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• W2014002213 abstract "Huntington disease (HD) is caused by a pathological elongation of CAG repeats in the huntingtin protein gene and is characterized by atrophy and neuronal loss primarily in the striatum. Mitochondrial dysfunction and impaired Ca2+ homeostasis in HD have been suggested previously. Here, we elucidate the effects of Ca2+ on mitochondria from the wild type (STHdhQ7/Q7) and mutant (STHdhQ111/Q111) huntingtin-expressing cells of striatal origin. When treated with increasing Ca2+ concentrations, mitochondria from mutant huntingtin-expressing cells showed enhanced sensitivity to Ca2+, as they were more sensitive to Ca2+-induced decreases in state 3 respiration and ΔΨm, than mitochondria from wild type cells. Further, mutant huntingtin-expressing cells had a reduced mitochondrial Ca2+ uptake capacity in comparison with wild type cells. Decreases in state 3 respiration were associated with increased mitochondrial membrane permeability. The ΔΨm defect was attenuated in the presence of ADP and the decreases in Ca2+ uptake capacity were abolished in the presence of Permeability Transition Pore (PTP) inhibitors. These findings clearly indicate that mutant huntingtin-expressing cells have mitochondrial Ca2+ handling defects that result in respiratory deficits and that the increased sensitivity to Ca2+ induced mitochondrial permeabilization maybe a contributing mechanism to the mitochondrial dysfunction in HD. Huntington disease (HD) is caused by a pathological elongation of CAG repeats in the huntingtin protein gene and is characterized by atrophy and neuronal loss primarily in the striatum. Mitochondrial dysfunction and impaired Ca2+ homeostasis in HD have been suggested previously. Here, we elucidate the effects of Ca2+ on mitochondria from the wild type (STHdhQ7/Q7) and mutant (STHdhQ111/Q111) huntingtin-expressing cells of striatal origin. When treated with increasing Ca2+ concentrations, mitochondria from mutant huntingtin-expressing cells showed enhanced sensitivity to Ca2+, as they were more sensitive to Ca2+-induced decreases in state 3 respiration and ΔΨm, than mitochondria from wild type cells. Further, mutant huntingtin-expressing cells had a reduced mitochondrial Ca2+ uptake capacity in comparison with wild type cells. Decreases in state 3 respiration were associated with increased mitochondrial membrane permeability. The ΔΨm defect was attenuated in the presence of ADP and the decreases in Ca2+ uptake capacity were abolished in the presence of Permeability Transition Pore (PTP) inhibitors. These findings clearly indicate that mutant huntingtin-expressing cells have mitochondrial Ca2+ handling defects that result in respiratory deficits and that the increased sensitivity to Ca2+ induced mitochondrial permeabilization maybe a contributing mechanism to the mitochondrial dysfunction in HD. Huntington disease (HD) 2The abbreviations used are: HD, Huntington disease; ΔΨm, mitochondrial membrane potential; PTP, permeability transition pore; 3-NP, 3-nitropropionic acid; KRH, Krebs-Ringer-HEPES; CM-H2TMRos, MitoTracker® Red; JC-1,5,5′,6,6′-tetrachloro-1,1′,3,3′-tetraethylbenzimidazolylcarbocyanine iodide; RCR, respiratory control ratio; FCCP, carbonyl cyanide 4-(trifluoromethoxy)phenylhydrazone; ROS, reactive oxygen species; 4-BrA23187, 4-bromo A-23187, free acid; ANT, adenine nucleotide translocator; ANOVA, analysis of variance; Cab, Ca2+ uptake buffer. 2The abbreviations used are: HD, Huntington disease; ΔΨm, mitochondrial membrane potential; PTP, permeability transition pore; 3-NP, 3-nitropropionic acid; KRH, Krebs-Ringer-HEPES; CM-H2TMRos, MitoTracker® Red; JC-1,5,5′,6,6′-tetrachloro-1,1′,3,3′-tetraethylbenzimidazolylcarbocyanine iodide; RCR, respiratory control ratio; FCCP, carbonyl cyanide 4-(trifluoromethoxy)phenylhydrazone; ROS, reactive oxygen species; 4-BrA23187, 4-bromo A-23187, free acid; ANT, adenine nucleotide translocator; ANOVA, analysis of variance; Cab, Ca2+ uptake buffer. is a neurodegenerative disease that is inherited in an autosomal dominant manner. It belongs to a family of CAG expansion diseases and is caused by the pathological elongation of the CAG repeats in exon one of the huntingtin protein gene (1The Huntington's Disease Collaborative Research GroupCell. 1993; 72: 971-983Abstract Full Text PDF PubMed Scopus (6990) Google Scholar). Symptoms and disease progression are caused by dysfunction and loss of neurons starting in the striatum (specifically medium spiny neurons), but progressing to cortex and to a lesser extent to other brain regions in the later stages of the disease (2Vonsattel J.P. DiFiglia M. J. Neuropathol. Exp. Neurol. 1998; 57: 369-384Crossref PubMed Scopus (1210) Google Scholar). Disease is caused by the toxic gain of function of mutant protein but some loss of function may also contribute to the pathogenesis (for review see Ref. 3Cattaneo E. Rigamonti D. Goffredo D. Zuccato C. Squitieri F. Sipione S. Trends Neurosci. 2001; 24: 182-188Abstract Full Text Full Text PDF PubMed Scopus (303) Google Scholar). The toxic gain of function of mutant huntingtin has not been clearly defined, but there are findings suggesting that mutant huntingtin causes transcriptional dysregulation (4Landles C. Bates G.P. EMBO Rep. 2004; 5: 958-963Crossref PubMed Scopus (348) Google Scholar), ubiquitin-proteasome system dys-function (5Valera A.G. Diaz-Hernandez M. Hernandez F. Ortega Z. Lucas J.J. Neuroscientist. 2005; 11: 583-594Crossref PubMed Scopus (46) Google Scholar), Ca2+ homeostasis dysfunction (6Tang T.S. Tu H. Chan E.Y. Maximov A. Wang Z. Wellington C.L. Hayden M.R. Bezprozvanny I. Neuron. 2003; 39: 227-239Abstract Full Text Full Text PDF PubMed Scopus (397) Google Scholar, 7Panov A.V. Gutekunst C.A. Leavitt B.R. Hayden M.R. Burke J.R. Strittmatter W.J. Greenamyre J.T. Nat. Neurosci. 2002; 5: 731-736Crossref PubMed Scopus (846) Google Scholar), and mitochondrial dysfunction (7Panov A.V. Gutekunst C.A. Leavitt B.R. Hayden M.R. Burke J.R. Strittmatter W.J. Greenamyre J.T. Nat. Neurosci. 2002; 5: 731-736Crossref PubMed Scopus (846) Google Scholar, 8Gu M. Gash M.T. Mann V.M. Javoy-Agid F. Cooper J.M. Schapira A.H. Ann. Neurol. 1996; 39: 385-389Crossref PubMed Scopus (609) Google Scholar, 9Mann V.M. Cooper J.M. Javoy-Agid F. Agid Y. Jenner P. Schapira A.H. Lancet. 1990; 336: 749Abstract PubMed Scopus (114) Google Scholar, 10Browne S.E. Bowling A.C. MacGarvey U. Baik M.J. Berger S.C. Muqit M.M. Bird E.D. Beal M.F. Ann. Neurol. 1997; 41: 646-653Crossref PubMed Scopus (736) Google Scholar). Mitochondrial dysfunction in HD has been suggested primarily by the studies showing impairment of mitochondrial complexes (II, III, and IV) specifically in the striatum in the late stages HD patients (8Gu M. Gash M.T. Mann V.M. Javoy-Agid F. Cooper J.M. Schapira A.H. Ann. Neurol. 1996; 39: 385-389Crossref PubMed Scopus (609) Google Scholar, 9Mann V.M. Cooper J.M. Javoy-Agid F. Agid Y. Jenner P. Schapira A.H. Lancet. 1990; 336: 749Abstract PubMed Scopus (114) Google Scholar, 10Browne S.E. Bowling A.C. MacGarvey U. Baik M.J. Berger S.C. Muqit M.M. Bird E.D. Beal M.F. Ann. Neurol. 1997; 41: 646-653Crossref PubMed Scopus (736) Google Scholar). Administration of the mitochondrial complex II inhibitor 3-nitropropionic (3-NP) in both rodents and nonhuman primates resulted in symptoms and neuropathology that resemble HD (11Beal M.F. Brouillet E. Jenkins B.G. Ferrante R.J. Kowall N.W. Miller J.M. Storey E. Srivastava R. Rosen B.R. Hyman B.T. J. Neurosci. 1993; 13: 4181-4192Crossref PubMed Google Scholar, 12Brouillet E. Hantraye P. Ferrante R.J. Dolan R. Leroy-Willig A. Kowall N.W. Beal M.F. Proc. Natl. Acad. Sci. U. S. A. 1995; 92: 7105-7109Crossref PubMed Scopus (426) Google Scholar). Further, studies have shown impairment of mitochondrial Ca2+ buffering in HD lymphoblast cell line and brain mitochondria from the full-length mutant huntingtin transgenic mice (YAC72) (7Panov A.V. Gutekunst C.A. Leavitt B.R. Hayden M.R. Burke J.R. Strittmatter W.J. Greenamyre J.T. Nat. Neurosci. 2002; 5: 731-736Crossref PubMed Scopus (846) Google Scholar, 13Panov A. Obertone T. Bennett-Desmelik J. Greenamyre J.T. Ann. N. Y. Acad. Sci. 1999; 893: 365-368Crossref PubMed Scopus (23) Google Scholar). Striatum, the primary region to get affected in HD is highly innervated by cortical glutaminergic projections (2Vonsattel J.P. DiFiglia M. J. Neuropathol. Exp. Neurol. 1998; 57: 369-384Crossref PubMed Scopus (1210) Google Scholar). Previously it has been demonstrated that mitochondrial dysfunction can lead to neuronal sensitization to glutamate leading to excitotoxic cellular dysfunction and cell death (14Calabresi P. Gubellini P. Picconi B. Centonze D. Pisani A. Bonsi P. Greengard P. Hipskind R.A. Borrelli E. Bernardi G. J. Neurosci. 2001; 21: 5110-5120Crossref PubMed Google Scholar, 15Henshaw R. Jenkins B.G. Schulz J.B. Ferrante R.J. Kowall N.W. Rosen B.R. Beal M.F. Brain Res. 1994; 647: 161-166Crossref PubMed Scopus (88) Google Scholar). Hence, even though HD is not a classic mitochondrial disease (16Wallace D.C. Science. 1999; 283: 1482-1488Crossref PubMed Scopus (2595) Google Scholar), elucidation of mitochondrial dysfunction mechanisms would likely provide important insight in HD pathogenesis. To study the effects of mutant huntingtin on mitochondrial function, conditionally immortalized cells of striatal origin that express endogenous, comparable levels of either wild type (STHdhQ7/Q7) or mutant (STHdhQ111/Q111) huntingtin were used (17Trettel F. Rigamonti D. Hilditch-Maguire P. Wheeler V.C. Sharp A.H. Persichetti F. Cattaneo E. MacDonald M.E. Hum. Mol. Genet. 2000; 9: 2799-2809Crossref PubMed Scopus (492) Google Scholar). These cell lines are prepared from wild type (HdhQ7/Q7) and mutant huntingtin knock-in mice (HdhQ111/Q111) (17Trettel F. Rigamonti D. Hilditch-Maguire P. Wheeler V.C. Sharp A.H. Persichetti F. Cattaneo E. MacDonald M.E. Hum. Mol. Genet. 2000; 9: 2799-2809Crossref PubMed Scopus (492) Google Scholar) and therefore the STHdhQ111/Q111 cell line is a genetically accurate cell model of HD. In our previous study (18Milakovic T. Johnson G.V. J. Biol. Chem. 2005; 280: 30773-30782Abstract Full Text Full Text PDF PubMed Scopus (206) Google Scholar), we investigated the effects of mutant huntingtin on mitochondrial electron transport chain complexes using STHdhQ7/Q7 and STHdhQ111/Q111 cell lines. Given the fact that the metabolic thresholds and enzyme activities of electron transport chain complexes were not different between the two cell lines, it is likely that the mitochondrial complex deficits are a later event in the course of HD pathogenesis, indeed in low grade HD cases no deficits in the enzyme activities of electron transport chain complexes were observed (19Guidetti P. Charles V. Chen E.Y. Reddy P.H. Kordower J.H. Whetsell W.O. Schwarcz Jr., R. Tagle D.A. Exp. Neurol. 2001; 169: 340-350Crossref PubMed Scopus (165) Google Scholar). In the present study, we examined the effects of Ca2+ on mitochondria from STHdhQ7/Q7 and STHdhQ111/Q111 cells. Isolated mitochondria were treated with increasing Ca2+ concentrations and mitochondrial function was assessed using different assays. We determined that mutant huntingtin-expressing cells have decreased Ca2+ uptake capacity, and exhibit increased sensitivity to Ca2+-induced decreases in respiration and ΔΨm. The ΔΨm defect was attenuated in the presence of ADP and the decrease in Ca2+ uptake capacity was abolished in the presence of Permeability Transition Pore (PTP) inhibitors. This study clearly demonstrates that mitochondrial Ca2+ buffering capacity in STHdhQ111/Q111 cells is compromised, and suggests increased sensitivity to Ca2+-induced mitochondrial permeabilization as a mechanism of mitochondrial dysfunction in HD. Materials—All chemicals were from Sigma-Aldrich unless otherwise noted. All buffers used in experiments with crude mitochondrial preparations were prepared in water (Sigma, catalogue no. 95305) that is standardized for Ca2+ content ([Ca2+]≤ 0.000001%). Cell Culture—In this study, conditionally immortalized striatal progenitor cell lines: STHdhQ7/Q7 cell line expressing endogenous wild type huntingtin and the homozygous mutant STHdhQ111/Q111 cell line expressing comparable levels of mutant huntingtin with 111 glutamines were used. Cell lines were prepared from wild type mice and homozygous HdhQ111/Q111 knock-in mice and were described previously (17Trettel F. Rigamonti D. Hilditch-Maguire P. Wheeler V.C. Sharp A.H. Persichetti F. Cattaneo E. MacDonald M.E. Hum. Mol. Genet. 2000; 9: 2799-2809Crossref PubMed Scopus (492) Google Scholar). Culturing conditions were the same as described in our previous study (18Milakovic T. Johnson G.V. J. Biol. Chem. 2005; 280: 30773-30782Abstract Full Text Full Text PDF PubMed Scopus (206) Google Scholar). Isolation of Mitochondria—Cells were grown on 150-mm plates until ≈ 80–90% confluency, washed twice with cavitation buffer (250 mm sucrose, 5 mm HEPES, 3 mm MgCl2, 1 mm EGTA, pH 7.3 corrected with 5 m KOH) and scraped into cavitation buffer using soft rubber scrapers. Cells were opened using N2 cavitation for 5 min at 250 psi on ice, and samples were additionally homogenized with 1 stroke in a glass Dounce homogenizer. Homogenates were centrifuged at 7000 × g for 10 min at 4 °C. Supernatants were aspirated, and pellets were resuspended in cavitation buffer and used as crude mitochondrial preparations. Protein concentrations in crude mitochondrial preparations were determined using the bicinchoninic acid assay (Pierce), and aliquots were then prepared that contained the indicated protein content for each measurement. Aliquots were centrifuged at 7000 × g for 10 min and kept on ice in cavitation buffer until use in each assay. Measurement of Mitochondrial Respiration—Respiration rates were measured using an oxygraph (Hansatech Instruments) as described previously (18Milakovic T. Johnson G.V. J. Biol. Chem. 2005; 280: 30773-30782Abstract Full Text Full Text PDF PubMed Scopus (206) Google Scholar). Crude mitochondrial preparations (0.5-mg aliquots) were resuspended in respiration buffer (130 mm KCl, 20 mm HEPES, 2 mm MgCl2, 2 mm EGTA, 2 mm potassium phosphate (KH2PO4/K2HPO4, 1:1.78), 1% essentially fatty acids free bovine serum albumin, pH 7.2 adjusted with 5 m KOH) to a final concentration of 1 mg/ml. The mitochondrial suspension (0.5 ml volume) was placed in the respiratory chamber and allowed to equilibrate for 2 min. Respiratory substrate (glutamate (10 mm) plus malate (10 mm) or succinate (5 mm) with rotenone (10 μm)) was then added and state 4 respiration was measured for 2 min, ADP (1.5 mm) was then added, and state 3 respiration was measured for a further 2–4 min. Rates were normalized to citrate synthase activity in the same samples. Citrate synthase activity was determined as previously described (18Milakovic T. Johnson G.V. J. Biol. Chem. 2005; 280: 30773-30782Abstract Full Text Full Text PDF PubMed Scopus (206) Google Scholar). Ca2+ Titration Experiments—Respiration buffers containing specific free Ca2+ concentrations (Ca2+-EGTA respiration buffers) were prepared on the day of the experiment. To calculate the amount of total Ca2+ that was needed to achieve the appropriate free Ca2+ concentration in the respiration buffer that contained 2 mm EGTA we used MaxChelator software (20Patton C. Thompson S. Epel D. Cell Calcium. 2004; 35: 427-431Crossref PubMed Scopus (325) Google Scholar). Each Ca2+-EGTA respiration buffer was prepared separately by diluting each specific 100× CaCl2 stock in the respiration buffer and correcting its pH to 7.2 using 0.1 m KOH in the respiration buffer. CaCl2 stocks were prepared from CaCl2·2H2O (minimum 99%), that was dried overnight and stored in a desiccation chamber until use. Crude mitochondrial preparations were dissolved in prepared Ca2+-EGTA respiration buffers, and respiration rates were measured as described above. The period between buffer addition to the mitochondrial preparation and initiation of state 3 was ∼5 min. Free Ca2+ concentrations in the Ca2+-EGTA buffers were checked using a calibrated Ca2+ electrode on the day of the experiment. Measured concentrations were averaged and presented on the X-axis of Ca2+ titration experiments graphs. Actually concentrations were always slightly higher than those calculated by software. Cytochrome c and NADH Respiration Experiments—Respiration experiments were performed as described above. State 3 respiration was measured for 2 min prior to the addition of cytochrome c (30 μm), and respiration was monitored for another 2 min. This was followed by the addition of NADH (5 mm), and respiration was monitored for an additional 2 min. Determination of Mitochondrial Ca2+ Uptake Capacity—Ca2+ uptake capacities were measured using a Ca2+ electrode (World Precision Instruments). Crude mitochondrial preparation was resuspended in Ca2+ uptake buffer (130 mm KCl, 20 mm HEPES, 2 mm MgCl2, 2 mm potassium phosphate (KH2PO4/K2HPO4, 1:1.78), 1% bovine serum albumin, pH 7.2 adjusted with 5 m KOH) and placed in the oxygraph respiratory chamber. The respiratory chamber was thermostatted at 37 °C, and its contents were constantly mixed with an electromagnetic stirrer bar. Glutamate (10 mm) and malate (10 mm) were added as respiratory substrates. Ca2+ and reference electrodes were added to the chamber from the top. Starting volume of the reaction was 2 ml. The chamber was kept open during an experiment. Ca2+ additions were performed using fine tubing and a Hamilton syringe. 5, 10, and 20 mm CaCl2 stocks were used to make 10, 20, 40, or 80 nmol of Ca2+ additions. The Ca2+ electrode measures extramitochondrial Ca2+ and increases in the signal present as downward deflections on the traces. To observe the effects of PTP inhibition on Ca2+ uptake capacity we used cyclosporine A (1 μm) plus ADP (50 μm) plus oligomycin (2 μg/ml). The PTP inhibitors were added to the respiratory chamber prior to Ca2+ additions (21Panov A.V. Andreeva L. Greenamyre J.T. Arch Biochem. Biophys. 2004; 424: 44-52Crossref PubMed Scopus (52) Google Scholar). To calculate Ca2+ uptake capacity, we counted number of Ca2+ additions until the addition after which no uptake was observed (trace horizontal). The number of additions was multiplied by the nmol of Ca2+ per addition, and normalized to protein content. Mitochondrial Membrane Potential (ΔΨm) Determination in Live Cells—Mitochondrial membrane potential was estimated using the specific mitochondrial probes: Mitotracker Red (CM-H2TMRos) and tetramethylrhodamine ethyl ester (TMRE) (Molecular Probes) (22Leski M.L. Hassinger L.C. Valentine S.L. Baer J.D. Coyle J.T. Synapse. 2002; 43: 30-41Crossref PubMed Scopus (11) Google Scholar, 23Collins T.J. Lipp P. Berridge M.J. Bootman M.D. J. Biol. Chem. 2001; 276: 26411-26420Abstract Full Text Full Text PDF PubMed Scopus (157) Google Scholar, 24Duchen M.R. Cell Calcium. 2000; 28: 339-348Crossref PubMed Scopus (270) Google Scholar, 25Esposti M.D. Hatzinisiriou I. McLennan H. Ralph S. J. Biol. Chem. 1999; 274: 29831-29837Abstract Full Text Full Text PDF PubMed Scopus (160) Google Scholar, 26Krysko D.V. Roels F. Leybaert L. D'Herde K. J. Histochem. Cytochem. 2001; 49: 1277-1284Crossref PubMed Scopus (68) Google Scholar). Cells were grown on poly-l-lysine-coated plates and cultured for 3 days. The cells were then loaded for 30 min with CM-H2TMRos in KRH-glucose, washed, and allowed to equilibrate for 15 min. Cell plates were then mounted in a chamber on the stage of a confocal laser scanning microscope (Leica model TCS SP2). Quantitative measurements of CM-H2TMRos fluorescence were performed by confocal microscopy (Leica model TCS SP2), using a 40× water immersion lens. CM-H2TMRos fluorescence images were obtained by excitation at 563 nm, reflection off a dichroic mirror with a cut-off wavelength at 564 nm, and longpass emission filtering at 590 nm. For TMRE experiments, cells were loaded with TMRE (100 nm) for 45 min in KRH-glucose, and then were mounted on the stage for confocal microscopy. TMRE fluorescence was detected exciting with a 561 nm He-Ne laser line very heavily attenuated (10% laser power), and the emission was collected at >563 nm (22Leski M.L. Hassinger L.C. Valentine S.L. Baer J.D. Coyle J.T. Synapse. 2002; 43: 30-41Crossref PubMed Scopus (11) Google Scholar). Signal from control cells and cells treated with different stimuli were compared using identical settings for laser power, confocal thickness, and detector sensitivity for each dye and separate experiment (23Collins T.J. Lipp P. Berridge M.J. Bootman M.D. J. Biol. Chem. 2001; 276: 26411-26420Abstract Full Text Full Text PDF PubMed Scopus (157) Google Scholar, 25Esposti M.D. Hatzinisiriou I. McLennan H. Ralph S. J. Biol. Chem. 1999; 274: 29831-29837Abstract Full Text Full Text PDF PubMed Scopus (160) Google Scholar, 26Krysko D.V. Roels F. Leybaert L. D'Herde K. J. Histochem. Cytochem. 2001; 49: 1277-1284Crossref PubMed Scopus (68) Google Scholar). The images were analyzed with LCS Leica confocal software and recorded as the mean Mitotracker Red or TMRE fluorescence signal per live cell. Measurement of Mitochondrial Membrane Potential (ΔΨm) in Mitochondrial Preparations—ΔΨm was measured using 5,5′,6,6′-tetrachloro-1,1′,3,3′-tetraethylbenzimidazolylcarbocyanine iodide (JC-1; Molecular Probes) according to a published protocol with modifications (27Ruan Q. Lesort M. MacDonald M.E. Johnson G.V. Hum. Mol. Genet. 2004; 13: 669-681Crossref PubMed Scopus (81) Google Scholar). Modifications were made so that method could be used with isolated mitochondria. To measure ΔΨm at different Ca2+ concentrations, crude mitochondrial preparation was aliquoted into the wells of 96-well plate (50 μg/well). The plate was centrifuged at 3220 × g for 10 min at 4 °C, and supernatants were carefully aspirated. Ca-EGTA respiration buffers with 0, 0.4, or 0.6 μm (software calculated) free Ca2+ or Ca2+ uptake buffer with 150, 500, or 1000 μm Ca2+ each supplemented with glutamate (10 mm), malate (10 mm), and with (for state 3) or without (for state 4) ADP (1.5 mm) were added to separate wells in duplicates (50 μl/well) and incubated at 37 °C for 10 min. Supernatants were carefully aspirated, and the same buffers but containing JC-1 (5 μg/ml) were added to the wells (50 μl/well). Wells in which FCCP (20 μm) was also added were considered as positive controls. The plate was incubated for 30 min, at 37 °C in dark, supernatants were aspirated, and fluorescence was read at 485/528 nm and 530/590 nm. The ratio between the fluorescence was used to describe ΔΨm as published previously (27Ruan Q. Lesort M. MacDonald M.E. Johnson G.V. Hum. Mol. Genet. 2004; 13: 669-681Crossref PubMed Scopus (81) Google Scholar). Measurement of Mitochondrial H2O2 Production—To determine mitochondrial ROS production we used an Amplex Red (Molecular Probes) assay (28Chen Q. Vazquez E.J. Moghaddas S. Hoppel C.L. Lesnefsky E.J. J. Biol. Chem. 2003; 278: 36027-36031Abstract Full Text Full Text PDF PubMed Scopus (1272) Google Scholar). Crude mitochondrial preparations were aliquoted (100 μg/well) and pelleted onto a 96-well plate as described for measuring ΔΨm. Mitochondrial pellets were covered with Ca-EGTA respiration buffers with 0, 0.4, or 0.6 μm (software calculated) free Ca2+ or Ca2+ uptake buffer with 150, 500, or 1000 μm Ca2+ each supplemented with glutamate (10 mm), malate (10 mm), Amplex Red (50 μm), horseradish peroxidase (0.01 units/ml or 0.1 units/ml), and with (for state 3) or without (for state 4) ADP (1.5 mm). Plate was read in the kinetic mode for 30 min at excitation/emission wavelengths 530/590 nm at 37 °C. Rates of H2O2 production were determined using a standard curve. Statistical Analysis—Results were analyzed using ANOVA, Student's t test, or paired t test as indicated. Differences were considered significant if p ≤ 0.05. Effects of Ca2+ on Respiration in Mitochondria from STH-dhQ7/Q7 (Wild Type) and STHdhQ111/Q111 (Mutant) Cells—It has been shown previously that at the free concentrations higher than 1 μm, Ca2+ causes strong inhibition of oxidative phosphorylation (29Moreno-Sanchez R. J. Biol. Chem. 1985; 260: 4028-4034Abstract Full Text PDF PubMed Google Scholar). To determine the effects of Ca2+ on oxidative phosphorylation in mitochondria isolated from the cells expressing endogenous levels of wild type (STHdhQ7/Q7) or mutant (STHdhQ111/Q111) huntingtin, we measured state 4 and state 3 respiration rates in respiration buffers containing increasing free μm Ca2+ concentrations. In these experiments, EGTA-based respiration buffer was used for 0 μm Ca2+ and Ca2+-EGTA respiration buffers were prepared as described under “Experimental Procedures.” At 0 μm Ca2+, we observed no differences in the state 4 or state 3 respiration rates between wild type and mutant cells when glutamate plus malate (complex I substrate) or succinate (complex II substrate) were used as substrates (Fig. 1A). As described earlier (29Moreno-Sanchez R. J. Biol. Chem. 1985; 260: 4028-4034Abstract Full Text PDF PubMed Google Scholar), with increasing free μm Ca2+ concentrations decreases in the state 3 rates were observed (Fig. 1B). However, this decrease was more pronounced in the mitochondria from the mutant huntingtin-expressing cells, reaching significance at lower Ca2+ concentrations than in the wild type (Fig. 1B). State 4 rates increased with increasing Ca2+ concentrations, reaching significance only in the mutant at the highest Ca2+ concentration used (Fig. 1B). To describe overall changes in the respiration rates, we calculated Respiratory Control Ratios (RCRs) at the different Ca2+ concentrations. RCR was calculated as the ratio between state 3 and state 4 rates. A decrease in RCR was observed with increasing Ca2+ concentrations and was more pronounced in mitochondria from mutant cells, reaching significance at the lower Ca2+ concentrations than in the wild type cells (Fig. 1C). These results indicate that mitochondria from STHdhQ111/Q111 (mutant) cells are more sensitive to Ca2+-induced changes in oxidative phosphorylation than mitochondria from STH-dhQ7/Q7 (wild type) cells. Ca2+ Uptake Capacity in Mitochondria from STHdhQ7/Q7 (Wild Type) and STHdhQ111/Q111 (Mutant) Cells—Several studies have suggested that there is reduced mitochondrial Ca2+ buffering capacity in HD. Panov et al. (7Panov A.V. Gutekunst C.A. Leavitt B.R. Hayden M.R. Burke J.R. Strittmatter W.J. Greenamyre J.T. Nat. Neurosci. 2002; 5: 731-736Crossref PubMed Scopus (846) Google Scholar, 13Panov A. Obertone T. Bennett-Desmelik J. Greenamyre J.T. Ann. N. Y. Acad. Sci. 1999; 893: 365-368Crossref PubMed Scopus (23) Google Scholar) demonstrated diminished Ca2+ uptake capacity in mitochondria from HD lymphoblast cell lines, and brain mitochondria from the full-length mutant huntingtin-overexpressing mice (YAC72) (7Panov A.V. Gutekunst C.A. Leavitt B.R. Hayden M.R. Burke J.R. Strittmatter W.J. Greenamyre J.T. Nat. Neurosci. 2002; 5: 731-736Crossref PubMed Scopus (846) Google Scholar), whereas others demonstrated diminished Ca2+ uptake in muscle mitochondria from R6/2 mice (30Gizatullina Z.Z. Lindenberg K.S. Harjes P. Chen Y. Kosinski C.M. Landwehrmeyer B.G. Ludolph A.C. Striggow F. Zierz S. Gellerich F.N. Ann Neurol. 2006; 59: 407-411Crossref PubMed Scopus (65) Google Scholar). To comprehensively describe the effects of Ca2+ on mitochondria in our model, we determined mitochondrial Ca2+ uptake capacity in STHdhQ7/Q7 (wild type) and STHdhQ111/Q111 (mutant) cells. For these experiments we used a Ca2+-sensitive electrode, as described under “Experimental Procedures.” To determine, mitochondrial Ca2+ uptake, isolated mitochondria (1.5 mg/2 ml) were placed in a 37 °C thermostatted chamber and challenged with 10-nmol Ca2+ pulses every 3 min. Representative traces are shown in Fig. 2C. Ca2+ uptake capacity was calculated as described under “Experimental Procedures.” We observed that mitochondria from STHdhQ111/Q111 (mutant) cells have significantly diminished Ca2+ uptake capacity compared with mitochondria from STHdhQ7/Q7 (wild type) cells (Fig. 2A). To determine the “initial uptake” rates we calculated the average of the rates after the second, third, and fourth additions of Ca2+ and determined that the “initial uptake” rates were significantly diminished in the mitochondria from the mutant cells (Fig. 2B). These results indicate that mitochondria from STHdhQ111/Q111 (mutant) cells have a Ca2+-buffering defect, as they can take up less Ca2+ than the mitochondria from wild type cells. Because ER contamination of the mitochondrial preparation was a possibility, we confirmed the mitochondrial nature of the Ca2+ uptake in our mitochondrial preparations, as the addition of uncoupler (FCCP) caused release of Ca2+, and pretreatment of the cells with thapsigargin (which blocks the Ca2+ uptake pump of the ER Ref. 31Paschen W. Doutheil J. Gissel C. Treiman M. J. Neurochem. 1996; 67: 1735-1743Crossref PubMed Scopus (72) Google Scholar) did not produce any change in the Ca2+ uptake capacity (Fig. 2D). Analysis of Mitochondrial Membrane Integrity Before and After Ca2+ Addition in STHdhQ7/Q7 (Wild Type) and STH-dhQ111/Q111 (Mutant) Cells—Ca2+ overload of mitochondria results in increased mitochondrial membrane permeability (32Forte M. Bernardi P. J. Bioenerg. Biomembr. 2005; 37: 121-128Crossref PubMed Scopus (70) Google Scholar). To further study the cause of differences between mitochondria from wild type and mutant cells in their sensitivity to Ca2+, we wanted to determine if the decrease in respiration observed in the presence of free μm Ca2+ concentrations was associated with increased permeability of the mitochondrial membrane. First, we analyzed the integrity of mitochondrial membrane in the basal conditions. NADH is the substrate for mitochondrial complex I. However, the inner mitochondrial membrane is not permeable to exogenous NADH (33Voet D. Voet J.G. Biochemistry. 2nd Ed." @default.
• W2014002213 created "2016-06-24" @default.
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• W2014002213 creator A5050126878 @default.
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• W2014002213 date "2006-11-01" @default.
• W2014002213 modified "2023-10-09" @default.
• W2014002213 title "Mutant Huntingtin Expression Induces Mitochondrial Calcium Handling Defects in Clonal Striatal Cells" @default.
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