Prion Protein Fragment PrP-(106–126) Induces Apoptosis via Mitochondrial Disruption in Human Neuronal SH-SY5Y Cells
2001; Elsevier BV; Volume: 276; Issue: 47 Linguagem: Inglês
10.1074/jbc.m103894200
ISSN1083-351X
AutoresConor N. O'Donovan, Deirdre Tobin, Thomas G. Cotter,
Tópico(s)Trace Elements in Health
ResumoThe synthetic peptide PrP-(106–126) has previously been shown to be neurotoxic. Here, for the first time, we report that it induces apoptosis in the human neuroblastoma cell line SH-SY5Y. The earliest detectable apoptotic event in this system is the rapid depolarization of mitochondrial membranes, occurring immediately upon treatment of cells with PrP-(106–126). Subsequent to this, cytochrome c release and caspase activation were observed. Caspase inhibitors demonstrated that while the peptide activates caspases they are not an absolute requirement for apoptosis. Parallel to caspase activation, PrP-(106–126) was also observed to trigger a rise in intracellular calcium through release of mitochondrial calcium stores. This leads to the activation of calpains, another family of proteases. A calpain inhibitor demonstrated that while calpains are activated by the peptide they also are not an absolute requirement for apoptosis. Interestingly a combination of caspase and calpain inhibitors significantly inhibited apoptosis. This illustrates alternative pathways leading to apoptosis via caspases and calpains and that blocking both pathways is required to inhibit apoptosis. These results implicate the mitochondrion as a primary site of action of PrP-(106–126). The synthetic peptide PrP-(106–126) has previously been shown to be neurotoxic. Here, for the first time, we report that it induces apoptosis in the human neuroblastoma cell line SH-SY5Y. The earliest detectable apoptotic event in this system is the rapid depolarization of mitochondrial membranes, occurring immediately upon treatment of cells with PrP-(106–126). Subsequent to this, cytochrome c release and caspase activation were observed. Caspase inhibitors demonstrated that while the peptide activates caspases they are not an absolute requirement for apoptosis. Parallel to caspase activation, PrP-(106–126) was also observed to trigger a rise in intracellular calcium through release of mitochondrial calcium stores. This leads to the activation of calpains, another family of proteases. A calpain inhibitor demonstrated that while calpains are activated by the peptide they also are not an absolute requirement for apoptosis. Interestingly a combination of caspase and calpain inhibitors significantly inhibited apoptosis. This illustrates alternative pathways leading to apoptosis via caspases and calpains and that blocking both pathways is required to inhibit apoptosis. These results implicate the mitochondrion as a primary site of action of PrP-(106–126). scrapie isoform of prion protein native cellular prion protein permeability transition change in mitochondrial membrane potential scrambled phosphate-buffered saline N-benzyloxycarbonyl-Val-Ala-Asp(O-Me) fluoromethyl ketone 3-[4,5-dimethylthiazol-2-yl]-2,5-diphenyltetrazolium bromide propidium iodide 5,5′,6–6′-tetrachloro-1,1′,3,3′-tetraethylbenzimidazolecarbocyanine iodide 530/30 band pass filter 585/42 band pass filter 1,2-bis(2-aminophenoxy)ethane-N,N,N′,N′-tetraacetic acid tetrakis- (acetoxymethyl ester) Prion-related encephalopathies are a family of neurodegenerative disorders including conditions such as scrapie in sheep, bovine spongiform encephalopathy in cattle, and Creutzfeldt-Jakob disease and Gerstmann-Straussler-Scheinker syndrome among others in humans. They are characterized by vacuolation of the neuropil, neuronal loss, and gliosis (1Masters C.L. Richardson Jr., E.P. Brain. 1978; 101: 333-344Crossref PubMed Scopus (309) Google Scholar). In many cases this is also accompanied by the extracellular accumulation of the scrapie isoform (PrPSC)1 of the normal cellular prion protein (PrPC), which can aggregate into fibrils in the extracellular matrix (2Mikol J. Biomed. Pharmacother. 1999; 53: 19-26Crossref PubMed Scopus (11) Google Scholar). PrPSC is widely believed to be the infectious agent of these diseases (3Prusiner S.B. Science. 1991; 252: 1515-1522Crossref PubMed Scopus (1749) Google Scholar), and the formation of PrPSC is thought to be via a post-translational conformational change by which PrPC complexes with PrPSC to yield two molecules of PrPSC (4Prusiner S.B. Proc. Natl. Acad. Sci. U. S. A. 1998; 95: 13363-13383Crossref PubMed Scopus (5167) Google Scholar). PrPC is a cell surface protein mainly expressed in the neuronal and glial cells of the central nervous system. The exact function of PrPC remains unknown, although recent studies have implicated it in copper metabolism (5Pauly P.C. Harris D.A. J. Biol. Chem. 1998; 273: 33107-33110Abstract Full Text Full Text PDF PubMed Scopus (543) Google Scholar, 6Ruiz F.H. Silva E. Inestrosa N.C. Biochem. Biophys. Res. Commun. 2000; 269: 491-495Crossref PubMed Scopus (75) Google Scholar) and signal transduction (7Mouillet-Richard S. Ermonval M. Chebassier C. Laplanche J.L. Lehmann S. Launay J.M. Kellermann O. Science. 2000; 289: 1925-1928Crossref PubMed Scopus (678) Google Scholar). Its expression is necessary for the pathogenesis of the spongiform encephalopathies (8Bueler H. Aguzzi A. Sailer A. Greiner R.A. Autenried P. Aguet M. Weissmann C. Cell. 1993; 73: 1339-1347Abstract Full Text PDF PubMed Scopus (1814) Google Scholar). Apoptosis is a physiologically important cellular suicide pathway, which has also been implicated in a number of pathological conditions (9Gorman A.M. McGowan A. O'Neill C. Cotter T. 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A synthetic peptide corresponding to residues 106–126 of human PrP (PrP-(106–126)) has previously been found to induce apoptosis in primary rat hippocampal cultures (13Forloni G. Angeretti N. Chiesa R. Monzani E. Salmona M. Bugiani O. Tagliavini F. Nature. 1993; 362: 543-546Crossref PubMed Scopus (896) Google Scholar), primary mouse cerebellar cultures (14Brown D.R. Schmidt B. Kretzschmar H.A. Nature. 1996; 380: 345-347Crossref PubMed Scopus (499) Google Scholar), the rat pituitary clonal cell line GH3 (15Florio T. Thellung S. Amico C. Robello M. Salmona M. Bugiani O. Tagliavini F. Forloni G. Schettini G. J. Neurosci. Res. 1998; 54: 341-352Crossref PubMed Scopus (79) Google Scholar), and more recently in mouse retinae in vivo (16Ettaiche M. Pichot R. Vincent J.P. Chabry J. J. Biol. Chem. 2000; 275: 36487-36490Abstract Full Text Full Text PDF PubMed Scopus (127) Google Scholar). PrP-(106–126) is one of a number of peptides corresponding to sequences within fragments of amyloid proteins isolated from the brains of patients suffering from Gerstmann-Straussler-Scheinker syndrome (17Tagliavini F. Prelli F. Ghiso J. Bugiani O. Serban D. Prusiner S.B. Farlow M.R. Ghetti B. Frangione B. EMBO J. 1991; 10: 513-519Crossref PubMed Scopus (171) Google Scholar). It retains the ability of PrPSC to aggregate into amyloid-like fibrils and the tendency to adopt a mostly β-sheet structure (18Tagliavini F. Prelli F. Verga L. Giaccone G. Sarma R. Gorevic P. Ghetti B. Passerini F. Ghibaudi E. Forloni G. Proc. Natl. Acad. Sci. U. S. A. 1993; 90: 9678-9682Crossref PubMed Scopus (235) Google Scholar). Residues 106–126 of PrP constitute a region maintained in all the PrP isoforms that have been found to accumulate in the brains of patients suffering from prion diseases. Under normal physiological conditions a catabolic pathway also leads to cleavage of this region of the prion protein at residues 110 and 111 (19Chen S.G. Teplow D.B. Parchi P. Teller J.K. Gambetti P. Autilio-Gambetti L. J. Biol. Chem. 1995; 270: 19173-19180Abstract Full Text Full Text PDF PubMed Scopus (453) Google Scholar). The importance of the 106–126 sequence of the prion protein makes it a useful model for the in vitro study of prion-induced cell death. In this study we determine, for the first time, the effect of PrP-(106–126) in human neuroblastoma cells in vitro and demonstrate its ability to induce apoptosis. Furthermore, we establish the mitochondrion as a target of PrP-(106–126) action and show that the peptide activates two distinct biochemical pathways involving caspases and calpains, which both result in apoptosis. PrP-(106–126) (KTNMKHMAGAAAAGAVVGGLG) and PrP-(106–126) scrambled (scr) (AVHTGLGAMAALNMVVGGAAGL) were synthesized and purified by MWG Biotech (Milton Keynes, UK). Peptides were dissolved in sterile phosphate-buffered saline (PBS) to a concentration of 1 mm before use and were freshly prepared before each experiment. The inhibitors z-VAD-fmk (Enzymes Systems Products, Livermore, CA) and calpeptin (Sigma) were added to cells 15 min prior to drug/peptide treatments. The degree of peptide aggregation was measured fluorimetrically by thioflavin-T binding (20Baumann M.H. Kallijarvi J. Lankinen H. Soto C. Haltia M. Biochem. J. 2000; 349: 77-84Crossref PubMed Scopus (32) Google Scholar). Aged preparations of peptides in PBS were added to 50 mmglycine, pH 9, with 2 μm thioflavin-T (Sigma) to a concentration of 100 μm. Samples were incubated at room temperature for 5 min, and then fluorescence was measured on a Spectramax Gemini fluorometer with excitation and emission maxima of 435 and 485 nm, respectively. Samples were prepared in triplicate. The adherent human neuroblastoma cell line SH-SY5Y was cultured in RPMI 1640 medium (Life Technologies, Inc.) supplemented with 10% fetal calf serum (Life Technologies, Inc.), 1% penicillin/streptomycin (Life Technologies, Inc.), and 1% l-glutamine (Life Technologies, Inc.). Cells were maintained at 37 °C in a humidified 5% CO2atmosphere. For experiments, cells were maintained in serum-free RPMI 1640 medium supplemented with 1% Growth Medium Supplement-A (Life Technologies, Inc.), 1% penicillin/streptomycin, and 1%l-glutamine. When being passaged or harvested for analysis cells were lifted using trypsin/EDTA. Cytotoxicity was assessed by the conversion of MTT (Sigma) to a formazan product. After appropriate incubation of cells with peptides, MTT was added to each well to a final concentration of 0.25 mg/ml and then incubated for 4 h at 37 °C. Microtiter plates were then centrifuged at 200 ×g for 5 min. The reaction was terminated by removal of the supernatant and addition of 100 μl of Me2SO to each well. Following thorough mixing to dissolve the formazan product, the plates were read at 620 nm on a microELISA plate reader. Assays were performed in replicate of four samples. Cell viability was assessed by flow cytometry that monitored annexin V binding and propidium iodide (PI) uptake simultaneously. After appropriate incubation with drugs/peptides, cells were resuspended in annexin V binding buffer and then treated with annexin V (1×) and PI (5 μg/ml) for 5 min at room temperature. Samples were then analyzed by fluorescence on a FACScan flow cytometer (Becton Dickinson, Oxford, UK). Fluorescence was measured through a 530/30 band pass filter (FL-1) to monitor annexin V binding and through a 585/42 band pass filter (FL-2) to monitor PI uptake. An initial increase in FL-1 fluorescence is indicative of apoptosis before an increase in FL-2, indicative of secondary necrosis, is observed. DNA was isolated and electrophoresed on 10% agarose gels by the method of McGahon et al. (21McGahon A.J. Martin S.J. Bissonnette R.P. Mahboudi A. Shi Y. Mogil R.J. Nishioka W.K. Green D.R. Methods Cell Biol. 1995; 46: 153-185Crossref PubMed Scopus (523) Google Scholar). Cells were lysed in RIPA buffer (50 mm Tris-HCl (pH 7.4), 1% Nonidet P-40, 0.25% sodium deoxycholate, 1 mm EGTA, 1 mmNa3VO4, 1 mm NaF, 150 mm NaCl, 0.1 mm phenylmethylsulfonyl fluoride, and 1 μl of protein inhibitor mixture (1 μg/ml antipain, 1 μg/ml aprotinin, 1 μg/ml chymostatin, 0.1 μg/ml leupeptin, and 1 μg/ml pepstatin)) and put on ice for 40 min. Samples were then centrifuged at 20,800 × g for 10 min. The supernatant was then transferred into Eppendorf tubes, and protein concentrations were determined using a Bio-Rad protein assay reagent. Protein was separated by SDS-polyacrylamide gel electrophoresis (15% gel, 30 μg of protein/sample) and then transferred to a nitrocellulose membrane. Proteins were detected using appropriate antibodies and the ECL detection reagent (Amersham Pharmacia Biotech). The antibodies used were mouse monoclonal anti-human Bcl-2 (DAKO, Cambridge, UK), rabbit monoclonal anti-Bcl-XL, (Calbiochem), rabbit monoclonal anti-Bax (DAKO, Cambridge, UK), monoclonal anti-cytochrome c(DAKO), and monoclonal anti-cytochrome c oxidase (DAKO). Cells were harvested and treated with 10 μmJC-1 (Molecular Probes, Leiden, Netherlands) for 15 min at 37 °C (22Salvioli S. Ardizzoni A. Franceschi C. Cossarizza A. FEBS Lett. 1997; 411: 77-82Crossref PubMed Scopus (897) Google Scholar). Mitochondrial membrane potential was then measured by fluorescence emission on a FACScan flow cytometer (Becton Dickinson). Fluorescence emission was collected through FL-1 on a log scale. FL-1 measures the fluorescence of JC-1 monomers, which increase in number as the mitochondrial membrane depolarizes. Cells were washed in 2 ml of PBS, fixed in 1% paraformaldehyde on ice for 30 min, and then washed in 1 ml of IFA-Tx (4% fetal calf serum, 10 mm HEPES (pH 7.4), 0.1% sodium azide, 0.1% Triton X-100, 150 mmsaline). Samples were then resuspended in 150 μl of IFA-Tx containing primary antibody (rabbit polyclonal anti-active caspase-3, 1:600 dilution (Pharmingen, San Diego, CA)), put on ice for 1 h, centrifuged at 200 × g, and washed twice in 1 ml of IFA-Tx. IFA-Tx was aspirated off, and to the remaining drop in each sample, 2 μl of secondary antibody (fluorescein isothiocyanate-labeled anti-rabbit (Sigma)) was added. Samples were left for 1 h on ice in darkness and then washed twice in 1 ml of IFA-Tx. Samples were then resuspended in 0.5 ml of PBS, and fluorescence was measured on a FACScan flow cytometer through FL-1. An increase in FL-1 fluorescence above the control is indicative of caspase activity. Samples treated with an irrelevant antibody (rabbit IgG (Sigma)) and with secondary antibody only were used as controls. Cells were washed once in PBS and then resuspended and lysed in 40 μl of 10 mm HEPES, 50 mm NaCl on ice for 40 min. After lysis the samples were centrifuged at 1000 × g for 5 min to pellet the membrane fraction. The supernatant (cytosolic fraction) was transferred to a clean Eppendorf tube, and the membrane fraction was resuspended in 40 μl of 10 mm HEPES, 50 mm NaCl. A protein determination assay was performed on both fractions, and an equal amount of protein was loaded into each well of a 96-well plate (ideally 100 μg of protein/well). To each well, 32 μl of fluorescent substrate (Suc-Leu-Tyr (AFL-117) (Enzyme Systems Products)) was added, and the total volume of each well was brought to 200 μl with imidazole buffer (100 mm imidazole, 5 mml-cysteine, 1 mm mercaptoethanol, 10 mm CaCl2, 4% Me2SO in H2O). Samples were incubated at 37 °C for 30 min after the addition of the substrate. Fluorescence was then measured on a fluorometer (Spectra Max Gemini, Molecular Devices) with excitation and emission wavelengths of 400 and 505 nm, respectively. Cells were washed once in PBS and then resuspended in 650 μl of PBS. Next 3.25 μl of 10 mmmonochlorobimane (Molecular Probes) was added to each sample. Samples were measured in triplicate, so 200 μl of each sample was aliquoted into three wells of a dark 96-well plate. Samples were incubated at room temperature for 15 min in darkness and then read on a fluorometer (Spectra Max Gemini, Molecular Devices) with excitation and emission maxima of 395 and 482 nm, respectively. The activity of catalase was measured by following the decrease in absorbance at 240 nm due to H2O2 decomposition (23Aebi H. Methods in Enzymology. 105. Academic Press, Inc., Orlando, FL1984: 121-126Google Scholar). Activity was measured as rate of change of absorbance (ΔO.D./min). Changes in intracellular calcium concentration were detected by loading cells with FLUO-3 (1×) (Molecular Probes) for 15 min prior to sample collection and measurement of fluorescence on a FACScan flow cytometer. Fluorescence emission was collected through FL-1 on a log scale, and an increase in FL-1 fluorescence is indicative of an increase in calcium levels. Cells were harvested, washed in 1 ml of homogenizing medium (1 msucrose, 1 m Tris-HCl (pH 7.4), 50 mm EGTA, and 1% bovine serum albumin), resuspended in 80 μl of homogenizing medium, and Dounce-homogenized. Samples were centrifuged at 1000 × g for 5 min at 4 °C, and the supernatant was transferred to a clean Eppendorf tube. The supernatant was centrifuged at 10,000 × g for 5 min at 4 °C. The pellet was then washed in 100 μl of Wash 1 (1 m sucrose, 1m Tris-HCl, 1% bovine serum albumin (pH 7.4), 50 mm EGTA, and 1 m KCl), centrifuged at 10,000 × g for 5 min at 4 °C, and washed in 100 μl of Wash 2 (1 m mannitol, 1 m KCl, 1m Tris-HCl, 1% bovine serum albumin (pH 7.4)). Finally the sample was centrifuged at 10,000 × g for 5 min at 4 °C and resuspended in RIPA buffer (see "Western Blotting"). Cytochrome c was visualized by Western blot analysis using anti-cytochrome c (Pharmingen). The supernatant was retained as the cytosolic fraction. PrP-(106–126) has previously been reported to induce cell death as a result of its ability to form aggregates (13Forloni G. Angeretti N. Chiesa R. Monzani E. Salmona M. Bugiani O. Tagliavini F. Nature. 1993; 362: 543-546Crossref PubMed Scopus (896) Google Scholar, 24Hope J. Shearman M.S. Baxter H.C. Chong A. Kelly S.M. Price N.C. Neurodegeneration. 1996; 5: 1-11Crossref PubMed Scopus (65) Google Scholar). The aggregation status of the peptide in this study was ascertained by means of a thioflavin-T assay (Fig.1 A). Thioflavin-T fluorescence increases in the presence of protein aggregates. PrP-(106–126) was seen to aggregate immediately at day 0 when prepared at 100 μm in PBS, and over time it gradually increases its aggregation status. The scrambled version of the peptide shows little aggregation even over time. PrP-(106–126) was observed to induce cell death in a dose-dependent manner over time as measured by MTT assay (Fig. 1 B). MTT is converted to a formazan product by mitochondrial enzymes, which become inactive as the cell dies. Measurement of this formazan product is an indicator of cell viability. The mechanism of cell death induced by PrP-(106–126) was shown to be apoptosis by annexin V binding (Fig. 1 C). Annexin V binds to phosphatidylserine, which flips from the inner to the outer leaflet of the cell membrane during apoptosis (73Martin S.J. Reutelingsperger C.P. McGahon A.J. Rader J.A. van Schie R.C. LaFace D.M. Green D.R. J. Exp. Med. 1995; 182: 1545-1556Crossref PubMed Scopus (2570) Google Scholar). Apoptosis could be detected as early as 2 h after treatment when 22% of the cell population were annexin-positive. Cells were incubated with annexin V and PI simultaneously The population of cells in the lower left quadrant represents viable cells (Fig. 1 C). An increase in FL-1 represents annexin V-positive cells in the lower right quadrant. This is the apoptotic population. The final shift in FL-2 up to the top right quadrant represents PI-positive cells, indicative of membrane permeability and secondary necrosis. The percentage of the overall population in each quadrant is given in the circles. The mechanism of cell death was further confirmed to be via apoptosis by the observation of DNA ladder patterns upon electrophoresis of isolated DNA from peptide treated cells (Fig.1 D). The internucleosomal cleavage of DNA into ∼200-base pair fragments is a typical biochemical hallmark of apoptosis in a number of systems (74McKenna S.L. Carmody R.J. Cotter T.G. Cell Engineering. Kluwer Academic Publishers, Dordrecht, The Netherlands1999Google Scholar). No effects were observed upon treatment with PrP-(106–126)scr. A number of typical apoptotic events are observed to occur after treatment of cells with 100 μmPrP-(106–126). Mitochondrial dysfunction characterized by a loss of transmembrane potential has been found to be a central event in many cases of apoptosis (25Carmody R.J. Cotter T.G. Sepsis. 1998; 2: 9-19Crossref Scopus (4) Google Scholar). Analysis of Ψm using the lipophilic fluorescent probe JC-1 demonstrates extensive loss of Ψm (represented by an increase in fluorescence of FL-1 due to increased numbers of JC-1 monomers) within 15 min of treatment with 100 μmPrP-(106–126) (Fig. 2 A). PrP-(106–126)scr (100 μm) had no significant effect. Although the response is maximal at 15 min, mitochondrial membrane depolarization is observed to commence immediately upon treatment of the cells with PrP-(106–126) when fluorescence in FL-1 is measured as a function of time on the FACScan flow cytometer (Fig. 2 B). Also within this 15-min time frame, cytochrome c is released from the mitochondria into the cytosol (Fig. 2 C), and caspase-3 activity is easily detectable by measuring the fluorescence of an anti-active caspase-3 antibody on a flow cytometer (Fig.2 D). Cytochrome cis a protein of the mitochondrial intermembrane space that is commonly released during apoptosis following membrane depolarization, which leads to caspase activation. The caspases are a family of serine threonine proteases responsible for the cleavage of a number of substrates during apoptosis (42Perkins C.L. Fang G. Kim C.N. Bhalla K.N. Cancer Res. 2000; 60: 1645-1653PubMed Google Scholar). Despite these early apoptotic events, at 15 min apoptosis is not detectable by annexin binding. In view of the rapid effects of PrP-(106–126) on the mitochondria we investigated possible effects on the Bcl-2 family of proteins. These are important regulators of apoptosis, composed of both pro- and antiapoptotic members, and are known to operate at the level of the mitochondrion (26Bruce-Keller A.J. Begley J.G. Fu W. Butterfield D.A. Bredesen D.E. Hutchins J.B. Hensley K. Mattson M.P. J. Neurochem. 1998; 70: 31-39Crossref PubMed Scopus (175) Google Scholar, 27Kitada S. Andersen J. Akar S. Zapata J.M. Takayama S. Krajewski S. Wang H.G. Zhang X. Bullrich F. Croce C.M. Rai K. Hines J. Reed J.C. Blood. 1998; 91: 3379-3389Crossref PubMed Google Scholar, 28Suzuki M. Youle R.J. Tjandra N. Cell. 2000; 103: 645-654Abstract Full Text Full Text PDF PubMed Scopus (909) Google Scholar). We examined the intracellular levels of Bcl-2, an antiapoptotic protein, when cells were treated with 100 μm PrP-(106–126) (Fig. 3 A). After a 15-min treatment with 100 μm PrP-(106–126) there was a dramatic reduction of Bcl-2 protein as compared with the untreated control. PrP-(106–126)scr (100 μm) caused no change in the levels of Bcl-2. We also examined the effects of the peptide on other Bcl-2 family members, Bcl-XL(antiapoptotic) and Bax (proapoptotic) (Fig. 3 A). In contrast to what was observed with Bcl-2, we found no change in the intracellular levels of Bcl-XL or Bax in response to 100 μm PrP-(106–126). As a caspase cleavage site was previously reported to be present in Bcl-2 (29Kirsch D.G. Doseff A. Chau B.N. Lim D.S. de Souza-Pinto N.C. Hansford R. Kastan M.B. Lazebnik Y.A. Hardwick J.M. J. Biol. Chem. 1999; 274: 21155-21161Abstract Full Text Full Text PDF PubMed Scopus (428) Google Scholar), the effect of z-VAD, a pan caspase inhibitor previously reported to block apoptosis, on Bcl-2 degradation was investigated (Fig. 3 B). The reduction of Bcl-2 induced by 100 μm PrP-(106–126) was found to be inhibited in cells that were pretreated with 100 or 200 μm z-VAD. Cells treated with 200 μm z-VAD or 100 μm PrP-(106–126)scr alone showed no change in their Bcl-2 levels. A concentration of 100 μm z-VAD was shown to block caspase-3 activity as measured by anti-active caspase-3 antibody on the FACScan flow cytometer (Fig. 3 C). Thus, the inhibition of caspases maintains the protein levels of the antiapoptotic Bcl-2 even in the presence of PrP-(106–126). The same treatment of cells with z-VAD did not inhibit mitochondrial depolarization (Fig. 3 D), placing this event upstream of caspase activation and Bcl-2 degradation. More importantly, z-VAD did not inhibit apoptosis despite blocking caspase-mediated Bcl-2 degradation, indicating that caspases are not necessary for PrP-(106–126) to induce cell death. With caspases found to be nonessential for apoptosis in this system, another mechanism must be at work. Maintaining focus on the mitochondrion, as this was the site of the earliest observed effect of PrP-(106–126), we investigated the possible role of oxidative stress due to disruption of mitochondrial function as this is the primary site of intracellular reactive oxygen species production. Oxidative stress has been shown to have a role in a number of apoptotic systems (30Verhaegen S. McGowan A.J. Brophy A.R. Fernandes R.S. Cotter T.G. Biochem. Pharmacol. 1995; 50: 1021-1029Crossref PubMed Scopus (170) Google Scholar, 31Slater A.F.G. Stefan C. Nobel I. van den Dobbelsteen D.J. Orrenius S. Cell Death Differ. 1996; 3: 57-62PubMed Google Scholar). We examined the intracellular levels of peroxides, superoxide anion, and nitric oxide as well as the activities of Mn-superoxide dismutase and Cu,Zn-superoxide dismutase (data not shown). We also looked at glutathione levels using the fluorescent monochlorobimane probe (Fig.4 A) and the activity of catalase by following the decrease in absorbance at 240 nm due to H2O2 decomposition (Fig. 4 B) in response to PrP-(106–126). Glutathione not only acts as a reactive oxygen species scavenger but also functions in the regulation of the intracellular redox state, and catalase is the primary defense mechanism against H2O2. None of the aforementioned changed appreciably in response to PrP-(106–126). Staurosporine (0.25 μm) and valinomycin (50 nm) were used as positive controls as both these drugs induce oxidative stress in SH-SY5Y cells. The peptide was not found to predispose the cells to death by a secondary oxidative insult either (data not shown). Another function of the mitochondrion is in the regulation of intracellular calcium levels, therefore we investigated whether PrP-(106–126) could be exerting an effect through the deregulation of calcium homeostasis. Using the fluorescent probe FLUO-3 we demonstrated an intracellular rise in calcium levels (represented by an increase in FL-1 fluorescence) in response to PrP-(106–126) (Fig.5 A). The response is immediate, and calcium levels reach a sustained peak by 30 s. This experiment was repeated in the presence of EGTA, a calcium chelator, to show that the source of the calcium was intracellular (data not shown). BAPTA-AM, an intracellular calcium chelator, was found to inhibit this increase in calcium levels, confirming the intracellular nature of the source of the calcium rise (Fig. 5 B). The two major calcium stores in the cell are the endoplasmic reticulum and the mitochondria. To determine which was releasing calcium into the cytosol in this system we used thapsigargin, which causes a rapid release of endoplasmic reticulum calcium stores. Upon treatment of cells with thapsigargin, an increase in calcium levels was observed using the FLUO-3 probe. Once that response had reached its maximum, cells were treated with 100 μm PrP-(106–126), and a further increase in calcium was observed (Fig. 5 C). This indicates that PrP-(106–126) releases calcium from a site other than the endoplasmic reticulum. The mitochondria make up the only other store of calcium in the cell large enough to account for the PrP-(106–126)-induced rise in calcium levels. We next examined the activity of calpains in response to the PrP-(106–126)-induced rise in intracellular calcium. The calpains are a group of calcium-activated proteases that have also been implicated in apoptosis. They can exist in active and inactive forms associated with the cell membrane or the cytosol (32Ray S.K. Wilford G.G. Matzelle D.C. Hogan E.L. Banik N.L. Ann. N. Y. Acad. Sci. 1999; 890: 261-269Crossref PubMed Scopus (60) Google Scholar, 33Ishihara I. Minami Y. Nishizaki T. Matsuoka T. Yamamura H. Neurosci. Lett. 2000; 279: 97-100Crossref PubMed Scopus (36) Google Scholar, 34Debiasi R.L. Squier M.K. Pike B. Wynes M. Dermody T.S. Cohen J.J. Tyler K.L. J. Virol. 1999; 73: 695-701Crossref PubMed Google Scholar). Within 15 min of treating cells with 100 μm PrP-(106–126) an increase in calpain activity was evident in both the cells in cytosolic and membrane fractions as measured using a fluorescent calpain substrate (Fig. 6 A). Pretreatment of cells with the calpain inhibitor calpeptin was found to inhibit the activity of the calpains induced by PrP-(106–126) but not cell death. However, when cells were pretreated with a combination of 100 μm z-VAD and 100 μm calpeptin, apoptosis induced by 100 μm PrP-(106–126) was significantly inhibited (Fig. 6 B). The toxicity of the PrP-(106–126) peptide in a human neuronal cell line has not been demonstrated previously. Host expression of PrPC has been illustrated to be a prerequisite of PrP-(106–126) toxicity (14Brown D.R. Schmidt B. Kretzschmar H.A. Nature. 1996; 380: 345-347Crossref PubMed Scopus (499) Google Scholar). Accordingly we observed the presence of PrPC in the SH-SY5Y cell line upon Western blot analysis (data not shown). In some instances the neurotoxicity of PrP-(
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