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• W1993274829 abstract "In the present study, we have determined the nature and the kinetics of the cellular events triggered by the exposure of cells to non-fibrillar amyloid-β peptide (Aβ). When cortical neurons were treated with low concentrations of soluble Aβ (1–40), an early reactive oxygen species (ROS)-dependent cytoskeleton disruption precedes caspase activation. Indeed, caspase activation and neuronal cell death were prevented by the microtubule-stabilizing drug taxol. A perturbation of the microtubule network was noticeable after being exposed to Aβ for 1 h, as revealed by electron microscopy and immunocytochemistry. Microtubule disruption and neuronal cell death induced by Aβ were inhibited in the presence of antioxidant molecules, such as probucol. These data highlight the critical role of ROS production in Aβ-mediated cytoskeleton disruption and neuronal cell death. Finally, using FRAP (fluorescence recoveryafter photo bleaching) analysis, we observed a time-dependent biphasic modification of plasma membrane fluidity, as early as microtubule disorganization. Interestingly, molecules that inhibited neurotubule perturbation and cell death did not affect the membrane destabilizing properties of Aβ, suggesting that the lipid phase of the plasma membrane might represent the earliest target for Aβ. Altogether our results convey the idea that upon interaction with the plasma membrane, the non-fibrillar Aβ induces a rapid ROS-dependent disorganization of the cytoskeleton, which results in apoptosis. In the present study, we have determined the nature and the kinetics of the cellular events triggered by the exposure of cells to non-fibrillar amyloid-β peptide (Aβ). When cortical neurons were treated with low concentrations of soluble Aβ (1–40), an early reactive oxygen species (ROS)-dependent cytoskeleton disruption precedes caspase activation. Indeed, caspase activation and neuronal cell death were prevented by the microtubule-stabilizing drug taxol. A perturbation of the microtubule network was noticeable after being exposed to Aβ for 1 h, as revealed by electron microscopy and immunocytochemistry. Microtubule disruption and neuronal cell death induced by Aβ were inhibited in the presence of antioxidant molecules, such as probucol. These data highlight the critical role of ROS production in Aβ-mediated cytoskeleton disruption and neuronal cell death. Finally, using FRAP (fluorescence recoveryafter photo bleaching) analysis, we observed a time-dependent biphasic modification of plasma membrane fluidity, as early as microtubule disorganization. Interestingly, molecules that inhibited neurotubule perturbation and cell death did not affect the membrane destabilizing properties of Aβ, suggesting that the lipid phase of the plasma membrane might represent the earliest target for Aβ. Altogether our results convey the idea that upon interaction with the plasma membrane, the non-fibrillar Aβ induces a rapid ROS-dependent disorganization of the cytoskeleton, which results in apoptosis. Alzheimer's disease amyloid-β 2′,7′-dichlorofluorescein diacetate 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide 4,6-diamidino-2-phenylindole phosphate-buffered saline 12-(N-methyl-N-(7-nitrobenz-2-oxa-1,3-diazol-4-yl))-sphingomyelin analysis of variance day in vitro, AMC, 7-amido-4-methylcoumarin fluoromethyketone 2′,7′-dichlorofluorescein A common feature of Alzheimer's disease (AD),1the most common form of dementia, is the accumulation and the aggregation of the amyloid-β peptide (Aβ), a 39- to 43-amino acid peptide derived from the proteolytic cleavage of the amyloid precursor protein (1Yankner B.A. Neuron. 1996; 16: 921-932Google Scholar, 2Selkoe D.J. Nature. 1999; 399 Suppl. 6738: A23-A31Google Scholar). Although Aβ represents a key factor in AD (3Selkoe D.J. J. Neuropathol. Exp. Neurol. 1994; 53: 438-447Google Scholar), the nature of the toxic form of Aβ early involved in AD pathology remains unclear. Whether it is the fibrillar or the non-fibrillar peptides that are the more deleterious remains a controversial issue (4Drouet B. Pincon-Raymond M. Chambaz J. Pillot T. Cell. Mol. Life Sci. 2000; 57: 705-715Google Scholar). The amyloid cascade hypothesis causally links AD clinico-pathological process and neuronal cell death to the aggregation and deposition of Aβ (5Anderson A.J., Su, J.H. Cotman C.W. J. Neurosci. 1996; 16: 1710-1719Google Scholar, 6Estus S. Tucker H.M. van Rooyen C. Wright S. Brigham E.F. Wogulis M. Rydel R.E. J. Neurosci. 1997; 17: 7736-7745Google Scholar, 7Morishima Y. Gotoh Y. Zieg J. Barrett T. Takano H. Flavell R. Davis R.J. Shirasaki Y. Greenberg M.E. J. Neurosci. 2001; 21: 7551-7560Google Scholar). However, this hypothesis has been challenged by recent evidences indicating that the non-fibrillar Aβ also plays a major role in AD (8Michikawa M. Gong J.S. Fan Q.W. Sawamura N. Yanagisawa K. J. Neurosci. 2001; 21: 7226-7235Google Scholar, 9Mucke L. Masliah E., Yu, G.Q. Mallory M. Rockenstein E.M. Tatsuno G., Hu, K. Kholodenko D. Johnson-Wood K. McConlogue L. J. Neurosci. 2000; 20: 4050-4058Google Scholar). A recent elegant study has demonstrated that the fibrils from AD brain are composed of amyloid peptide moieties arranged at right angles to the backbone of the amyloid P protein wrapped in glycosaminoglycans (10Inoue S. Kuroiwa M. Kisilevsky R. Brain Res Rev. 1999; 29: 218-231Google Scholar). Thus, the fibrils are not simply made of chains of self-aggregated Aβ and do not comprise long chains of multimeric Aβ, similar to those used to evaluate the neurotoxicity of the fibrillar Aβ in vitro and in vivo. Moreover, the synaptic loss in AD brain has been correlated with the soluble pool of Aβ peptides rather than the fibrillar one, implying that the non-fibrillar Aβ may be a crucial pathological factor in AD (11McLean C.A. Cherny R.A. Fraser F.W. Fuller S.J. Smith M.J. Beyreuther K. Bush A.I. Masters C.L. Ann. Neurol. 1999; 46: 860-866Google Scholar, 12Lue L.F Kuo Y.M. Roher A.E. Brachova L. Shen Y. Sue L. Beach T. Kurth J.H. Rydel R.E. Rogers J. Am. J. Pathol. 1999; 155: 853-862Google Scholar, 13Wang J. Dickson D.W. Trojanowski J.Q. Lee V.M. Exp. Neurol. 1999; 158: 328-337Google Scholar). Several studies, based on the use of transgenic mice, have demonstrated that neurodegeneration and specific spatial learning deficits might occur without amyloid plaque formation (14Koistinaho M. Ort M. Cimadevilla J.M. Vondrous R. Cordell B. Koistinaho J. Bures J. Higgins L.S. Proc. Natl. Acad. Sci. U. S. A. 2001; 98: 14675-14680Google Scholar, 15Koistinaho M. Kettunen M.I. Goldsteins G. Keinanen R. Salminen A. Ort M. Bures J. Liu D. Kauppinen R.A. Higgins L.S. Koistinaho J. Proc. Natl. Acad. Sci. U. S. A. 2002; 99: 1610-1615Google Scholar, 16Chui D.H. Tanahashi H. Ozawa K. Ikeda S. Checler F. Ueda O. Suzuki H. Araki W. Inoue H. Shirotani K. Takahashi K. Gallyas F. Tabira T. Nat. Med. 1999; 5: 560-564Google Scholar, 17Hsia A.Y. Masliah E. McConlogue L., Yu, G.Q. Tatsuno G., Hu, K. Kholodenko D. Malenka R.C. Nicoll R.A. Mucke L. Proc. Natl. Acad. Sci. U. S. A. 1999; 96: 3228-3233Google Scholar). These results emphasize the necessity to clarify the initial response of neurons to the non-fibrillar Aβ and to identify the cellular targets involved in non-fibrillar Aβ-induced neurotoxicity. Tailing with these observations, our studies and others rely on the hypothesis of a close association between neuronal loss and a proapoptotic effect of soluble forms of Aβ (18Pillot T. Drouet B. Queillé S. Labeur C. Vandekerckhove J. Rosseneu M. Pinçon-Raymond M. Chambaz J. J. Neurochem. 1999; 73: 1626-1634Google Scholar, 19Drouet B. Fifre A. Pinçon-Raymond M. Vandekerckhove J. Rosseneu M. Gueant J.L. Chambaz J. Pillot T. J. Neurochem. 2001; 76: 117-127Google Scholar, 20Lambert M.P. Barlow A.K. Chromy B.A. Edwards C. Freed R. Liosatos M. Morgan T.E. Rozovsky I. Trommer B. Viola K.L. Wals P. Zhang C. Finch C.E. Krafft G.A. Klein W.L. Proc. Natl. Acad. Sci. U. S. A. 1998; 95: 6448-6453Google Scholar, 21Walsh D.M. Lomakin A. Benedek G.B. Condron M.M. Teplow D.B. J. Biol. Chem. 1997; 272: 22364-22372Google Scholar). Indeed, it has been established that the amphiphilic non-aggregated Aβ may intercalate into the plasma membrane of neurons, directly altering membrane activities and inducing neuronal cell death (18Pillot T. Drouet B. Queillé S. Labeur C. Vandekerckhove J. Rosseneu M. Pinçon-Raymond M. Chambaz J. J. Neurochem. 1999; 73: 1626-1634Google Scholar, 22Drouet B. Pinçon-Raymond M. Chambaz J. Pillot T. J. Neurochem. 1999; 73: 758-769Google Scholar, 23Rhee S.K. Quist A.P. Lal R. J. Biol. Chem. 1998; 273: 13379-13382Google Scholar, 24Favit A. Grimaldi M. Nelson T.J. Alkon D.L. Proc. Natl. Acad. Sci. U. S. A. 1998; 95: 5562-5567Google Scholar, 25Lin H. Bhatia R. Lal R. FASEB J. 2001; 15: 2433-2444Google Scholar). Accumulative evidences have laid emphasis on the critical role of an oxidative stress in AD and in the neurotoxicity induced by Aβ (26Varadarajan S. Yatin S. Aksenova M. Butterfield D.A. J Struct Biol. 2000; 130: 184-208Google Scholar, 27Miranda S. Opazo C. Larrondo L.F. Munoz F.J. Ruiz F. Leighton F. Inestrosa N.C. Prog. Neurobiol. 2000; 62: 633-648Google Scholar). However, most of the molecular mechanisms involved in the neuronal cell death induced by non-fibrillar forms of Aβ are yet to be characterized. The aim of the present paper was to identify the primary targets of the non-fibrillar Aβ and the chronology of the early cellular events involved in apoptotic neuronal cell death upon Aβ exposure. Microtubules fulfill a plethora of cellular functions, including axonal and dendritic growth and stability (28Sheetz M.P. Nat. Rev. Mol. Cell. Biol. 2001; 2: 392-396Google Scholar, 29Baasm P.W. Curr. Opin. Cell Biol. 1997; 9: 29-36Google Scholar). We have proved recently that the non-fibrillar Aβ (1–40) induces apoptosis to rat cortical neurons (18Pillot T. Drouet B. Queillé S. Labeur C. Vandekerckhove J. Rosseneu M. Pinçon-Raymond M. Chambaz J. J. Neurochem. 1999; 73: 1626-1634Google Scholar, 22Drouet B. Pinçon-Raymond M. Chambaz J. Pillot T. J. Neurochem. 1999; 73: 758-769Google Scholar). In the present study, we have investigated whether the microtubule network could be an early cellular target for the non-fibrillar Aβ. We have demonstrated the following sequence: low concentrations of soluble Aβ (1–40), or of its shorter (29–40) C-terminal domain, induce apoptotic neuronal cell death by perturbing the fluidity of the plasma membrane, leading to a disruption of the neurotubule network depending on the induction of an early oxidative stress. Aβ (1–40), Aβ (29–40), the caspase substrate, and inhibitor peptides were purchased from Bachem, and DCFH-DA was from Molecular Probes. Unless otherwise indicated, materials used for cell culture were obtained from Invitrogen. The drug stabilizing cytoskeleton, taxol, and all other chemicals were of high purity grade from Sigma. To overcome problems of amyloid peptide solubility at high concentrations, fresh peptide stock solutions were prepared at 5 mg × ml−1 in hexafluoro-2-propanol (Sigma) as described previously (30Pillot T. Goethals M. Vanloo B. Talussot C. Brasseur R. Rosseneu M. Vandekerckhove J. Lins L. J. Biol. Chem. 1996; 271: 28757-28765Google Scholar). For the incubation of the peptides with the neurons, aliquots of peptide stock solution were quickly dried under nitrogen and directly solubilized at the experimental concentrations into the culture medium. Peptide solutions were then applied onto the cells. Under those conditions, all the amyloid peptides remained soluble for the determination of their neurotoxic properties (18Pillot T. Drouet B. Queillé S. Labeur C. Vandekerckhove J. Rosseneu M. Pinçon-Raymond M. Chambaz J. J. Neurochem. 1999; 73: 1626-1634Google Scholar). Cortical neurons from embryonic day 16–17 Wistar rat fetuses were prepared as described previously (18Pillot T. Drouet B. Queillé S. Labeur C. Vandekerckhove J. Rosseneu M. Pinçon-Raymond M. Chambaz J. J. Neurochem. 1999; 73: 1626-1634Google Scholar). Briefly, dissociated cells were plated at 4.5–5.0 104cells/cm2 in plastic dishes pre-coated with poly-l-ornithine (1.5 μg × ml−1; Sigma). The cells were cultured in a chemically defined Dulbecco's modified Eagle's medium-F12 medium free of serum (Invitrogen) and supplemented with insulin (5.10−7m), putrescine (60 μm), sodium selenite (30 nm), transferrin (100 μm), progesterone (1.10−7m), and 0.1% (w/v) ovalbumin. Cultures were kept at 35 °C in a humidified 6% CO2atmosphere. After six to seven DIV, cortical population was determined to be at least 95% neuronal by immunostaining as described previously (18Pillot T. Drouet B. Queillé S. Labeur C. Vandekerckhove J. Rosseneu M. Pinçon-Raymond M. Chambaz J. J. Neurochem. 1999; 73: 1626-1634Google Scholar, 31Keller J.N. Pang Z. Geddes J.W. Begley J.G. Germeyer A. Waeg G. Mattson M.P. J. Neurochem. 1997; 69: 273-284Google Scholar). Experiments were performed on six to seven DIV neurons. Cell viability was first determined by morphological observation and cell counting after 5 min of trypan blue staining (0.4%; Sigma) to evaluate membrane integrity, and the metabolic activity was assessed by the MTT reduction assay (18Pillot T. Drouet B. Queillé S. Labeur C. Vandekerckhove J. Rosseneu M. Pinçon-Raymond M. Chambaz J. J. Neurochem. 1999; 73: 1626-1634Google Scholar, 22Drouet B. Pinçon-Raymond M. Chambaz J. Pillot T. J. Neurochem. 1999; 73: 758-769Google Scholar). Moreover, the release of lactate dehydrogenase into the culture medium was assessed using a cytotoxicity detection kit (Roche Molecular Biochemicals) according to the recommendations of the manufacturer. Cell nuclei were visualized using 4,6-diamidino-2-phenylindole (DAPI; Sigma). The cells, grown on a glass coverslips, were washed in PBS, incubated at room temperature for 10 min with DAPI (0.1 μg × ml−1), washed with PBS, and examined under a microscope equipped for epifluorescence. To evaluate the percentage of apoptotic cells, five independent fields of microscope were counted (around 100 cells) in three separate experiments, with two determinations each. Under control conditions, neuronal cells exhibited 12–15% of apoptotic cells at nine DIV. For experiments in the presence of caspase inhibitors, cells were incubated 2 h with 50 or 100 μm inhibitors before and throughout Aβ peptide exposure. Alternatively, DNA fragmentation was monitored by enzyme-linked immunosorbent assay for the detection of oligonucleosomes using a kit form purchased from Roche Molecular Biochemicals. Briefly, cortical neurons were plated in 24-well dishes (around 200.000 cells per well) and treated at seven DIV for 24 h with Aβ. After having been washed, the cells were lysed directly on wells, and oligonucleosomes were determined according to the recommendations of the manufacturer. The caspase activities were measured by means of the cleavage of the substrates DEVD-pNA, YVAD-pNa, LEHD-AMC, and IEPD-AMC (Bachem). Briefly, at the indicated time points following peptide treatments, the cells were rinsed three times with ice-cold PBS and incubated for 20 min on ice in a buffer of 25 mm Hepes, pH 7.5, 1% (v/v) Triton X-100, 5 mm EDTA, 1 mm EGTA, 5 mm MgCl2, 5 mm dithiothreitol, 1 mm phenylmethylsulfonyl fluoride, 10 μg/ml each of pepstatin and leupeptin, and 5 μg/ml aprotinin. The lysate was centrifuged for 15 min at 12,000 rpm and assayed for protein by Bradford (Bio-Rad). 50 μg of proteins were incubated for 2 h with 100 μm caspase substrates initially dissolved in Me2SO. The cleavage of the caspase substrates was monitored by absorbance measurements at 405 nm for DEVD-pNA and YVAD-pNa and by fluorescence emission at 460 nm after exciting LEHD-AMC and IEPD-AMC at 360 nm, using a Fluostar reader plate (BMG-Labtechnologies). The measurement of cell oxidation is based on the oxidation of the non-fluorescent compounds, DCFH-DA, to a fluorescent derivative, DCF, in a peroxidase-mediated reaction. Increases in fluorescence emission reflect enhanced cellular oxidative stress. Briefly, treated cortical neurons were loaded with 100 μm DCFH-DA for 45 min. Before analysis, cells were washed three times in PBS, and DCF fluorescence was recorded directly on culture dishes by a Fluostar reader plate (BMG-Labtechnologies), using 488-nm excitation and 510-nm emission filters. Cortical neurons were fixed for 2 h at 4 °C in 2.5% glutaraldehyde and 0.5% tannic acid in 0.1m cacodylate buffer, pH 7.4. Then the neurons were postfixed for 2 h at 4 °C in 2% osmic acid in phosphate buffer. After having been dehydrated in a graded alcohol series, the samples were embedded in Epon resin (Poly/Bed 812; Polysciences, Warrington, PA), and ultra thin sections (70-nm) were obtained using a Reichert Ultracut. Thin sections were counterstained with uranyl acetate, and lead citrate and examined with a Jeol 100CX microscope. For immunofluorescence studies, the neurons were cultured on glass coverslips that had been coated overnight with 15 μg/ml poly l-ornithine. Following the treatments, the neurons were fixed in PBS containing 4% paraformaldehyde for 30 min at room temperature. The cells were permeabilized with 0.1% Triton X-100 made up in PBS containing 3% bovine serum albumin for 30 min and then incubated with a monoclonal anti-β-tubulin antibody (1:500) (Chemicon) for 1 h under constant agitation. After several washes in PBS, the cells were incubated for 1 h with a fluorescein isothiocyanate-conjugated donkey anti-mouse IgG (1: 250) (Santa Cruz Biotechnology), washed with PBS, labeled with DAPI as described above, and mounted in Fluoprep (BioMérieux). The microtubules were visualized with a Nikon microscope using a PlanFluor X40/1.3 objective. For semiquantitative analysis of microtubule organization, at least five microscope fields/condition were imaged using a Nikon DXM1200 digital camera, and microtubule organization in 100–120 cells/field was classified as normal, mildly disrupted, or severely disrupted. The cells were incubated for shorter times with Aβ (1–40) in Hanks' balanced salt solution at room temperature and loaded with 4 μm NBD-SM (1 mg/ml stock solution in chloroform). In the case of longer treatments (up to 24 h), the neurons were incubated with Aβ (1–40) in normal culture conditions, then rinsed with Hanks' balanced salt solution and stained with the fluorescent lipid, and analyzed 10 min thereafter. The measurement of the lateral diffusion of the molecules in the membrane by means of FRAP (fluorescence recoveryafter photo bleaching) has already been described (32Blonk C.G. Don A. Van Aalst H. Birmingham J.J. J. Microscopy. 1992; 169: 363-374Google Scholar). A fluorescent probe, here a labeled lipid, was incorporated into the cell membranes. When a small defined area of the labeled membrane was photobleached by a high powerful laser pulse, the intensity of fluorescence was reduced immediately. As the fluorescent probe present in the vicinity underwent a constant diffusion motion, it diffused into the bleached area, gradually increasing the fluorescence intensity after bleaching. Thus, the kinetics of the fluorescent recovery depends on the diffusion rate of the probe, measured as the diffusion coefficient, D 20,w. Analyses were performed at 37 °C with a Zeiss LSM 510 confocal laser scanning microscope. We used the 488-nm line of a 25-milliwatt argon laser with a Zeiss C-Apochromat, ×63, numerical aperture 1.2, oil-immersion objective. The pinhole diameter was set to 1 airy unit, which correspond to a 1.2-μm depth of field, to reduce the contribution of cytoplasm fluorescence as much as possible. The area of bleaching was defined as a circle of 3.0-μm diameter, centered on an apical body cell membrane. The area was photobleached at full laser power (100% power, 100% transmission) for 230 ms. The extent of the bleaching typically reached 50 to 80%. Before bleaching, five images were monitored to define the initial fluorescence. The post-bleached images were scanned at 0.6% transmission with a delay of 100 ms during the last 60 s, resulting in a total acquisition of 80 points for 90 s, the time required to complete the maximum recovery of fluorescence. No photobleaching was observed during recovery. The sets of scans in which the fraction mobile of fluorophore was less than 65% or more than 110% and bleaching less than 35% were discarded. Cells presenting debris, heterogeneous labeling, or movements during scanning were not used for FRAP measurements.D 20,w parameters were calculated according to the method described by Kubitscheck et al.(33Kubitscheck U. Wedekind P. Peters R. Biophys. J. 1994; 67: 948-956Google Scholar). STAT VIEW computer software was used for the statistical analysis. Most of the data were from three separate experiments with three to four determinations each. Values were expressed as means + S.E. Differences between control and treated groups were analyzed using Student's t test. Multiple pairwise comparisons among the groups of data were performed using ANOVA followed by a Scheffe's post hoc test. Statistical differences were determined at p < 0.05. To investigate the kinetics of microtubule network disorganization, cortical neurons were exposed to 5 μmsoluble Aβ (1–40) for short incubation times at the end of which no morphological feature of apoptosis (e.g. membrane bleeding, cell shrinkage, and chromatin condensation) was detected (see Figs.1 and 2). After a 3-h Aβ (1–40) exposure, we observe a dramatic perturbation of the neurotubule network in most of the neurites of the treated neurons (Fig. 1 c), as compared with the control cells in which neurotubules elongated within the neurites normally, in a parallel organization (Fig. 1 a). Similar results were obtained using the shorter Aβ (29–40) which displays also membrane perturbing properties (data not shown). These observations made through electron microscopy were confirmed using immunocytochemistry. In the control cells, a dense and constant microtubule network radiates from the cell bodies to the periphery (Fig. 2, A and B). By contrast, in the cortical neurons treated with 5 μmnon-fibrillar Aβ (1–40) for 1 and 3 h (Fig. 2, C andD, respectively), we observed a peripheral fragmentation and loss of microtubules without any fragmentation or condensation of nuclear DNA (Fig. 2 C). Despite this severe microtubule disruption, the treated cells maintained their spreading shape, implying that the non-fibrillar Aβ (1–40) did not affect neurofilaments. Higher peptide concentrations or longer incubation times resulted in a more extensive and rapid loss of the microtubule network (not shown).Figure 2The non-fibrillar Aβ (1–40) peptide disrupts neuronal microtubules.Cortical neurons were incubated in the absence (A andB) or in the presence of 5 μm soluble Aβ (1–40) (C and D) for 1 and 3 h, respectively. The microtubule organization was visualized using immunofluorescence with an anti β-tubulin monoclonal antibody.View Large Image Figure ViewerDownload (PPT) To determine the kinetics of onset of the early microtubule perturbations induced by the non-fibrillar Aβ (1–40) and of the apoptotic neuronal cell death, we investigated the effects of taxol, a microtubule-stabilizing drug, on Aβ-induced cytoskeleton disruption and neurotoxicity. Cortical neurons incubated with 100 nm taxol only displayed a typical microtubule organization (Fig. 1, b and b′) as described previously (34Michaelis M.L. Ranciat N. Chen Y. Bechtel M. Ragan R. Hepperle M. Liu Y. Georg G. J. Neurochem. 1998; 70: 1623-1627Google Scholar). Interestingly, the non-fibrillar Aβ (1–40) was unable to disrupt the neuronal microtubule network in the cells preincubated for 2 h with 100 nm taxol before Aβ (1–40) addition (Fig. 1 D). These results have been confirmed using immunocytochemistry and with taxol being added to the cortical neurons at the same time as Aβ (1–40) (not shown). We next investigated the effects of taxol on non-fibrillar Aβ-induced neuronal cell death. As described previously (18Pillot T. Drouet B. Queillé S. Labeur C. Vandekerckhove J. Rosseneu M. Pinçon-Raymond M. Chambaz J. J. Neurochem. 1999; 73: 1626-1634Google Scholar), the treatment of cortical neurons with 5 μm Aβ (1–40) resulted in a time-dependent decrease in cell viability monitored by the MTT assay (Fig. 3 A). Upon a 6-h exposure to Aβ (1–40), the MTT reduction level decreased significantly to 18.3% (p < 0.05) compared with the control level. Interestingly, whereas taxol alone displayed no effect on MTT even after 48 h of treatment, its presence protected the neurons against Aβ (1–40)-induced neurotoxicity (Fig. 3 A). After a 48-h exposure to Aβ in the presence of 100 nm taxol, the MTT reduction level remained at 82% of control, whereas the protective effects of taxol diminished after prolonged incubations. Moreover, the presence of taxol in the culture medium inhibited the release of lactate dehydrogenase after a 48-h Aβ (1–40) treatment (Fig. 3 B). This suggests that the stabilization of the microtubule organization by taxol prevents Aβ-induced neuronal cell death. In agreement with these morphological and biochemical observations, we demonstrated that taxol inhibited apoptosis induced by low concentrations of non-fibrillar Aβ (1–40). The apoptotic nuclei were visualized and quantified after DAPI staining of cultures treated with 5 μm Aβ (1–40) in the absence and presence of 100 nm taxol. Upon Aβ exposure, cortical neurons shared a time-dependent increase in the number of apoptotic nuclei, which was statistically different from control after 24 h of incubation and reached 58.6 + 3.1% (p < 0.001) after 48 h of incubation (Fig. 4 A). The presence of taxol in the culture medium almost completely inhibited the apoptotic cell death induced by Aβ (1–40) after a 24-h incubation, and the effects persisted for up to 48 h (23.4 + 2.5% of apoptotic nuclei) (Fig. 4 A). To further improve our results, we quantified the cytoskeleton perturbation induced by the non-fibrillar Aβ (1–40) using immunocytochemistry with an anti-β-tubulin antibody. The Aβ-induced microtubule disruption was time-dependent (Fig. 4 B). As early as 1 h after the addition of 5 μm soluble Aβ (1–40) to the cells, we observed a greater number of neurons exhibiting a mild disruption of microtubules (25.2 + 1.8%, p < 0.01). After a 3-h incubation, the microtubule network was mildly or severely disrupted in 35.2 + 3.5% (p < 0.001) of the treated cells, and almost all the neurons displayed disturbed microtubules after a 24-h incubation (Fig. 4 B). Interestingly, the presence of taxol completely abolished microtubule perturbation induced by the non-fibrillar Aβ peptide (Fig. 4 B). These data strongly emphasize that early cytoskeleton disruption is required in the neuronal cell death induced by non-fibrillar Aβ peptide. Moreover, Fig. 4 C showed only few cells exhibiting both disorganized microtubules and apoptotic nuclei. Altogether, these results strongly suggest that microtubule perturbations precede, and might be involved in, a pathway leading to apoptosis upon soluble Aβ exposure. In a previous report, we demonstrated that the caspase 3 inhibitor, DEVD-CHO peptide, reduced neuronal cell death induced by the non-fibrillar Aβ (1–40) significantly (18Pillot T. Drouet B. Queillé S. Labeur C. Vandekerckhove J. Rosseneu M. Pinçon-Raymond M. Chambaz J. J. Neurochem. 1999; 73: 1626-1634Google Scholar). Here, we clearly show that the apoptotic cell death induced by non-fibrillar Aβ (1–40) requires the activation of caspases 3 and 9 by directly measuring caspase-like activity in the lysates of the treated cells (Fig. 5). The activity of caspases 3 and 9 increased significantly (p < 0.05 as compared with the control cells) after a 6-h incubation with 5 μm Aβ (1–40) (Fig. 5, A and D, respectively), whereas the activation of caspases 1 and 8 was not detected (Fig. 5,B and C, respectively). To establish a causative relationship between microtubule disruption and caspase activation, we performed kinetic experiments of microtubule perturbation in the absence and presence of caspase inhibitors (Fig. 6). Fig. 6 A demonstrates that both caspase inhibitors markedly reduced apoptosis induced by 5 μm non-fibrillar Aβ (1–40). However, unlike taxol, the caspase inhibitors had no effect on the early cytoskeleton perturbations induced by Aβ (1–40), even after a prolonged incubation (Fig. 6 B). Moreover, the presence of 100 nm taxol during Aβ exposure inhibited the activation of caspases 3 and 9 significantly (not shown). These results suggest that the microtubule perturbation might occur before caspase activation under treatment with non-fibrillar Aβ (1–40).Figure 6Effects of caspase inhibitors on soluble Aβ (1–40)-induced apoptosis and microtubule disruption. Cortical neurons were preincubated or not for 2 h with 100 μm caspase inhibitors and then treated for the indicated incubation time with 5 μmnon-fibrillar Aβ (1–40). Apoptotic nuclei were visualized and quantified after DAPI staining (A), and microtubule perturbations were quantified by immunofluorescence using an antibody against β-tubulin (B). Data are means (± S.E.) of three different experiments with four determinations each and are normalized to the effect of vehicle, designated as 100% (*, p < 0.05; **, p < 0.01; ***, p < 0.001). No significant differences were found between caspase inhibitor-treated and control cells.View Large Image Figure ViewerDownload (PPT) We showed that cell exposure to 5 μmnon-fibrillar Aβ (1–40) peptide induced a time-dependent increase ROS formation, as measured by the oxidative stress-sensitive dye DCFH-DA. A 1- to 3-h treatment with Aβ caused a significant increase in ROS production (Fig. 7). During these short incubation times," @default.
• W1993274829 created "2016-06-24" @default.
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• W1993274829 date "2003-01-01" @default.
• W1993274829 modified "2023-10-17" @default.
• W1993274829 title "Apoptotic Neuronal Cell Death Induced by the Non-fibrillar Amyloid-β Peptide Proceeds through an Early Reactive Oxygen Species-dependent Cytoskeleton Perturbation" @default.
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