delayed apoptosis, i.e. the ethanol-induced apoptosis was evident only after chronic exposure. On the other hand, ethanol did not affect the necrotic index. In conclusion, ethanol decreases the cycling pool of CGPs by inducing cell cycle delay and promoting apoptosis. Ethanol-mediated disturbance of the cyclin-dependent kinase system may be an important mechanism to account for cell cycle arrest in neuronal precursor cells.  2001 Elsevier Science B.V. All rights reserved. Theme: Disorders of the nervous system Topic: Neurotoxicity Keywords: Apoptosis; Cyclin-dependent kinase; CDK inhibitor; Fetal alcohol syndrome; Neuron

1. Introduction Ethanol is a potent teratogen and fetal exposure to ethanol results in considerable central nervous system dysfunction [1]. One consequence of early ethanol exposure is the depletion of selected populations of neuronal cells in brain. The developing cerebellum is especially vulnerable to ethanol exposure. Animal studies have established that the numbers of both cerebellar Purkinje cells and granule neurons are significantly reduced by ethanol exposure during development [9,11,29,47,62,63]. Ethanol-induced loss of cerebellar granule neurons may *Corresponding author. Tel.: 11-304-293-7208; fax: 11-304-2937823. E-mail address: [email protected] (J. Luo).

occur as a result of (a) disruption of proliferation of cerebellar granule progenitors (CGPs), and (b) direct killing of CGP and / or cerebellar granule neurons, or both. We hypothesize that disruption of proliferation of CGPs is a major mechanism underlying ethanol-induced neuronal depletion. In the present study, we test this hypothesis by investigating the effects of ethanol on cell cycle kinetics using an in vitro model, primary culture of rat CGPs. Furthermore, we explore the cellular and molecular mechanisms by which ethanol disrupts cell cycle kinetics. In vitro models offer several advantages for elucidation of the cellular and molecular mechanisms of ethanol action. The models provide opportunities to: (1) precisely control experimental conditions and the cellular environment (e.g. to control and maintain ethanol levels and to apply defined amounts of exogenous substances), (2)

0165-3806 / 01 / $ – see front matter  2001 Elsevier Science B.V. All rights reserved. PII: S0165-3806( 01 )00294-2

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evaluate cellular activities, such as cell cycle kinetics and activity of regulatory enzymes, and (3) assess the direct effects of ethanol on a single cell type. With this experimental system, we demonstrate here that ethanol causes a significant cell cycle delay and a severe reduction in the number of the cycling population of CGPs. The movement of cells through the cell cycle is regulated by a family of protein kinases called cyclin-dependent kinases (CDKs). Activated CDKs catalyze the phosphorylation of downstream effector targets, resulting in the onset of transcriptional events necessary for cell cycle progression. The activity of CDKs is positively regulated by cyclins and negatively regulated by CDK inhibitors (CKIs). In the present study, we further investigate the effects of ethanol on the CDK system. Our results show that expression and activity of several key proteins of the CDK system are significantly altered by ethanol exposure.

2. Materials and methods

2.1. Culture of cerebellar granule progenitors Sprague–Dawley rats were obtained from Hilltop Laboratory Inc. (Scottdale, PA). Cultures of cerebellar granule progenitors (CGPs) were created using a previously described method [21,53,55,71]. It has been demonstrated that the purity of CGPs generated using this paradigm is greater than 90.0%. Briefly, 3-day-old rat pups were decapitated and cerebella carefully removed. It has been shown that CGPs derived from this age of rats undergo maximal proliferation [53]. The brain tissue was minced with a sterile razor blade and suspended in 10 ml of trypsin solution (0.025%) at 378C. After incubation for 15 min, an equal volume of a solution containing DNAse (130 Knuitz units / ml) and trypsin inhibitor (0.75 mg / ml) was added and the tissue was sedimented by a brief (5 s) centrifugation. The tissue was dissociated by trituration, and the cell suspension was centrifuged through a 4.0% bovine serum albumin solution, which is a critical step for increasing neuronal viability. The cell pellet was resuspended in Eagle’s minimal essential medium containing the following supplements: 10.0% fetal bovine serum (FBS), 25 mM KCl, 1 mM glutamine, 33 mM glucose, and penicillin (100 units / ml) / streptomycin (100 mg / ml). Cell numbers were determined with a hemocytometer. For MTT assay and ELISA, 100 ml of cell suspension containing either 503 10 4 cells / ml or 100310 4 cells / ml were plated into poly-Dlysine (PDL)-coated wells of 96-well tissue culture trays (Costar, Cambridge, MA). The concentrations produced the seeding density of 1.6 and 3.2310 4 / cm 2 , respectively. For immunohistochemical and Western blot studies, cells were plated into removable chamber slides (Fisher Scientific, Pittsburgh, PA) or 35-mm culture dishes at a density of 3.2310 4 / cm 2 . Cells were incubated at 378C in a

humidified environment containing 5.0% CO 2 for at least 4 h to permit cell adhesion before initiating ethanol treatment.

2.2. Double-labeling immunohistochemistry Previous study has established that the procedure described above generates CGPs with purity greater than 90.0%, and that the derived CGPs are capable of proliferation in culture [53,56]. To verify that the cycling cells are CGPs, but not glia in our culture system, a double-labeling technique was employed. In this procedure, cells were double-labeled with an anti-BrdU monoclonal antibody and an anti-glial fibrillary acidic protein (GFAP). 5Bromo-29-deoxyuridine (BrdU), a thymidine analog, incorporates into DNA as a proliferating cell passes through the S-phase of the cell cycle. BrdU incorporation is an index of cell proliferation. Briefly, cultures were pulselabeled with BrdU (20 mM) for 4 h. The BrdU-labeled cells were fixed and then incubated with a mixture of an anti-BrdU monoclonal antibody and an anti-GFAP polyclonal antibody (Santa Cruz Biotechnology) diluted at 1:100 for 1 h. Cells were washed, and incubated with biotinylated goat anti-rabbit IgG (Amersham) diluted at 1:100 for 1 h. After incubation, cultures were washed, and exposed to a mixture of Texas red-coupled streptavidin (Amersham) diluted at 1:100 and fluorescein-coupled goat anti-mouse IgG (Amersham) diluted at 1:100 for 1 h. The slides were covered and viewed under an Olympus Provis fluorescent microscope.

2.3. Ethanol exposure protocol Because of ethanol’s volatility, a method utilizing sealed containers [2,42,59] was used to maintain ethanol levels in the culture medium. With this method, ethanol was added directly to the culture medium in either tissue culture trays or dishes. Then, the trays or dishes were placed in sealed containers in which there was an ethanol-containing water bath in the bottom. The concentration of ethanol in the bath was the same as that in the culture medium. Ethanol from the bath evaporates into the air of the sealed container and maintains the ethanol concentration in the culture medium. A small volume of CO 2 (60 cc) was injected into the container prior to sealing. The ethanol bath was changed daily to maintain the ethanol concentration. In control cultures, the water bath contains no ethanol. All containers were incubated at 378C. Previous studies have shown that this sealed container method accurately maintains ethanol concentrations in the culture medium [42].

2.4. MTT assay The MTT assay was employed to determine the number of viable cells in culture (Cat. [: 1 465 007, Roche

Z. Li et al. / Developmental Brain Research 132 (2001) 47 – 58

Molecular Biochemicals, Indianapolis, IN). The assay is based on the cleavage of the yellow tetrazolium salt MTT [3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl tetrazolium bromide] to purple formazan crystals by metabolically active cells [73]. Briefly, the CGPs were plated into 96-well microtiter plates at a density of 3.2310 4 / cm 2 . Cultures were exposed to ethanol (0, 50, 100, 200, and 400 mg / dl) for 4 days. After ethanol exposure, 10 ml of MTT labeling reagent were added to each well and plates were incubated at 378C for 4 h. Following MTT incubation, the cultures were solubilized and the spectrophotometric absorbance of the samples was detected by a microtiter plate reader. The wavelength to measure absorbance of formazan product is 570 nm, with a reference wavelength of 750 nm.

2.5. Analysis of cell cycle kinetics As cells proliferate, they pass through their mitotic cell cycle. The kinetics of the cell cycle, i.e. the length of the cell cycle, the S-phase and the growth fraction, were measured by a cumulative labeling method. This method is based on the fact that BrdU incorporates into DNA as a proliferating cell passes through the S-phase of the cell cycle [19,28,51]. Cells were plated into wells of removable chamber slides at a density of 3.2310 4 / cm 2 . After exposure to ethanol (400 mg / dl) for 48 h, 20 mM BrdU (Sigma) was added to the medium. At various times after BrdU addition (1, 2, 4, 12, 24, 32, 40, 48, and 72 h), the medium was removed and the cells were washed with PBS. Cells were fixed with 70.0% ethanol for 30 min at 48C. The fixed cells were immersed in 2.0 N HCl for 20 min to denature DNA. The acid was neutralized by a 30-s wash with PBS. Cells were then treated with 0.30% Triton X-100 for 20 min. Subsequently, the cells were incubated with normal goat serum for 30 min to block any nonspecific immunochemical reactions. This was followed by reaction with an anti-BrdU antibody (Sigma) at 1:100 dilution in PBS for 60 min at room temperature. After 60 min, the primary antibody was removed and the cultures were washed three times with PBS. A secondary antibody, goat anti-mouse IgG conjugated with alkaline phosphatase (1:100 dilution; Zymed Laboratories Inc., San Francisco, CA), was added and the incubation continued for another 60 min at room temperature followed again by three washes with PBS. The cultures then were treated with alkaline phophatase substrate (BCIP/ NBT substrate kit; Zymed Laboratories Inc.) for 20 min to reach full-color development. The slides were covered and examined under a phase-contrast microscope. The numbers of BrdU-positive and negative cells within an area of 0.5 mm30.5 mm were counted. Four fields were chosen at random in each slide and counted. A labeling index (LI; the number of labeled cells divided by the total number of cells counted) was calculated for each time point. The change in labeling over time (from 1 h to 72 h post-BrdU application) was fitted to formulae to determine

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the lengths of the total cell cycle (T C ), and the length of the S-phase (T S ), and the growth fraction (GF5the fraction of the cells undergoing proliferation) [43,58]. Briefly, LI at time 0 h (T 0 ) will be extrapolated from the empirical data for the LI for T 1 to T 72 . The theoretical initial labeling index (T 0 ) will be equal to the quotient of the length of the S-phase (T S ) and the total length of the cell cycle (T C ). During the period defined as T C 2 T S (i.e. the total time spent in the G2-, M-, and G1-phases), the labeling ratio will increase at a rate of 1 /T C . Thus, by determining the slope of the curve (describing the change in the labeling ratio over time, and the best fit to the empirically determined labeling indices), we were able to calculate the T C . Finally, by dividing the value for the y-intercept (T S /T C , the labeling index at T 0 ) by the slope (1 /T S ), we can calculate the T S . Control slides were prepared in which either the BrdU, primary antibody, or secondary antibody was omitted. In all cases, the staining was consistently negative in the controls.

2.6. Immunoprecipitation and Cdk2 activity Cdk2 immunoprecipitates were generated with a previously described method [45]. Briefly, the cell lysate was collected, and an aliquot containing 200 mg of protein was incubated with rabbit polyclonal antibody against Cdk2 (1:50 in PBS) for 1 h at 48C. Twenty microlitres of Protein A / G conjugated to agarose (Santa Cruz Biotech) were added to the lysate, and the mixture was incubated overnight at 48C. Immunoprecipitates were collected by centrifugation at 10,0003g for 10 min. The pellet was washed three times with 0.5 ml RIPA buffer (150 mM NaCl, 50 mM Tris (pH 8.0), 1.0% Nonidet P-40, 0.10% SDS, 0.50% deoxycholic acid sodium, 0.10 mg / ml phenylmethylsulfonyl fluoride, 1.0 mM sodium orthovanadate, and 3.0% Aprotinin (Sigma)). After the final wash, the supernatant was aspirated. The pellet was resuspended in 20 ml of assay dilution buffer (20 mM MOPs, pH 7.2, 25 mM beta-glycerol phosphate, 5 mM EGTA, 1 mM sodium orthovanadate, 1 mM dithiothreitol). Cdk2 activity was determined using a commercial kit (Upstate Biotechnology, Lake Placid, NY). The assay kit is based on the principle that Cdk2 mediates the transfer of the g-phosphate of adenosine-59-[ 32 P] triphosphate ([g- 32 P]ATP), to a specific substrate, Histone H1. The phosphorylated Histone H1 is then separated from the residual [g- 32 P]ATP using P81 phosphocellulose paper and quantified using a scintillation counter. This experiment was replicated four times.

2.7. Western blot Immunoblotting analysis of protein expression was performed with a previously described procedure [45]. Briefly, cells were washed with PBS and lysed with RIPA buffer for 10 min. Adherent cell fragments were scraped

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from the dish and the entire lysate was transferred to microfuge tubes. Solubilized cells were centrifuged, the proteinaceous supernatant was collected, and the protein concentration was determined [41]. Aliquots of the supernatant were loaded onto the lanes of a sodium dodecylsulfate (SDS) 10.0% polyacrylamide gel. Each aliquot contained 60 mg of protein. The proteins were separated by electrophoresis, and the separated proteins were transferred to nitrocellulose filters. The filters were washed with 5.0% nonfat dry milk in 0.010 M phosphate-buffered saline (pH 7.4) and 0.10% Tween-20 (TPBS) at room temperature for 1 h to block non-specific immunoreactivity. Subsequently, the filters were incubated with primary antibodies directed against Cyclin D 1 , Cyclin D 2 , Cyclin A, Cdk2, p16, p21, p19, p27 and actin (Santa Cruz Biotech) diluted at 1:200– 800 in TPBS for 1.5 h at room temperature. After two quick washes in TPBS, the filters were incubated with a secondary antibody conjugated to horseradish peroxidase (Amersham) diluted at 1:2000 in TPBS for 1 h. The immune complexes were detected with the enhanced chemiluminescence method (Amersham, Arlington Hts., IL). The blots were stripped and re-probed with an anti-actin antibody. Since ethanol does not affect the expression of actin, the variation in loading was normalized against the amount of actin expression in each lane. Each experiment was repeated 3–4 times for each primary antibody. The relative amount of each cell cycle-related protein imaged on the films was quantified using a Sigma Gel program (Jandel, San Rafael, CA).

2.8. Apoptosis assays Two distinct forms of eukaryotic cell death can be classified by morphological and biochemical criteria: apoptosis and necrosis. Apoptosis is characterized by membrane blebbing and the activation of an endogeneous endonuclease. This activated nuclease cleaves doublestranded DNA at the most accessible internucleosome linker region, generating the DNA fragments of mono- and oligonucleosomes [12,70]. Two independent assays, in situ TUNEL enzymatic labeling assay and cellular DNA fragmentation ELISA, were employed to analyze and quantify the apoptosis of cultured CGPs.

2.8.1. In situ TUNEL enzymatic labeling assay This assay offers an in situ cellular detection of apoptosis by measuring DNA strand breaks in individual cells. The assay uses an optimized terminal transferase (TdT) to label free 39OH ends in genomic DNA with fluoresceindUTP (Cat. [ 1 684 809; Roche Molecular Biochemicals). Briefly, the cells were fixed with 4.0% paraformaldehyde in PBS for 30 min at room temperature. Cell membranes were permeabilized by treating the cells with 0.10% Triton X-100 in PBS for 15 min. The permeabilized cells were washed with PBS for 5 min. After washing, the cells were

incubated with the TUNEL reaction mixture containing TdT and fluorescein-dUTP, followed by a treatment with anti-fluorescein antibody conjugated with alkaline phosphatase for 30 min. The cultures were then incubated with alkaline phosphatase substrate (BCIP/ NBT) for 20 min to reach full-color development. The slides were covered and examined under a phase-contrast microscope. The numbers of TUNEL-positive and negative cells within an area of 0.5 mm30.5 mm were counted. Four fields were chosen at random in each slide and counted. An apoptotic index (the number of labeled cells divided by the total number of cells counted) was calculated.

2.8.2. Cellular DNA fragmentation ELISA This assay is based on assessment of accumulation of DNA fragments in the cytoplasm of apoptotic cells. The enrichment of mono- and oligo-nucleosomes in the cytoplasm is due to the fact that DNA degradation occurs several hours before plasma membrane breakdown [20]. In this assay, the accumulation of cytosolic histone-bound DNA fragments was quantified using a commercial ELISA kit (Cat [: 1 544 675, Roche Molecular Biochemicals). The measurement of apoptosis by this assay is sensitive and consistent with other morphometric indices of apoptosis [78]. Briefly, the CGPs were plated into PDL-coated 96-well plates at a density of 3.2310 4 / cm 2 . Cells were exposed to ethanol (0, 200, and 400 mg / dl) for either 2 or 4 days. After ethanol treatment, the cultures were washed twice with 0.01 M PBS, and cultured cells were lysed. Twenty microlitres of cell lysate from each well were mixed with 80 ml of antibody solution in the coated wells. The loaded wells were incubated at room temperature for 2 h. Substrate was added to each well after it was washed three times in incubation buffer. After incubation at room temperature for 10–20 min, the optical density was measured using a microtiter plate reader with a light filter of 405 nm. The readings were used to measure the degree of apoptosis. 2.9. Necrosis assay Necrosis is accompanied by increased ion permeability of the plasma membrane, resulting in cell swelling and rupture of the plasma membrane. Colorimetric assay for the quantification of necrotic cell death is based on the measurement of lactate dehydrogenase (LDH) activity released from the cytosol of damaged cells into the supernatant due to the breakdown of the plasma membrane. LDH activity in the culture supernatant was measured with an ELISA kit (Cat [: 1 644 793, Roche Molecular Biochemicals). This assay is shown to be a sensitive and precise assessment of necrosis in various in vitro cell systems (e.g. Refs. [18,56]). In brief, the CGPs were plated and treated with ethanol the same way as in the apoptosis assay. After treatment, 100 ml of the cell-free supernatant were collected and mixed with 100 ml reaction

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mixture (the substrates for LDH) for 20 min in the dark at room temperature. LDH converts substrate into a redcolored formazan salt product. The intensity of color, which represents the activity of LDH was measured with a microtiter plate reader with a light filter of 490 nm. The readings were used to characterize the degree of necrosis. For a positive control, the CGPs were treated with Triton X-100 (0.05%) for 12 h and LDH release was measured. Triton X-100 treatment resulted in a 10–15-fold increase of LDH release (data not shown).

2.10. Statistical analysis Treatment groups were compared using one-way analysis of variance (ANOVA). Differences in which P,0.05 were considered statistically significant. In cases where significant differences were detected, specific post-hoc comparisons between treatment groups were examined with the Student–Newman–Keuls tests.

Fig. 2. The proliferation of CGPs in culture. The CGPs were plated at a density of 3.2310 4 / cm 2 . At 24 h after plating (day 1), cell number was determined with the MTT assay. Subsequently, the cultures were exposed to ethanol (0, 50, 100, 200 and 400 mg / dl) for 4 days (day 5) and cell number was determined. Each data point (6S.E.M.; bars) is the mean of four independent trials.

3. Results

3.1. The proliferation of CGPs in culture The cell number correlated well with the MTT absorbance at seeding densities ranging from 0.3 to 10310 4 / cm 2 , indicating that the MTT assay is an accurate method to measure the number of neuronal cells in culture (Fig. 1). The number of viable CGPs increased over time (Fig. 2).

Over 4 days in culture, the number of CGPs increased by 74%. To verify that the proliferating cells in culture were CGPs and not glia, we double-stained the cells with two different antibodies. One was an anti-BrdU monoclonal antibody and another was an anti-glial fibrillary acidic protein (GFAP, a marker for astroglia) polyclonal antibody. Our result showed that more than 95.0% of BrdU positive cells are negative for GFAP expression (data not shown), indicating that the proliferating cells are not astrocytes. At lower concentrations (50–200 mg / dl), ethanol did not have a significant effect on the number of CGPs. However, ethanol at 400 mg / dl significantly reduced the numbers of CGPs in culture (Fig. 2). Since ethanol at 400 mg / dl significantly alters the numbers of CGPs, this concentration was used throughout the subsequent studies.

3.2. Cell cycle kinetics of CGPs in culture

Fig. 1. Determination of the number of CGPs by MTT assay. The CGPs were plated at various densities ranging from 0.3 to 20310 4 / cm 2 . At 24 h after plating, the number of viable cells was determined with the MTT assay as described in the Materials and methods. Each data point (6S.E.M.; bars) is the mean of four independent trials.

The BrdU labeling index (LI, the ratio of BrdU-positive cells to total cells) at various time points post-BrdU application was determined, and the results are presented in Figs. 3 and 4. There were two phases in the change of LI over time (Fig. 4). In the first phase, LI increased steadily as more proliferating cells were labeled. The data fit a linear regression line. The LI reached a maximum when the entire cycling population (the growth fraction, GF) was labeled, and then LI stayed constant. From these data, the lengths of total cell cycle (T C ) and S-phase (T S ), and the GF were calculated [58]. The parameters of cell cycle kinetics are presented in Table 1. Ethanol exposure (400 mg / dl) significantly (P,0.05) prolonged the durations of both the total cell cycle and S-phase by 135%, and 220%, respectively. On the other hand, ethanol reduced GF by 50% (P,0.05).

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Fig. 3. Photomicrographs of BrdU immunolabeling. CGPs derived from 3-day-old rats were cultured with or without ethanol (400 mg / dl) for 48 h. Subsequently, cells were treated with BrdU (20 mM) in the presence or absence of ethanol for 24 h. The incorporation of BrdU by CGPs was determined immunohistochemically as described in the Materials and methods. (A) Control culture. (B) Ethanol-treated culture. BrdU-positive cells are indicated by arrows.

3.3. Cdk2 activity and the expression of cell cyclerelated proteins

Fig. 4. Analysis of cell cycle kinetics. After cells plated at a density of 3.2310 4 / cm 2 were exposed to ethanol (400 mg / dl) for 2 days, CGPs were treated with BrdU in the presence or absence of ethanol. The plot depicts the BrdU labeling index (LI) over time. Each data point (6S.E.M.; bars) is the mean of four independent trials.

Table 1 Parameters of the cell cycle kinetics of CGPs in culture

Control Ethanol

Cdk2 is a key kinase that regulates both the movement from G1 to S and the progression of the S phase. Therefore, we examined the effects of ethanol on the activity and expression of Cdk2. The CGPs were exposed to ethanol (400 mg / dl) for various durations (1, 2 and 4 days), and Cdk2 was isolated by immunoprecipitation. The Cdk2 activity was determined using an in vitro phosphorylation assay. As shown in Fig. 5, ethanol significantly inhibited the phosphotransferase activity of Cdk2 at every selected time point (P,0.05). The expression of Cdk2 was analyzed with Western blot. Fig. 6 documents ethanol down-regulation of expression of Cdk2. Since the activity of CDKs is positively regulated by cyclins and negatively regulated by CDK inhibitors (CDKIs), we further examined the expression of various cyclins and CDKIs. The expression of Cyclin D 1 , and three CDKIs (p16, p19, and p21) was very weak and barely detectable in cultured CGPs (data not shown). The CGPs strongly expressed Cyclin A, Cyclin D 2 and p27. Ethanol exposure significantly down-regulated the expression of Cyclin A, Cyclin D 2 and p27, whereas it did not affect actin (Fig. 6).

3.4. Survival of CGPs in culture

T C (h)

T S (h)

GF

44.6265.53 60.2765.07*

8.2061.20 18.1163.02*

0.4460.04 0.2260.04*

Cells were exposed to ethanol (0 or 400 mg / dl). The durations of S-phase (T S ) and total cell cycle (T C ) and growth fraction (GF) were calculated from the data presented in Fig. 4. *Denotes a statistically significant differences from controls. The values shown are the mean6S.E.M. of four independent trials.

Using in situ TUNEL assay, apoptotic cells were observed in control cultures, with about 7.8% of the cells being TUNEL-positive (Fig. 7). Ethanol exposure (400 mg / dl, 4 days) significantly increased (176%; P,0.05) the number of TUNEL-positive cells to 13.8%. Quantitative study using DNA fragmentation ELISA showed that a 2-day exposure to ethanol (200 or 400 mg / dl) did not alter

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the number of apoptotic cells (Fig. 8). In contrast, a 4-day ethanol exposure at 400 mg / dl resulted in a statistically significant increase (151%; P,0.05) in apoptotic cells. Thus, ethanol-induced apoptosis of CGPs was durationand concentration-dependent. Necrotic death of CGPs was quantified by assessing LDH activity in culture supernatant. As shown in Fig. 9, ethanol did not induce necrosis of CGPs.

4. Discussion

Fig. 5. The effect of ethanol on Cdk2 activity. The CGPs were exposed to ethanol at 400 mg / dl for various durations (1, 2 and 4 days) and total cellular proteins were extracted. Cdk2 was isolated from 200 mg total proteins by immunoprecipitation. The Cdk2 activity was determined using an in vitro phosphorylation assay as described in the Materials and methods. Cdk2-mediated phosphorylation was quantified using a scintillation counter and expressed as percentage of control. Each data point (6S.E.M.; bars) is the mean of four independent trials. *Denotes a statistically significant difference (P,0.05).

This is the first report showing that ethanol exposure disrupts the cell cycle kinetics in primary cultures of cerebellar granule progenitors (CGPs), establishing that cultured CGPs represent a valuable in vitro model for studying ethanol-induced cell cycle disruption to neuronal precursor cells. We demonstrate that ethanol prolongs the duration of the cell cycle and disturbs the cyclin-dependent kinase system. The expression and activity of several key proteins that regulate cell cycle progression are dramatically altered by ethanol. These results provide an important insight into the cellular and molecular mechanisms by which ethanol impairs the proliferation of neuronal precursors. Cerebellum granule neurons, the major cerebellar neuronal population, are generated in the proliferative external

Fig. 6. The expression of cell cycle-related proteins. The CGPs were exposed to ethanol at 400 mg / dl for 4 days and total proteins were isolated. Equal amounts of proteins (60 mg) were subjected to Western blot analysis. (A) The expression of cell cycle-related proteins was analyzed with Western blot. (B) The relative amount of these proteins was quantified microdensitometrically, and expressed as percentage of control. The experiment was replicated four times. *Denotes a statistically significant difference (P,0.05).

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Fig. 7. Analysis of CGP apoptosis by in situ TUNEL assay. Cells were plated at a density of 3.2310 4 / cm 2 , and exposed to ethanol (400 mg / dl) for 4 days. Apoptotic index (the number of TUNEL-positive cells divided by the total number of cells counted) was determined. (A) A photomicrograph shows apoptotic cells in control (Ct) and ethanol-treated (Et) cultures. Arrows indicate TUNEL-positive cells. (B) Apoptotic index in control and ethanol-treated cultures. Each data point (6S.E.M.; bars) is the mean of four independent trials. *Denotes a statistically significant difference (P,0.05).

germinal layer (EGL) during the first 2–3 postnatal weeks in the rat [4,50]. During the early postnatal period, cells in EGL undergo extensive proliferation to generate a large pool of cerebellar granule progenitors (CGPs). The developing CGPs then exit the cell cycle, migrate inward past Purkinje cell bodies to their final destination in the internal granule layer (IGL) where they differentiate into cerebellar granule neurons [6,31]. Our study shows that CGPs isolated from 3-day-old rat pups (a time when CGPs are being generated and proliferating in vivo) retain their proliferative capability in vitro. Like us, other investigators demonstrate that CGPs can divide in vitro, and their proliferative activity is directly influenced by the culture environment (e.g. the contents of serum, growth factor or ion supplement) [17,53,55,75]. Actively cycling cells make up 44% of the total population under our experimental conditions, and the proliferation of CGPs in vitro is dependent upon the plating density, i.e. the rate of cell proliferation is increased at higher seeding density (data not shown). This indicates that the CGPs can regulate their own proliferation in an autocrine or paracrine manner as suggested by Gao et al. [26]. The ethanol concentration (400 mg / dl) required for depletion of cultured CGPs is relatively high, but is physiologically relevant to the blood alcohol concentration

(BAC) reported to cause Fetal Alcohol Syndrome [13]. The lack of a dose response suggests that there is a threshold concentration between 200 and 400 mg / dl in which ethanol reduces the number of cultured CGPs. In general, the concentrations of ethanol used for in vitro studies are higher than those required in vivo to produce similar effects. One example relates to the effects of ethanol exposure on cell proliferation. The duration of the cell cycle is increased by 29% in the telencephalic ventricular zone of rats with a BAC of |180 mg / dl [52]. Similarly, the length of the cell cycle of cultured neuroblastoma cells (used as a model for proliferating neuronal cells) is prolonged (137%) by treatment with 400 mg / dl of ethanol [43]. Analysis of cell cycle kinetics reveals that ethanol inhibits CGP proliferation by prolonging the duration of the cell cycle (T C ) and by reducing the growth fraction (GF). Strikingly, ethanol increases the T S of CGPs by more than twofold. The ethanol-induced changes in T C and T S are 15 and 10 h, respectively. Thus, by deduction, ethanol-mediated increases in T C result mainly from prolonged T S , although ethanol may also affect the length of the G 1 -phase. An effect of ethanol on the length of G 2 or M-phase cannot be eliminated, however studies of cultured non-neuronal cells (leukemia cells or hepatoma

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Fig. 9. The necrosis of CGPs in culture. Cells were seeded at a density of 3.2310 4 / cm 2 , and exposed to ethanol (0, 200, and 400 mg / dl) for 4 days. Released activity of LDH, an index of necrotic death, was measured. Each data point (6S.E.M.; bars) is the mean of four independent trials.

Fig. 8. Quantification of apoptosis of CGPs with a cellular DNA fragmentation ELISA. Cells were plated at a density of 3.2310 4 / cm 2 , and exposed to ethanol (0, 200, and 400 mg / dl) for 2 (top panel) or 4 (bottom panel) days. Apoptosis was measured with a cellular DNA fragmentation ELISA as described in the Materials and methods. Each data point (6S.E.M.; bars) is the mean of four independent trials. *Denotes a statistically significant difference (P,0.05).

cells) consistently show that ethanol does not affect T G2 and T M [15,16,32,33]. The finding that ethanol prolongs T C in the CGPs agrees with various in vitro studies using neuron-like cells, such as neuroblastoma cells and PC12 cells [43,46], however, the target site of ethanol action apparently differs. Previous results using transformed cell lines indicate that ethanol primarily prolongs the duration of the G1 phase of the cell cycle, whereas the current study suggests that the S phase of the cell cycle in the CGPs is

the primary target of ethanol. The mechanisms for such differences in response to ethanol are currently unknown. It may reflect distinctive features (such as genetic properties, growth factors, ion channel systems, or intracellular signal components) inherent to different cell types, suggesting that the cellular response to ethanol may be celltype-specific. It has been shown that other exogenous insults targeting the S phase of the cell cycle also induce loss of cerebellar granule neurons. For example, X- or ultraviolet-irradiation is known to inhibit the progression through S-phase of the cell cycle (e.g. Refs. [3,8,74]). Interestingly, X-irradiation during cerebellar development substantially reduces the number of cerebellar granule neurons, and produces behavioral abnormalities with characteristics of Fetal Alcohol Syndrome [5,39,61,65]. It is likely that ethanol and Xirradiation share common mechanisms that perturb development of the cerebellum. The growth fraction of CGPs is significantly (250%) decreased by ethanol. This finding is consistent with our previous study showing the GF of neuroblastoma cells is reduced by in vitro ethanol exposure [43]. The BrdU labeling index (LI) is mainly determined by the GF. A substantial decrease in GF may explain why LI is lower in ethanol-treated cultures, even though the S phase is prolonged. When the entire cycling population is labeled, the extended S phase will not affect the LI. The ethanolinduced decrease in the GF of CGPs may result from (1) a permanent exit of CGPs from the cell cycle, and (2) the preferential killing of cycling cells. Our data support the suggestion that decreased GF results, at least partially,

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from ethanol-induced cell death. The effects of ethanol on apoptosis will be discussed later. Progression of eukaryotic cells from one phase of the cell cycle to another is mediated by the sequential activation and inactivation of a family of protein kinases called cyclin-dependent kinases (CDKs). At least eight CDKs (Cdk1–Cdk8) have been identified in mammalian cells, but only Cdk1, Cdk2, Cdk4 and Cdk6 are required to regulate the cell cycle [7,57]. Cdk2 is considered to be a key protein that regulates both the passage from G1 into S and the actual onset of DNA replication (S-phase) [25,64]. Monomeric CDKs have very low protein kinase activity, and they require binding of regulatory subunits named cyclins as an initial step in their activation process [54,67]. A large family of cyclins (Cyclin A through H) binds and activates different CDKs at different times of the cell cycle. Cyclin A is a key protein that activates Cdk2 to drive cells into S-phase and ensure the orderly progression through S-phase [27,38,64,80]. Cyclin Ds are the major activators that govern the G1–S transition [24,48]. The activity of CDKs is negatively regulated by CDK inhibitors (CDKIs) [68]. Two different families of CDKI (KIP and INK4) have been identified. The KIP family is composed of at least three members, p21, p27, and p57. Members of KIP, such as p21 and p27, are known to inhibit the activity of Cdk2 [7,14,30,69,72,76,77]. The INK4 family includes p14, p15, p18 and p19, and primarily inhibits the activities of Cdk4 and Cdk6 [7,34,64,66]. Our results show that ethanol dramatically down-regulates the activity and the expression of Cdk2, suggesting that cell cycle arrest results from the dysfunction of Cdk2. It appears that ethanol-mediated inhibition of CDK activity does not result from the preferential killing of proliferating cells, at least during early exposure, because the inhibition of CDK activity occurs within 1 day of exposure, whereas ethanol-induced apoptosis is not evident until day 5 of ethanol exposure. Ethanol-induced inhibition may occur as a result of (1) a direct effect on Cdk2 activation, (2) inhibition of Cdk2 activators (such as Cyclin A and Cyclin Ds), (3) promotion of Cdk2 inhibitors (such as p21 and p27), or (4) a combination of these possibilities. Our results show that ethanol significantly down-regulates the expression of Cyclin A and Cyclin D 2 , supporting the suggestion that ethanol inhibits the activators of Cdk2. Especially, cyclin D 2 has been known to play an important role in regulating cerebellar histogenesis, and knocking out Cyclin D 2 dramatically depletes cerebellar granule neurons [35]. Ethanol-induced down-regulation of p27 does not support the idea that ethanol inhibits CDKs by promoting CDKIs. This down-regulation may rather reflect a compensatory response to chronic suppression of Cdk2 activation. Apoptosis is naturally occurring in cultured CGPs (about 7%). It is currently unclear whether the apoptosis occurs in the proliferating CGPs or post-mitotic granule cells. Ethanol induces a delayed apoptosis in CGPs, i.e. apoptosis is

not evident until 4 days of ethanol exposure. There are limited in vitro studies examining the effects of ethanol on the survival of post-mitotic cerebellar granule neurons [40,44,60]. In contrast to the effect on the CGPs, ethanolinduced death of post-mitotic neurons occurs rapidly. At 400 mg / dl, ethanol produces a significant cell loss in post-mitotic cultures of cerebellar granule neurons within 24 h [44,60]. Several mechanisms have been proposed for ethanol-induced death of post-mitotic neurons. These include inhibition of N-methyl-D-aspartate (NMDA) receptors [10], and the blockade of signaling by neurotrophic factors [44,79]. The mechanisms for ethanol-induced apoptosis in the CGPs are unknown. It has been demonstrated that apoptosis is closely linked to cell cycle control. Damage to the cell cycle or DNA integrity is a potent trigger of apoptosis (see reviews [22,23,36,37,49]). Our results show that ethanol-induced cell cycle arrest precedes apoptotic processes, suggesting that delayed apoptosis in CGPs may be a result of severe cell cycle disruption, especially the prolongation of the S-phase. In summary, ethanol-induced loss of cerebellar granule neurons may result from the suppression of proliferation of CGPs. Disturbance of a cyclin-dependent kinase system is a potential mechanism to account for the ethanol-mediated cell cycle arrest in neuronal precursors. It has been shown that ethanol exposure during development depletes neurons in various brain areas, and the temporal window of vulnerability to ethanol is different among different neuronal populations and anatomical locations [47]. The present study shows that CGPs are especially susceptible to ethanol exposure, providing important insights into the cellular and molecular mechanisms underlying the temporal susceptibility to ethanol in the developing cerebellum.

Acknowledgements We thank Drs. William Beresford and James Culberson for their critical reading of this manuscript. This research was supported by grants from the National Institutes of Health (AA12968 and CA 90385).

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