Download Post-Translational Modifications and Dysfunction of Mitochondrial

Articles in PresS. Am J Physiol Endocrinol Metab (July 12, 2016). doi:10.1152/ajpendo.00127.2016
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Post-Translational Modifications and Dysfunction of
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Mitochondrial Enzymes in Human Heart Failure
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Freya L Sheeran1 and Salvatore Pepe1,2,3
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Heart Research, Clinical Sciences, Murdoch Childrens Research Institute, and
Department of Paediatrics, University of Melbourne, Royal Children’s Hospital,
Melbourne, Australia.
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Department of Surgery at Alfred Hospital, Monash University, Melbourne, Australia.
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Address for Correspondence:
Associate Professor Salvatore Pepe, Department of Cardiology, Royal Children’s
Hospital, 50 Flemington Road, Parkville VIC 3052, Australia.
Email: [email protected]
Key Words: human heart failure, mitochondria, oxidative stress
Copyright © 2016 by the American Physiological Society.
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Abstract
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Deficiency of energy supply is a major complication contributing to the syndrome of
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heart failure (HF). As the concurrent activity profile of mitochondrial bioenergetic
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enzymes has not been studied collectively in human HF, our aim was to examine the
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mitochondrial enzyme defects in left ventricular myocardium obtained from
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explanted end-stage failing hearts. Compared to non-failing donor hearts, activity
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rates of complexes I and IV and the Krebs cycle enzymes isocitrate dehydrogenase,
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malate dehydrogenase and aconitase were lower in HF, as determined
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spectrophotometrically. However, activity rates of complexes II, III and citrate
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synthase did not differ significantly between the two groups. Protein expression,
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determined by western blotting, did not differ between the groups, implying post-
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translational perturbation. In the face of diminished total glutathione and coenzyme
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Q10 levels, oxidative modification was explored as an underlying cause of enzyme
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dysfunction. Of the three oxidative modifications measured, protein carbonylation
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was significantly increased by 31% in HF (p<0.01; n=18), while levels of 4-
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hydroxynonenal and protein nitration though elevated, did not differ. Isolation of
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complexes I, IV and F1FoATP synthase by immunocapture revealed that proteins
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containing iron-sulphur or heme redox centres were targets of oxidative
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modification. Energy deficiency in end-stage failing human left ventricle involves
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impaired activity of key electron transport chain and Krebs cycle enzymes, without
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altered expression of protein levels. Augmented oxidative modification of crucial
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enzyme subunit structures implicates dysfunction due to diminished capacity for
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management of mitochondrial reactive oxygen species, thus contributing further to
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reduced bioenergetics in human HF.
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Key Words –heart failure, mitochondria, respiration, oxidative stress
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Introduction
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Energy supply deficit in the failing human heart has been well established in
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contributing to the decline in cardiac function (3, 24, 52) and is a key prognostic
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marker for patient mortality, similar to NYHA class alone (34). It has been proposed
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that energy starvation leads to the progressive worsening of heart failure (16),
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limiting energy available for cellular contraction, enzyme activities, cellular repair
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and membrane turnover, and if levels reach a critical threshold can ultimately lead to
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apoptotic or necrotic cell death (20, 57). Mitochondrial oxidative phosphorylation
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(OXPHOS) in mammalian cells accounts for up to 90% of ATP supply and has been
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reported to decline in human heart failure (47). While a small number of studies
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have found decreases in the activities of individual OXPHOS complexes I (2, 22, 44),
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III (7, 17) and IV (2, 7, 40), the concurrent global contribution of these enzymes and
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Krebs enzymes that support NADH supply to OXPHOS have not previously been
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reported in end-stage human heart failure.
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The aim of the present study was to characterize where specific enzyme dysfunction
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occurs concurrently in the mitochondrial Krebs and oxidative phosphorylation
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pathways in the end-stage failing human heart. Activities of the electron transport
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chain complexes I-IV, the Krebs cycle enzymes isocitrate dehydrogenase (ICDH),
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malate dehydrogenase (MDH), α-ketoglutarate dehydrogenase (KGDH), aconitase
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and citrate synthase (CS) were measured and protein expression of the OXPHOS
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enzymes quantified to determine if altered activities related to changes in protein
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content. As superoxide release from complex I is a major source of ROS production,
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mitochondrial-derived ROS was explored as a potential cause of enzyme dysfunction.
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Methods
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Tissue Processing
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Left ventricular (LV) tissues from non-failing donor (NF, n=20) and end-stage
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(explanted) failing hearts (HF, n=30; 18 ischemic heart disease, 12 dilated
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cardiomyopathy) were obtained by patient consent from the Alfred Hospital
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(Melbourne, Australia) at the time of heart transplantation and approved by the
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Alfred Hospital Human Ethics Committee for Discarded Tissue Research. The NF
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donor hearts that were excluded from transplantation due to unavailable timely
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match or technical constraints were approved for research by donor family consent
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and Victorian Organ Donation Service, Australian Red Cross. Patient age was similar
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between the two groups (NF: 52 ± 4 years; HF: 51 ± 2 years (Mean ± SEM), with a
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higher proportion of males in HF compared to the non-failing group (NF:
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13male/7female vs HF: 25male/5female). Samples were immediately snap-frozen in
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liquid N2 upon collection and stored at −80°C. For enzyme assays and western
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blotting, tissues were ground under LN2 in a pre-chilled mortar and pestle and
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homogenized using a glass Dounce homogenizer in 10x volume of ice-cold buffer (0.1
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M KH2PO4, pH 7.4 with 1% mammalian protease inhibitor cocktail). Samples were
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centrifuged at 600g to remove unbroken cells and nuclei and the supernatants
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stored at −80°C until required. Protein content was estimated using the BCA Assay
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(Sigma Aldrich, MO, USA).
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Enzyme Assays
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Activity of complexes I, II, III and IV was measured using a Hewlett-Packard model
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8453 spectrophotometer, essentially as described by Birch-Machin (5). Activities of
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complexes I, II, III and IV were measured spectrophotometrically in the absence and
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presence of inhibitors specific for the respective complexes (4) and are expressed as
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inhibitor-sensitive activities. CS activity was measured according to the methods by
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Srere (49), while activities of ICDH and KGDH were measured using the methods of
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Benderdour (4) and Lucas (25) respectively. MDH activity was determined by
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incubating 33 µl tissue extract with reaction buffer (0.2 mM oxalacetic acid in 0.1 M
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KH2PO4, pH 7.4; 1 ml final volume) for 2 minutes at 25°C. The reaction was initiated
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by the addition of NADH (0.26 mM final) and the absorbance monitored at 340 nm
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for 4 minutes. Aconitase activity was measured as described by Maack (26) using a
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coupled aconitase-ICDH enzyme reaction using the reduction of NADP+ at 340 nm for
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detection. Pyruvate dehydrogenase (PDH) activity was measured as described by
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Hansford (15). Measurement of pyridine nucleotides was made using the
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NAD+/NADH quantification kit (Biovision Inc. CA, USA) by means of a plate-reader
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based enzyme-cycling assay. Extracts were prepared from frozen tissues according
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to the kit protocol and were assayed on 2 parallel plates, one for total NAD (NADt)
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and one for reduced NAD (NADH). For NADH only, prior to assay, 100 μl extract was
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heated at 60°C for 30 minutes, followed by cooling, to decompose NAD; for NADt,
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extracts were assayed untreated. NAD concentrations were determined using
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supplied reference standards and expressed as nmol NAD/mg protein. Oxidized NAD
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was calculated as [NADt] minus [NADH].
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Western Blotting
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Protein expression of representative subunits from complexes I, II, III, IV, V, ANT and
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porin was quantified on individual blots using standard western blotting procedures.
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For each blot, 1 μg tissue extract was loaded per lane and run on a 15% SDS-PAGE
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gel. Bands were transferred to a PVDF membrane for 1 hour at 100 V. Membranes
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were briefly washed in Tris-buffered saline (TBS) + 0.05% Tween-20 (TBST) and
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blocked for 1 hour at RT with TBST + 5% skim milk powder (blocking buffer). For
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each respective blot, membranes were incubated overnight with primary antibody at
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the following concentrations: complex I (39 kDa subunit) 0.2 μg/mL; complex II (70
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kDa subunit) 0.1 μg/mL; complex III (core I subunit) 0.1 μg/mL; complex IV (Vib kDa
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subunit) 1 μg/mL; complex IV (mt-DNA encoded COX I subunit) 1 μg/mL; F1FoATP
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synthase (β subunit) 0.1 μg/mL; porin 0.5 μg/mL. All OXPHOS antibodies were
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monoclonal, obtained from Molecular Probes, USA. Polyclonal antibody to ANT was
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purchased from Calbiochem, CA, USA, and used at a working concentration of 2.5
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μg/mL. Blots for Complexes I-V and ANT were normalized to expression of porin,
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which was probed on the same membranes and expressed as a density ratio. For
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Krebs cycle enzymes, primary antibodies (Abcam, MA, USA) were used at the
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following concentrations: mitochondrial isocitrate dehydrogenase (IDH2; 1 µg/mL);
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MDH 1 µg/mL, KGDH 0.25 µg/mL and aconitase 0.5 µg/mL, with expression
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normalized to that of mitochondrial porin. Membranes were then washed and
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incubated with secondary antibody (goat α-mouse/HRP or α-rabbit/HRP; 1:2000
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dilution; Biorad, CA, USA) for 1 hour at RT. Detection was made to film by ECL
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chemiluminescent reagent (Packard Biosciences, CT, USA). Band densities were
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quantified using Quantity One software (Bio-Rad, Australia).
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Oxidative Markers and Antioxidant Capacity
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Protein carbonylation was measured in tissue extracts using the Oxyblot Protein
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Oxidation Detection kit (Chemicon, Australia) according to the manufacturer’s
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protocol. Quantification of protein nitration was made using an anti-nitrotyrosine
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competitive ELISA kit (Upstate Chemicals) using nitrated BSA as standards. Levels of
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HNE-protein adducts were determined using an ‘in house’ ELISA assay using HNE-
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modified BSA as standards, essentially as described by Benderdour (4). Total
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antioxidant capacity was measured based on the ability of endogenous antioxidants
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to prevent the oxidation of ABTS (2,2’-Azino-di-[3-ethylbenzthiazoline sulphonate])
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by metmyoglobin (Cayman Chemicals, MI, USA). Total glutathione (GSHt) was
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quantified using the GSH cycling assay, based on the reduction of 5,5’-dithiobis(2-
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nitrobenzoic acid) (DTNB) relative to the oxidation of NADPH (GSH-GSSG-412 kit,
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Oxis Research, OR, USA). For GSHt measures, homogenates were prepared from
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frozen tissues in sodium phosphate buffer, pH 7.4, while for oxidized glutathione
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(GSSG), a parallel set of tissue samples was processed with the addition of 3 mM 1-
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methyl-2-vinylpyridinium trifluoromethanesulfonate (M2VP) to rapidly scavenge
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reduced GSH present, thus preventing its participation in the GSH cycling reaction.
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Homogenates were incubated with an equal volume of ice-cold 5% metaphosphoric
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acid (MPA) to precipitate protein, centrifuged at 10,000 g for 2 minutes and
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neutralized with 5 M NaOH, before being used for the GSH assay as directed.
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Reaction rates were compared to those of supplied standards and expressed as nmol
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GSH/mg protein or pmol GSSG/mg protein). Reduced GSH was calculated as [GSHt]
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minus (2 x [GSSG]). Coenzyme Q10 (CoQ10), which is almost exclusively
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mitochondrial, was quantified by solvent extraction using hexanes/ethanol and
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analyzed using HPLC (42) . Activity of aldose reductase (AR) was measured in tissue
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extracts essentially as described by Srivastava et al(50). All antioxidant measures
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were normalized to tissue protein content, determined using the BCA assay (Sigma
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Aldrich).
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Immunocapture of Electron Transport Complex Proteins
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Purified complex I, IV and F1FoATP synthase protein was isolated from LV tissue
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extracts using monoclonal antibody-based immunocapture protocols (Mitosciences,
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OR, USA). Briefly, 1 mg tissue protein was solubilized with 10% dodecyl-β-D-
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maltoside on ice for 30 minutes and centrifuged at 16,000g for 10 minutes at 4°C.
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The solubilized protein (supernatant) was incubated with 10 µl bead matrix in PBS
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(cross-linked to 25 µg monoclonal antibody) for 3 hours at room temperature with
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continuous rotation. Excess sample was washed from the beads with 3 consecutive
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washes in 100 volumes of 50 mM Tris-Cl, pH 7.4, followed by gentle centrifugation at
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1000g for 1 minute. Purified complex protein was eluted from the beads with 20
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mM glycine, pH 2.5, neutralized with 1.5 M Tris-Cl, pH 8.8 and stored at −80°C until
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required. Protein content was determined using the BCA assay (Sigma Aldrich). For
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detection of subunits subject to oxidative modification, 10 µg of purified protein per
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lane was run under standard western blotting conditions and immunoblotted using
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antibodies to DNP (carbonylation; Oxyblot Protein Oxidation kit; Chemicon
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International, USA), HNE (monoclonal; Oxis Research) and nitrotyrosine (monoclonal;
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Upstate Chemicals, NY, USA).
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Statistical Analysis
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All results are presented as mean ± SEM. For each measure, group contrasts were
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performed between the non-failing and heart failure groups using the Student’s t-
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test, with significance accepted at p<0.05. For end-stage HF samples, no significant
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differences were noted between values determined from any of the assays using
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tissues from ischemic heart disease and dilated cardiomyopathy patients, thus these
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were combined in the HF group and compared to NF.
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Results
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Electron Transport Chain Activities
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Figure 1 demonstrates a reduction in the activities of complexes I and IV in HF
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compared to NF. While activity of complex I was 29% lower in the failing group
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(p=0.007; Figure 1A), the decline in activity was most pronounced in complex IV, with
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a 38.7% reduction in activity in the failing group compared to non-failing controls
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(p=0.0025; Figure 1D). Activities of complexes II or III were not significantly different
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between the NF and HF groups (Figures 1B and IC). Activity of citrate synthase, a
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marker of mitochondrial content, was similar between the two groups (p=0.4859;
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Figure IE). Activity of SERCA2, a biochemical marker of heart failure, was reduced by
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28.4% in the heart failure group (p=0.037; Figure 1F), which was concurrent with a
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significant up-regulation in atrial natriuretic peptide (ANP) and β-myosin heavy chain
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(MHC7) gene expression in these identical samples (data not shown).
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Mitochondrial OXPHOS Proteins
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As shown in Figure 2, there was no change in protein expression of complexes I-IV or
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F1FoATP synthase between NF and HF tissues (p=NS; Figures 2A-2E). Samples were
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normalized to outer membrane porin content, which was found to be similar
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between the two groups (p=0.9094; Figure 2H). To account for potential changes to
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the mitochondrial genome which may affect protein expression, COX I protein
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expression was also measured. As shown in Figure 2G there was no difference in
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COX I expression between the non-failing and failing groups. The adenine nucleotide
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translocase (ANT), which forms a supercomplex with F1FoATP synthase and the
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phosphate carrier, was also unchanged in protein expression (Figure 2F). These
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results indicate that protein content of both cytosolic and mitochondrial-encoded
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subunits does not change in heart failure, which supports the findings of Scheubel et
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al (44) who found no change in the gene expression of any of the mitochondrial-
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encoded subunits in the failing human heart.
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Krebs Cycle Enzymes
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Due to the association between NADH-linked Krebs cycle enzymes and complex I, we
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explored whether Krebs cycle enzymes may also be affected in heart failure, which
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has yet to be determined. As seen in Figure 3, activity of two of the three NADH-
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linked Krebs cycle enzymes was significantly lower in the failing heart. Loss of
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activity was the most pronounced in ICDH (NADH), being 43% lower in the failing
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group (p=0.0003; Figure 3A), despite unchanged protein expression (Figure 4A),
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while a 25% reduction was seen in MDH activity (p=0.0090; Figure 3B). KGDH
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activity, however, was not significantly different between the two groups (p=0.8705;
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Figure 3C), although slightly greater protein expression was noted (Figure 4C). While
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total NAD was similar between the two groups, there was a significant decline in the
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proportion of reduced NADH in the failing tissue, with a concurrent increase in
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oxidized NAD (p<0.001; Figure 3D). A significant correlation existed between ICDH
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activity and reduced [NADH] (r2=0.382, p=0.0082; Figure 3E), which was not present
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between MDH activity and [NADH] (r2=0.0009, p=0.8788). As aconitase activity is
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highly vulnerable to oxidative stress (14), we measured this finding a 46% reduction
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in activity in the HF vs NF group (p=0.0167; Figure 3F), in spite of an elevation in
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protein expression (Figure 4D). Together, these results identify key Krebs enzymes,
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in particular those associated with complex I to be functionally vulnerable to
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oxidative stress.
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Oxidative Stress and Antioxidant Measures
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Increased protein carbonylation has been described in a number of human
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pathologies and is considered a broad marker of oxidative protein damage (12). In
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the HF group, there was a 31% increase in protein carbonyls (p=0.0288; Figure 5A).
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In comparison, levels of nitrated or HNE-modified proteins did not differ between
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the two groups (Figures 5B and 5C). While total antioxidant capacity did not differ
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between the two groups (p=0.259; Figure 5D), there was a significant loss of GSH and
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CoQ10 in the heart failure tissues (p=0.022; Figure 5E and p=0.047; Figure 5F
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respectively). Activity of aldose reductase, which contributes to the clearance of
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HNE, was not significantly different between the two groups. Thus in the failing
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human heart, in the face of diminished antioxidant and REDOX capacity (CoQ10 and
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GSH), protein oxidation accumulates.
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Oxidation of OXPHOS Protein Subunits
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We examined whether oxidative modification of OXPHOS protein subunits was
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evident in HF. In complex I, four subunits were subject to both carbonyl modification
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and protein nitration, being the 75 kDa, 51 kDa, 49 kDa and 24 kDa subunits, while
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the 39 kDa subunit was positive to carbonyl modification alone (Figures 6A and 6B).
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Notably, the first four of these possess Fe-S clusters. Complex IV contained two
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subunits which were subject to both carbonyl modification and protein nitration,
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being the COX I (57 kDa) and COX II (26 kDa) subunits (Figures 6C and D). These
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subunits contain the copper and heme centers and are directly involved in the
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transfer of electrons from cytochrome c to molecular oxygen. F1FoATP synthase
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contained two subunits which were modified by both carbonyl groups and protein
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nitration; the F1β subunit (52 kDa) and the F0 beta subunit (24.7 kDa) (Figures 6F and
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6G). An additional band was subject to carbonyl modification; tentatively identified
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as the F1γ subunit (30.1 kDa). However, as the ANT co-captures with F1FoATP
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synthase (product sheet), it is possible that this band may also be the ANT protein
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(33 kDa). These results show that OXPHOS proteins containing redox centers are
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predominantly the targets of oxidative modification.
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Discussion
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HF features a severe imbalance between energy demand and supply, with availability
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of ATP up to 30-50% lower in human failing hearts (3, 33, 52). Coupled with
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oxidative modification of contractile myofibrillar proteins (8) the failing human heart
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is progressively unable to sustain cardiac output requirements. Studies in skinned
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muscle fibre bundles from both human and animal failing LV report lower state III
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respiration (ADP-coupled synthesis) denoting impacted mitochondrial oxidative
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phosphorylation (OXPHOS) (47). Lowered activities of individual electron transport
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chain (ETC) enzymes, namely complexes I, III and IV, have been previously reported
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(1, 2, 7, 17, 22, 40, 44). However, it has not been established whether this
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represents widespread concurrent mitochondrial perturbation in the failing heart, or
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is limited to specific target sites. Thus the aim of the present study was to
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characterize where specific enzyme dysfunction occurs concurrently in the
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mitochondrial Krebs and oxidative phosphorylation pathways in the end-stage failing
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human heart.
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The mitochondrial electron transport chain (ETC) forms a major regulatory site for
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mitochondrial respiration, harnessing energy released from oxidative
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phosphorylation (OXPHOS) to drive the synthesis of ATP. To date, there have been
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numerous studies reporting defects in individual OXPHOS enzymes in the failing
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human heart with complexes I (1, 2, 22, 44) , III (7, 17) and IV (1, 2, 7, 40) being the
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predominant targets. While these studies highlighted functional loss of activity in
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individual enzymes, in skinned muscle fibre bundles, Lemieux (22) and colleagues
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further demonstrated defective complex I-driven coupled respiration not only in the
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end-stage failing heart, but in the early stages of heart disease, suggesting
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mitochondrial dysfunction as a primary cause leading to metabolic insufficiency,
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rather than a secondary event. In this study we report a 29% and 38% respective
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decline in the activities of complexes I and IV in the failing heart (Figure 1A and 1D),
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supporting previous studies in human tissues (1, 40, 44), while activities of
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complexes II, III and the mitochondrial marker, citrate synthase, were not
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significantly different between the two groups.
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While some of our findings support those of previously reported studies above,
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some variation between other published human studies, see review by Lemieux and
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Hoppel (21), may relate to the limitations of studies that include: fewer subjects than
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our current study, measures of only a few select targets (and not a wider concurrent
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series of enzymes), variable donor etiology and history, variable donor heart storage
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during transport, variable HF etiology and disease progression. Unlike animal
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experimental studies where control and treatment groups have strictly controlled
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conditions (and are imposed on healthy hearts) human tissues studies have distinct
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limitations. Notably variation in prior donor heart history, donor cause of death,
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variable progression of brain death, collection and storage methods, and timing are
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expected to contribute to variability of measured endpoints. In a study that
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compared potential donor heart biopsies taken from ICU patients versus accident
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victims, a marked loss of ATP was evident in tissue from ICU patients (33), likely due
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to differing prior chronic illness state, ischemic and tissue retrieval times. This is also
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demonstrated in experimental studies of donor storage, for example canine hearts
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stored in UW solution resulted in a 30% loss of ATP after 12 hours (46). Although we
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found no difference in citrate synthase activity between NF and HF, two studies
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reported moderate reductions in citrate synthase activity in heart failure (18, 22),
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However, they did not adjust CS activity per gram of protein but rather per gram of
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wet tissue weight, thus subject to confounding from differences in edema that may
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be influenced by crystalloid buffer donor heart storage. However, despite such
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limitations of human heart samples, as opposed to prolonged and chronic cardiac
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remodelling and subsequent maladaptive failure, in the absence of a true
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experimental ‘control’ these ‘non-failing’ donor tissues are valuable surrogates for
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concurrent comparisons having suffered relatively acute stresses of brain death and
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subsequent collection. Ultimately experimental animal models do not fully account
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for the reality of clinical conditions. Thus the importance and novelty of the present
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study is that the present series of measures are concurrent (with equivalent
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conditions) across each group for protein activities and expression of OXPHOS and
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Krebs enzymes.
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Does lowered activity reflect a decline in protein expression due to potential
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alterations in mitochondrial content or dysfunction due to post-translational
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modification of protein functional sites? While decreases in mitochondrial DNA
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replication, mitochondrial biogenesis regulators, citrate synthase and increases in
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mitochondrial DNA oxidation have been reported in the failing human heart (19),
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studies by Scheubel (44) and Bornstein (6) demonstrated no change in mtDNA copy
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number or gene expression of any of the mitochondrial-encoded ETC protein
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subunits in failing tissues. In addition, a limited number of epidemiological studies
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have found only point mutations or deletions in the ETC genes in a small subset of
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patients (2, 17). While reduced protein expression of mtDNA-encoded ND1, ND6
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(NADH dehydrogenase subunits 1 and 6) and cytochrome b protein, but not nDNA-
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encoded SDHA (complex II) have been previously measured (19), studies of actual
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protein expression in OXPHOS and Krebs cycle enzymes, which are modified by
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factors other than DNA content, have been limited. To address this question,
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protein expression of representative subunits from complexes I-IV, F1FoATP synthase,
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ANT and the mtDNA-encoded COX I subunit were measured by individual western
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blots and normalized to the mitochondrial outer membrane protein, VDAC-1 (porin).
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Notably, there was no significant difference in porin expression between the two
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groups, nor any difference in protein expression of subunits from complexes I-IV,
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F1FoATP synthase, ANT or COX I when normalized to porin (Figure 2A-H). These
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results therefore imply post-translational disturbance of enzymatic function akin to
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ischemia-reperfusion injury, whereby function is lost despite protein levels being
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maintained (38).
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Krebs Cycle Enzymes
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Although studies on human heart failure have predominantly focused on OXPHOS
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enzymes, little is known about whether other mitochondrial enzymes, in particular
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the Krebs cycle enzymes, are affected in heart failure. Krebs cycle enzymes provide
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reducing equivalents (NADH and FADH2) to the electron transport chain and are key
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regulators of mitochondrial oxidative metabolism. It has also been reported that
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KGDH and PDH can generate superoxide and hydrogen peroxide under normal
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conditions (51). Given the link between complex I and ROS-induced injury, we
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extended our study to determine whether complex I (NADH) – linked Krebs cycle
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enzymes were altered in the failing heart. As shown in Figure 3, activity of NADH-
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linked enzymes isocitrate dehydrogenase (ICDH) and malate dehydrogenase (MDH)
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was significantly lower, despite unchanged protein levels (Figures 4A and 4B), while
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activities of Krebs cycle enzymes not associated with complex I (complex II /succinate
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dehydrogenase; citrate synthase) were not different between the two groups. The
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association between oxidative stress and perturbation of enzyme function in HF was
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further confirmed by a 46% decline in the activity of aconitase (Figure 3F), a well-
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established marker of oxidative injury due to its iron-sulphur centres (14).
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It has been previously demonstrated that NADH directly influences the rate of
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mitochondrial state III (ADP-linked) respiration, while such a correlation did not exist
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between complex I activity and the state III respiratory rate (25). In addition, critical
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loss of NAD induces morphological changes to mitochondrial membranes and
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cellular death via the necrotic death pathway (20). While total NAD levels were
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unchanged between the two groups, there was a significant decline in levels of
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reduced NADH in the failing group, with concomitant increases in oxidized NAD and
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a lower NADH:NAD ratio (Figure 3D). [NADH] correlated well with ICDH activity
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(Figure 3E), which highlights its role in the regulation of mitochondrial respiration,
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though there is no direct correlation between MDH and [NADH]. Studies of creatine
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kinase (CK) in human heart failure suggest a multiplicative effect of both loss of CK
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activity and a reduction in the creatine pool on ATP synthesis in the failing heart(33).
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Thus it also follows that both reductions in activities of Krebs cycle enzymes
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combined with reduced substrate availability would have an additive effect in
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contributing to lowered ATP synthesis rates. The susceptibility of ICDH to oxidative
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damage has also been demonstrated in a spontaneously hypertensive (SHHF) rat
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model of heart failure, by which loss of activity due to oxidative damage was present
404
early in failure before many of the classical markers of heart failure were present (4).
405
The above results therefore highlight that select mitochondrial enzymes are targeted
406
in heart failure, namely those associated with sites of ROS injury, such as complexes I
407
and IV, complex I-linked Krebs cycle enzymes and aconitase.
408
409
Post-Translational Oxidative Modification of Electron Transport Chain Enzymes
410
The association between increased ROS products and contractile dysfunction in
411
human heart failure has been well documented. Raised 4-hydroxynonenal (HNE) and
412
malondialdehyde (MDA) levels have been reported in the serum (27, 30) and tissues
413
(32, 43) of heart failure patients, while increased inflammatory isoprostane levels
414
have been reported in the pericardial fluid of patients undergoing coronary bypass
415
(28). In this study we extended these findings, demonstrating increased tissue
416
protein oxidation (carbonyl modification) (Figure 5A) in the failing myocardium,
417
concomitant with a moderate lowering of glutathione and coenzyme Q10 levels
418
(Figures 5E & 5F).
419
420
A major source of cellular ROS arises from the mitochondrial respiratory chain, often
421
termed ‘electron leak’, originating predominantly from complexes I and III (54, 55). It
422
is these proteins which are more commonly the subject of oxidative attack, such as
423
occurs in ischemia-reperfusion injury (36, 39). Complex I facilitates the first step of
424
electron transport, transferring electrons from NADH to a non-covalently bound
425
flavin mononucleotide (FMN), through a series of iron-sulphur clusters to the
426
terminal acceptor, ubiquinone. The precise origin of ‘electron leak’ within complex I
427
remains controversial, although the FMN, iron-sulphur (Fe-S) clusters and
428
ubiquinone have been proposed as potential sites (23, 31).
429
430
In isolated complex I from human heart mitochondria, we revealed several subunits
431
which were specific targets of oxidative modification. These included the 75 kDa, 51
432
kDa, 49 kDa, 39 kDa and 24 kDa subunits (Figures 6A and 6B). Notably, all the
433
subunits subject to carbonyl and nitrotyrosine modification contain iron-sulphur
434
clusters (10). While the 39 kDa subunit does not possess an Fe-S centre,
435
modification of the subunit by HNE was found to occur under basal conditions in
436
isolated bovine sub-mitochondrial particles (11). Given that almost all of the
437
subunits modified by carbonyl and nitrotyrosine residues contain Fe-S centres, it
438
therefore appears likely that the source of ROS within complex I resides proximal to
439
the Fe-S clusters.
440
441
In turn, two distinct protein subunits of complex IV displayed modification to all
442
three markers of oxidative damage: carbonyl (DNP), nitrotyrosine and HNE binding
443
(Figure 6C, 6D and 6E). These were identified as subunits I and II, of which the
444
findings are consistent with the functional role of these subunits. The four redox
445
centres of complex IV, two copper and two heme, are located within subunits I and II
446
and are involved in the transfer of electrons from cytochrome c to molecular oxygen.
447
Thus these subunits, can be considered potential sites of electron ‘leak’. Structural
448
analysis of the complex IV protein in eukaryotes has indicated that the heme and
449
copper centres also contain a high proportion of histidine and cysteine residues (9),
450
which are known targets of oxidative attack.
451
452
The results of our study indicate that the β (F1), b (Fo) and γ (F1) subunits from ATP
453
synthase isolated from failing human hearts are particularly susceptible to oxidative
454
attack, containing both carbonyl and nitrotyrosine adducts (Figure 6F and 6G).
455
These findings are consistent with those described by Choksi (11) in isolated bovine
456
heart mitochondria, who demonstrated that under basal respiratory conditions the β
457
subunit of F1FoATP synthase is subject to oxidative modification by carbonyl, HNE
458
and nitrotyrosine adducts. In comparison, the Capaldi group analysed the entire
459
human mitochondrial proteome for the presence of N-formylkynurenine, a product
460
of dioxidation of tryptophan residues using MALDI-TOF mass spectroscopy. In their
461
findings, 51 peptides from 39 mitochondrial proteins were found to contain N-
462
formylkynurenine in their tryptic fragments, including nine subunits of complex I, the
463
most oxidation being in the 39 kDa and 51 kDa subunits, and the d, ATPase 6 and g
464
subunits of F1FoATP synthase (53). In a murine model of cardiomyopathy due to
465
Trypanosoma cruzi infection, increased mitochondrial oxidative stress, correlating
466
with loss of complex I and III activity and increased lipid peroxidation (TBARS) has
467
been described. Analysis of oxidatively modified subunits (carbonylation) in this
468
model by BN-PAGE and immunoblotting revealed catalytic sites of the respiratory
469
chain components were most susceptible to oxidative modification, including the
470
catalytic core and Fe-S containing subunits of complex I, in addition to five subunits
471
of complex III, subunit I of complex IV and the α, β and γ subunits (F1 portion) of
472
F1FoATP synthase (56). A limitation of our study is that we were unable to utilize
473
mass spectroscopy to confirm the identity of modified proteins due to limited
474
additional tissue availability to per patient and the relatively low abundance of these
475
proteins.
476
477
Analysis of complexes I and IV revealed that subunits associated with redox centres
478
were increased targets of oxidative modification. Given that no redox centres are
479
present in F1FoATP synthase, the question then arises as to why certain subunits, in
480
particular the β subunit, are oxidatively targeted. One possibility relates to the
481
‘supercomplex’ arrangement of membrane-bound proteins within the mitochondrial
482
inner membrane, which provides structural stability and maximal function of ETC
483
complexes. Supercomplex formation is facilitated through a close protein-lipid
484
interaction between membrane-bound proteins and inner membrane lipids such as
485
cardiolipin (CL), which is crucial for optimal enzyme function, in particular that of
486
complexes I, III and IV (13, 37). Reductions in the proportion of membrane
487
cardiolipin, which is particularly susceptible to oxidative attack due to its highly
488
unsaturated bonds, have been previously measured in hearts from patients with
489
heart failure, together with a shift away from the L4CL-tetralinoleoyl-CL formation
490
(48). Increased dissociation of ETC complexes from supercomplex formation to free
491
enzyme complexes has been demonstrated in a canine model of heart failure (41),
492
leading to lowered state III respiratory rates (and thus less ATP synthesis) despite
493
membrane integrity remaining intact with unchanged activities and protein content
494
of complexes I to IV. The diminished activity state of F1FoATP synthase and thus
495
lower availability of ATP is observed with decreased SERCA activity (see Figure 1F).
496
497
Despite convincing evidence of elevated ROS products and their correlation with the
498
progression of disease, the importance of oxidative stress in the pathology of heart
499
failure, in particular the decline in mitochondrial oxidative phosphorylation, has not
500
been fully recognized due to limited capacity to demonstrate direct cause. A direct
501
causal link between increased mitochondrial-derived oxidative stress and the
502
initiation of heart failure was established by Nojiri et al (35), who used a heart
503
specific MnSOD mouse knockout model to show increased cardiac enlargement,
504
depressed contractile function, diminished heart ATP content, lowered activities of
505
complexes I-III and II-III and increased levels of superoxide and lipid peroxides (MDA)
506
compared to controls. Elevated ROS levels have also been demonstrated in
507
cardiomyopathy due to a primary mitochondrial disorder (45). The evidence
508
presented in our study suggests that targeted oxidative modification of
509
mitochondrial proteins occurs in end-stage heart failure with ischemic and dilated
510
cardiomyopathy etiology, impacting enzymatic function and ultimately ATP supply,
511
causing a decline in contractile performance.
512
513
Conclusion
514
In the present study we have for the first time examined key left ventricle myocardial
515
Krebs cycle enzymes concurrently with OXPHOS enzymes in end-stage human heart
516
failure. Elevated oxidative stress in the failing heart results in specific, targeted
517
oxidative modification of mitochondrial OXPHOS enzymes (complexes I, IV)
518
associated with sites of superoxide production, leading to loss of enzymatic function
519
without alteration of protein levels. Together with dysfunctional Krebs enzymes
520
(ICDH, MDH, aconitase), altered mitochondrial membrane lipid environment,
521
supercomplex organization, reduced oxidative phosphorylation and ATP synthesis,
522
these changes confer a multi-component impact on energy depletion and exacerbate
523
contractile dysfunction in the failing human heart.
524
525
526
527
528
529
Disclosures
No conflicts of interest, financial or other, are declared by the authors.
Acknowledgements
530
FS was supported by the National Health and Medical Research Council of Australia
531
(NHMRC Dora Lush Postgraduate Scholar) and an Early Career Post-Doctoral
532
Fellowship (GNT1016543). The work was supported in part by NHMRC Project
533
funding (SP), and the Victorian Government’s Operational Infrastructure Support
534
Program to the Murdoch Childrens Research Institute.
535
536
537
538
539
540
541
542
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Figure Legends
Figure 1. Enzyme activities of mitochondrial OXPHOS enzymes. Activities of
complexes I-IV, citrate synthase and SERCA2 was measured in 5-100 μg tissue
protein extract as described. Activities of complexes I, IV and SERCA2 were
significantly lower in the Heart Failure group (complex I: p<0.0007; complex IV:
p=0.001; SERCA2: p=0.037), whereas activities of complexes II, III and citrate
synthase did not significantly differ between the two groups. For all figures, values
equal mean ± SEM. Non-Failing, n=15; Heart Failure, n=30.
Figure 2. Expression of protein subunits from I, IV and F1FoATP synthase. Protein
expression of mitochondrial OXPHOS enzymes was determined using representative
subunits from complexes I-IV, F1FoATP synthase, ANT and VDAC1 (porin). Protein
expression was determined using western blotting, with 1 μg protein loading/well, as
described in the Methods section. Values represent mean density of target protein
normalized to individual porin expression. p=NS all groups. n=18 per group.
Figure 3. Enzyme activities of Krebs cycle enzymes. Enzyme activity of ICDH, MDH,
KGDH and aconitase was measured as described. Values were expressed as
nmol/min/mg protein. Activities of ICDH (p<0.001), MDH (p<0.01) and aconitase
(p<0.05) were significantly lower in the failing group, while activity of KGDH did not
differ between groups. Non-Failing, n=20; Heart Failure, n=25.
Figure 4. Protein expression of Krebs cycle enzymes. Protein expression of ICDH,
MDH, KGDH and aconitase was measured through western blotting, using 5 µg
protein loading/lane. Values represent mean density of the target protein
normalized to individual sample porin expression. While protein expression of
aconitase and KGDH were slightly higher in the failing group (p<0.05), there was no
difference in ICDH and MDH expression between the Non-Failing and Heart Failure
groups (p=NS). Non-Failing, n=20; Heart Failure, n=30.
Figure 5. Antioxidant and oxidative stress measures in the human myocardium.
Levels of oxidative markers (protein carbonylation, HNE and protein nitration) and
antioxidant levels (total antioxidant capacity, glutathione and coenzyme Q10) were
measured in heart homogenates as described in the Methods. While total
myocardial antioxidant capacity was not significantly different between groups,
levels of glutathione and coenzyme Q10 were significantly lower in the failing group
(both p<0.05 control vs HF). Non-Failing, n=20; Heart Failure, n=30.
Figure 6. Oxidatively modified subunits of complexes I, IV and F1FoATP synthase.
Semi-purified protein subunits were prepared from Non-Failing and Heart Failure LV
tissues using immunocapture and probed for antibodies to carbonyl (DNP)
modification, nitrated protein and HNE protein using western blotting. Blots
represent protein subunits specifically targeted by oxidative modification. NonFailing, n=3; Heart Failure, n=5.
Complex I Activity
80.0
Activity (nmol/min/mg protein)
70.0
60.0
***
50.0
40.0
30.0
20.0
10.0
0.0
Non-Failing
Heart Failure
Patient Group
'JHVSFB
Complex II Activity
Activity (nmol/min/mg protein)
35.0
30.0
25.0
20.0
15.0
10.0
5.0
0.0
Non-Failing
Heart Failure
Patient Group
'JHVSFC
Complex III Activity
Activity (nmol/min/mg protein)
400.0
350.0
300.0
250.0
200.0
150.0
100.0
50.0
0.0
Non-Failing
Heart Failure
Patient Group
'JHVSFD
Complex IV Activity
Activity (nmol/min/mg protein)
700.0
600.0
500.0
**
400.0
300.0
200.0
100.0
0.0
Non-Failing
Heart Failure
Patient Group
'JHVSFE
Citrate Synthase
Activity (μmol/min/mg protein)
2.25
2.00
1.75
1.50
1.25
1.00
0.75
0.50
0.25
0.00
Non-Failing
Heart Failure
Patient Group
'JHVSFF
SERCA2 Activity
Activity (nmol/min/mg protein)
140
120
100
*
80
60
40
20
0
Non-Failing
Heart Failure
Patient Group
'JHVSFG
Protein Expression (CI/Porin)
1.40
Complex I (39 kDa subunit)
1.20
1.00
0.80
0.60
0.40
0.20
0.00
Non-Failing
'JHVSFB
Heart Failure
Complex II (70 kDa subunit)
Protein Expression (CII/Porin)
1.40
1.20
1.00
0.80
0.60
0.40
0.20
0.00
Non-Failing
'JHVSFC
Heart Failure
Complex III (Core I)
Protein Expression (CIII/Porin)
1.20
1.00
0.80
0.60
0.40
0.20
0.00
Non-Failing
'JHVSFD
Heart Failure
Complex IV (Vib subunit)
Protein Expression (CIV/Porin)
1.40
1.20
1.00
0.80
0.60
0.40
0.20
0.00
Non-Failing
'JHVSFE
Heart Failure
))R$73V\QWKDVH (E subunit)
Protein Expression (CV/Porin)
1.40
1.20
1.00
0.80
0.60
0.40
0.20
0.00
Non-Failing
'JHVSFF
Heart Failure
ANT-1 Expression
Protein Expression (ANT/Porin)
1.20
1.00
0.80
0.60
0.40
0.20
0.00
Non-Failing
'JHVSFG
Heart Failure
COX I Protein Expression
Protein Expression (COX I/Porin)
2.00
1.80
1.60
1.40
1.20
1.00
0.80
0.60
0.40
0.20
0.00
Non-Failing
'JHVSFH
Heart Failure
Porin Protein Expression
Protein Expression (Mean Density)
700
600
500
400
300
200
100
0
Non-Failing
'JHVSFI
Heart Failure
Isocitrate Dehydrogenase
Activity (μmol/min/mg protein)
6.0
5.0
4.0
***
3.0
2.0
1.0
0.0
Non-Failing
'JHVSFB
Heart Failure
Malate Dehydrogenase
Activity (nmol/min/mg protein)
60.0
50.0
**
40.0
30.0
20.0
10.0
0.0
Non-Failing
'JHVSFC
Heart Failure
D-Ketoglutarate Dehydrogenase
Activity (nmol/min/mg protein)
35.0
30.0
25.0
20.0
15.0
10.0
5.0
0.0
Non-Failing
'JHVSFD
Heart Failure
Pyridine Nucleotides
7
Non-Failing
Heart Failure
NAD (nmol/mg protein)
6
*
5
4
3
*
2
1
0
Total NAD
NADH only
NAD only
Pyridine Nucleotides
'JHVSFE
Correlation Between
ICDH Activity and NADH
7.0
NADH (nmol/mg protein)
6.0
5.0
4.0
R2 = 0.3822
3.0
p = 0.0082**
2.0
1.0
0.0
0
1
2
3
4
5
6
ICDH Activity (μmol/min/mg protein)
'JHVSFF
7
Aconitase Activity
Activity (nmol/min/mg protein)
1.80
1.60
1.40
1.20
1.00
*
0.80
0.60
0.40
0.20
0.00
Non-Failing
'JHVSFG
Heart Failure
IDH2 Protein Expression
1.80
Density Ratio (IDH2/Porin)
1.60
1.40
1.20
1.00
0.80
0.60
0.40
0.20
0.00
Non-Failing
'JHVSFB
Heart Failure
MDH Protein Expression
0.70
Density Ratio (MDH/Porin)
0.60
0.50
0.40
0.30
0.20
0.10
0.00
Non-Failing
'JHVSFC
Heart Failure
KGDH Protein Expression
*
0.80
Density Ratio (KGDH/Porin)
0.70
0.60
0.50
0.40
0.30
0.20
0.10
0.00
Non-Failing
'JHVSFD
Heart Failure
Aconitase Protein Expression
Density Ratio (Aconitase/Porin)
2.50
*
2.00
1.50
1.00
0.50
0.00
Non-Failing
'JHVSFE
Heart Failure
Protein Carbonylation
*
Mean Density (normalized to protein)
25000
20000
15000
10000
5000
0
Non-Failing
'JHVSFB
Heart Failure
Nitrated Protein
Nitrated Protein
(ng nitrated protein/mg total)
300
250
200
150
100
50
0
Non-Failing
'JHVSFC
Heart Failure
HNE-Protein
HNE (nmol HNE/mg total protein)
3.00
2.50
2.00
1.50
1.00
0.50
0.00
Non-Failing
'JHVSFD
Heart Failure
Total Antioxidant (nmol/mg protein)
Antioxidant Capacity
55
50
45
40
35
30
25
20
15
10
5
0
Non-Failing
'JHVSFE
Heart Failure
Glutathione
140.0
*
Non-Failing
Heart Failure
GSSG (pmol/mg protein)
GSH (nmol/mg protein)
120.0
100.0
80.0
60.0
*
*
40.0
20.0
0.0
Total
'JHVSFF
Reduced
Oxidized
(GSSG)
Coenzyme Q10
350.0
CoQ10 (umol/mg protein)
300.0
*
250.0
200.0
150.0
100.0
50.0
0.0
Non-Failing
'JHVSFG
Heart Failure
170
109
79
60
47
35
25
Non-Failing
Heart Failure
Carbonyl Modification of Mitochondrial
Complex I Protein.
'JHVSFB
170
109
79
60
47
35
25
18
kDa
Non-Failing
Heart Failure
Nitrotyrosine modification of Isolated
Complex I protein.
'JHVSFC
170
109
79
60
47
35
25
18
MW
(kDa)
Non-Failing
Heart Failure
Carbonyl modification of isolated Complex IV
protein.
'JHVSFD
MW
kDa
170
109
79
60
47
35
25
18
Non-Failing
Heart Failure
Nitrotyrosine modification of isolated
Complex IV protein.
'JHVSFE
MW
(kDa)
170
109
79
60
47
35
25
18
Non-Failing
Heart
Failure
HNE modification of isolated
Complex IV protein.
'JHVSFF
MW
(kDa)
109
79
60
47
35
25
18
Non-Failing
Heart Failure
Carbonyl Modification of Mitochondrial
Complex V Protein.
'JHVSFG
MW
(kDa)
109
79
60
47
35
25
18
Non-Failing
Heart Failure
Nitrotyrosine modification of isolated
Complex V protein.
'JHVSFH