Oxidative modification of mitochondrial respiratory complexes in response to the stress of Trypanosoma cruzi infection

Free Radical Biology & Medicine, Vol. 37, No. 12, pp. 2072–2081, 2004 Copyright D 2004 Elsevier Inc. Printed in the USA. All rights reserved 0891-5849/$-see front matter

doi:10.1016/j.freeradbiomed.2004.09.011

Original Contribution OXIDATIVE MODIFICATION OF MITOCHONDRIAL RESPIRATORY COMPLEXES IN RESPONSE TO THE STRESS OF Trypanosoma cruzi INFECTION JIAN-JUN WEN* and NISHA GARG*,y,z *Department of Microbiology and Immunology, University of Texas Medical Branch, Galveston, TX, USA; yDepartment of Pathology, Center for Biodefense & Emerging Infectious Diseases, University of Texas Medical Branch, Galveston, TX, USA; and zSealy Center for Vaccine Development, University of Texas Medical Branch, Galveston, TX, USA (Received 11 June 2004; Revised 24 August 2004; Accepted 10 September 2004) Available online 8 October 2004

Abstract — Previously, we have shown deficiencies in the activities of the mitochondrial respiratory complexes and reduced mitochondrial ATP generation capacity in chagasic hearts infected by Trypanosoma cruzi. In this study, we determined whether the oxidative stress that occurs in response to T. cruzi infection contributes to the catalytic impairment of respiratory complexes and to subsequent mitochondrial dysfunction in murine myocardium. Our data show that oxidative injuries, as determined by the levels of lipid peroxides and protein carbonyls, are incurred in cardiac mitochondria as early as 3 days postinfection and persist throughout the infection and disease. The individual components of the respiratory complexes were separated by two-dimensional, blue-native gel electrophoresis, and carbonyl adducts were detected by Western blotting. We observed substantial carbonylation of the specific subunits of mitochondrial respiratory complexes in infected murine hearts. Of note is the oxidative modification of NDUFS1, NDUFS2, and NDUFV1, which form the catalytic core of the CI complex; UQCRC1, UQCRC2, and UQCRQ, the subunits of the core subcomplex, and UQCRH and CYC1, which form the cyt c1 subcomplex of CIII; and a g chain that is essential for ATP synthesis by CV complex. The extent of oxidative modifications of the subunits correlated with the catalytic defects of the respiratory complexes in the infected myocardium. Taken together, our data demonstrate that respiratory complexes are oxidatively damaged in response to the stress of T. cruzi infection. These data also suggest involvement of the specific susceptibility of the protein subunits, and not generalized mitochondrial oxidative damage in respiratory chain impairment of chagasic hearts. D 2004 Elsevier Inc. All rights reserved. Keywords—Trypanosoma cruzi, Chagasic disease, Endogenous mitochondrial oxidative stress, Protein carbonyls, Respiratory complexes, Mitochondrial dysfunction, Free radicals

the cardiac tissues, that progressively leads to heart failure and death [2]. The progressive severity of CCM has been shown to be associated with an increasing order of oxidative damage to lipids and proteins in the heart [3]. An increase in oxidative stress rather than a decline in antioxidant defense mechanisms was suggested to be the major contributory factor in accumulation of oxidative damage in chagasic hearts [3]. Oxidative stress might occur in the course of parasite infection and disease development as a consequence of tissue destruction produced by toxic secretions of the parasite, of immune-mediated cytotoxic reactions [4,5], or of secondary damage to mitochondria [6–8]. Mitochondrial disruption is noted to be one of the major characteristics of cellular abnormalities during chagasic disease devel-

INTRODUCTION

Chagasic cardiomyopathy (CCM) is caused by the protozoan parasite Trypanosoma cruzi, and is widely distributed in the Americas [1]. The disease is characterized by an acute phase, during which parasites invade all tissues and organs and multiply. After the immune system exerts control of parasites, patients enter an indeterminate phase with very low parasitemia and no apparent pathology. Susceptible hosts then enter a chronic phase with increasing tissue damage, mostly in

Address correspondence to: Dr. Nisha Garg, Department of Microbiology and Immunology, University of Texas Medical Branch, 301 University Boulevard, Galveston, TX 77555, USA; Fax: +1 409 747 6869; E-mail: [email protected]. 2072

Mitochondrial oxidative damage during T. cruzi infection

opment. In cardiac biopsies of chagasic patients [9] and experimental models [10,11], swollen mitochondria, which may also be dysfunctional, are shown to accumulate. We have demonstrated, in a murine model of T. cruzi infection and chagasic disease development, a substantial decline in respiratory chain complex activities [12]. The alterations in NADH–ubiquinone oxidoreductase (CI) activity were found to be more pronounced during the acute infection phase, while ubiquinol– cytochrome c reductase (CIII) activity was constitutively repressed throughout the infection and disease phase, and F1F0 ATP synthase (CV) inhibition appeared in chronic hearts [12]. These changes in respiratory chain activity were closely related to cumulative oxidative damage in the myocardium [3]. A number of studies have suggested that alteration in mitochondrial respiratory chain activity is one of the major systems in the myocyte for reactive oxygen species (ROS) generation. CI and CIII are recognized as prime sites for electron leakage to molecular oxygen, resulting in free radical generation in mitochondria [6,13–15]. The rate of mitochondrial free radical production is exponentially increased when the CI or CIII complex of the respiratory chain function at a sub optimal level [14]. The deleterious production of ROS in response to a decrease in CI or CIII activities and the resultant impairment of electron transfer in experimental models of ischemia and heart failure are well documented [6,13,14,16]. Considering the observations of a decline in CI and CIII activities in T. cruzi-infected murine myocardium, it is proposed that chagasic hearts sustain oxidative stress due to mitochondria-generated ROS toxicity [12]. Because the mitochondrial membrane is rich in polyunsaturated fatty acids, lipid peroxidation and its toxic aldehydic end products, i.e., 4-hydroxynonenal (HNE) and malonyldialdehyde (MDA), may also be generated in mitochondria [17]. HNE is highly reactive with protein molecules, causing direct oxidation of arginine, lysine, proline, or threonine residues [18]. The purpose of this study was to examine whether mitochondria are the targets of ROS-induced oxidative modifications in chagasic hearts. We specifically focused on the respiratory chain complexes whose activities are altered during the course of infection and disease development. We employed blue-native polyacrylamide gel electrophoresis (BN-PAGE) to separate the respiratory chain complexes CI–CV, and second-dimension denaturing SDS–PAGE to resolve the individual subunits of these complexes. Immunoblotting analysis was then used to detect oxidatively modified proteins. Carbonylated proteins were subjected to N-terminal sequencing to establish their identity. Our data show sustained oxidative damage to mitochondrial membranes during the

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course of chagasic disease development. Further, we have identified specific protein subunits of the mitochondrial respiratory chain complexes that are susceptible to and targeted for ROS-mediated oxidative modification. We discuss the differential sensitivity of the specific proteins to endogenous oxidative stress and its role in mitochondrial respiratory chain dysfunction in CCM. MATERIALS AND METHODS

Mice and parasites C3H/HeN male mice (Harlan Labs), aged 6 to 7 weeks, were used in the current study. The SylvioX10/4 strain of T. cruzi and C2C12 cells (murine skeletal muscle hybridoma cells) were purchased from American Tissue Culture Collection (ATCC, Rockville, MD, USA). T. cruzi trypomastigotes were maintained and propagated by the continuous in vitro passage of parasites in monolayers of C2C12 cells. Mice were infected by intraperitoneal injection of 25,000 culturederived trypomastigotes. Animal experiments were performed according to the National Institutes of Health Guide for Care and Use of Experimental Animals and approved by the UTMB Animal Care and Use Committee. Mitochondria isolation Tissues were washed and suspended in isolation buffer (5 mM Hepes, pH 7.2 containing 210 mM mannitol, 70 mM sucrose, 1 mM EGTA, and 0.1% fatty acid-free BSA bovine serum albumin; tissue:buffer ratio, 1:10 w/v). Mitochondrial isolation was carried out by published methods [12]. The crude mitochondria were stored in aliquots at 808C for further use. For each analysis, fresh aliquots were thawed, centrifuged once, and resuspended in the isolation buffer. When tissue samples were processed for the estimation of protein carbonyl derivatives (PCOs) and thiobarbituric acidreactive substances (TBARS), h-mercaptoethanol (hME, 2%) and butylated hydroxytoluene (500 AM), respectively, were added to all buffers. Protein concentrations were determined by the Bradford method [19]. Lipid peroxidation We employed the biochemical assay of TBARS to evaluate lipid peroxidation as described [20] with slight modifications [3]. For further confirmation, lipid hydroperoxides (LHPOs) were measured by a ferrous oxidation–xylenol orange (FOX) assay as described [21]. Electrophoresis BN-PAGE was carried out by published methods [12,22]. Briefly, mitochondria were isolated in 20 mM Tris–HCl, pH 7.4, 250 mM sucrose, 2 mM K2EGTA.

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Mitochondria (100 Ag protein) were suspended in extraction buffer (50 mM imidazole pH 7.0, 5 mM aminocaproic acid, 50 mM NaCl; protein:buffer ratio, 1:10), solubilized in 10% dodecyl maltoside (protein:detergent ratio, 1:6). The solubilized proteins were mixed with 5% Coomassie brilliant blue G-250, 0.5 M aminocaproic acid, 10% glycerol (Coomassie:detergent ratio, 1:4), and resolved on a 5–12% native polyacrylamide gradient gel with 4% stacker gel on a Bio-Rad minigel apparatus. Gels were imaged and densitometric analysis of the individual complexes conducted using a FluorChem 8800 image-analyzing system (Alpha Innotech). For resolution of the subunits of the respiratory complexes, gel slices were soaked in 1% SDS and 150 mM h-ME to denature the complexes. Gel slices were slid horizontally on top of the denaturing resolving gel, using hot agarose (0.7% in 0.5% SDS, 15 mM h-ME) as a lubricant and sealant, and protein subunits were separated by SDS–PAGE. We used 10% acrylamide gels on a miniProtean 3 system (Bio-Rad) to resolve the subunits of the CII–CV complexes and 8–12% linear gradient gels on a Protean II XL system (Bio-Rad) to resolve the CI subunits. Gels were fixed for at least 30 min in 7% acetic acid and 10% methanol, and stained in 0.005% Coomassie blue G250 (Bio-Rad). Gels were destained in the same buffer as fixation and imaged as above. HNE treatment and catalytic staining Respiratory complexes from normal murine heart mitochondria were resolved by BN-PAGE and in-gel treated with 256 AM 4-HNE (Cayman) for 3 h at 48C. Catalytic staining reactions to evaluate the activity of the respiratory complexes were performed immediately after the HNE treatment, as described [12]. Quantitative data were estimated as the ratio between the densitometry units obtained for catalytic stained complex and the corresponding Coomassie stained complex  100. Western blot analysis The BN-gel slices containing individual complexes were soaked in 1% SDS to denature the proteins, and the proteins transferred to a PVDF membrane using a TransBlot SD Semi-Dry transfer system (Bio-Rad). Immunoblot analysis was performed as described previously [11]. Mouse monoclonal antibodies raised against the 39-kDa subunit of CI (anti-CI39, 1:5000); 70-kDa subunit of CII (anti-SDH 70, 1:1000); FeS subunit of CIII (anti-ISP, 1:5000); subunit IV of CIV (anti-CIV20, 1:1000); and h subunit of CV (anti-CVh, 1:1000) were from Molecular Probes. Protein carbonyls PCOs were detected as described [23] with slight modifications [3]. For the detection of total PCOs,

mitochondria (20 Ag protein) were denatured and derivatized in 3% SDS, 10 mM 2,4-dinitrophenylhydrazine (DNPH) dissolved in 10% trifluoroacetic acid. After neutralization with an equal volume of 2 M Tris, 30% glycerol, DNP-derivatized protein samples were mixed with an equal volume of 2X sample buffer (4% SDS, 20% glycerol, 4% h-ME, 0.04% bromphenol blue, 120 mM Tris–HCl, pH 6.8), and resolved by SDS–PAGE on 8–10% gradient gels (8 Ag protein/well). Gels were transferred to PVDF membranes. PCOs were probed with rabbit anti-DNP antibody (1:4000 dilution, Sigma), followed by HRP-conjugated goat a rabbit IgG (1:4000 dilution, Sigma), and detected using the SuperSignal chemiluminescent substrate (Pierce, Rockford, IL, USA). Images were visualized, digitized, and quantified by densitometry using a FluorChem 8800 (Alpha Innotech) image analyzing system. For the detection of carbonylation of the respiratory complexes, BN-PAGE gel slices containing individual complex were derivatized with DNPH for 30 min and neutralized, and the subunits resolved by SDS–PAGE as above. Carbonylated proteins were detected by immunoblotting using anti-DNP antibody. The protein carbonylation data presented in Figs. 2–8 are representative of three independent experiments. Protein identification Individual protein bands showing carbonyl modification were excised from second dimension SDS–polyacrylamide gels run simultaneously with the gels that were Western blotted. The proteins were eluted from the gel and subjected to N-terminal sequencing on a 494/HT PROCISE Sequencing system (Applied Biosystems) using the standard Edman sequencing method as recommended by the manufacturer. The services of the Protein Chemistry Laboratory of the Biomolecular Resource Facility, UTMB, were used for protein sequencing. RESULTS

As a model system, we chose C3H/HeN mice infected with the SylvioX10/4 strain of T. cruzi. This mouse– parasite combination has been extensively characterized in our laboratory as a standard model of human CCM [11]. As recognized in humans, the course of disease development in mice is divided into the immediate early phase (3–8 days postinfection, dpi) of parasite infection; acute phase of parasite replication (20–45 dpi); and progressive disease phase (N100 dpi), marked by minimal parasite burden, diffused inflammation, cellular fibrosis, and tissue degeneration in the myocardium [11]. Figure 1 shows the lipid peroxidation of cardiac mitochondria, measured as MDA formation in response

Mitochondrial oxidative damage during T. cruzi infection

Fig. 1. Levels of mitochondrial lipid peroxidation in T. cruzi-infected mice. C3H/HeN mice infected with T. cruzi trypomastigotes were sacrificed at various intervals postinfection. Shown are the MDA and LHPO contents of cardiac mitochondria of infected and control mice. Data are the mean values obtained from three independent experiments, two animals per experiment F SD. P b .05.

to T. cruzi infection and disease development. We found an immediate increase in MDA levels in response to T. cruzi infection that was detectable as early as 3 dpi (28.4 and 22.3 nmol MDA/mg protein in infected and control hearts, respectively, N25% increase). The MDA level in the cardiac mitochondria of infected mice subsided during the early acute phase (21–40 dpi); however, MDA content gradually increased by twofold with progressive severity of chronic disease (105–180 dpi) in infected mice relative to controls ( p b .05). To validate the results of TBARS assay, we measured the triphenylphosphine (TPP)-specific LHPO contents in cardiac mitochondria of infected mice. TPP is a specific reductant of hydroperoxides and allows definitive identification of LHPO [21]. We observed an increase in LHPO levels in the cardiac mitochondria of infected mice parallel to the TBA–MDA elevation with progressive disease development (Fig. 1). Total oxidized and carbonylated proteins of the cardiac mitochondria were visualized by immunoblotting analysis after SDS–PAGE (Fig. 2A). The anti-DNPH signal was less marked in the immediate early phase (3–8 dpi) of infection. The PCO content in cardiac mitochondria began to increase during the acute phase (15–45 dpi) of parasite replication, and remained consistently enhanced throughout the chronic phase of disease progression (N100 dpi) (Fig. 2A, data not shown). A similar level of Coomassie blue staining of cardiac mitochondrial proteins resolved on SDS–polyacrylamide gel confirmed that there was equal loading of the protein samples (Fig. 2B). Previously, we showed that the activities of the respiratory complexes substantially decreased during the course of infection and disease development in chagasic hearts [12]. To probe for a cause–effect relationship between inhibition of complex activities and oxidative stress in mitochondria (Figs. 1, 2), we resolved the mitochondrial complexes from normal

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murine hearts on BN-gel, incubated with 4-HNE (256 AM), and determined respiratory complex activities by catalytic staining. According to the literature, CI is a 980 kDa complex of N42 subunits [24]; CII is a 130 kDa complex of 4 subunits [25]; CIII is a 250 kDa complex of 11 polypeptides, the functional CIII being present as a dimer of molecular mass of 500 kDa [26]; CIV is a 200 kDa complex constituted by 13 components [27]; and CV is a 600 kDa complex of 16 subunits [28]. On the basis of molecular weight, by BN-PAGE, these complexes separate in order of highest to lowest molecular mass: CI, CV, CIII, CIV, and CII (Fig. 3A, lane 1). We confirmed the identity of the complexes by immunoblot analysis using the antibodies specific for complex components (Fig. 3A, lanes 2–6). Catalytic staining of the individual complexes revealed HNE-induced 50, 70, and 55% decreases in CI, CV, and CIV activities, respectively, compared with those in controls (Fig. 3C). Histochemical staining of CIII failed to provide reproducible results. Together, these results show the HNE adducts mediated inactivation of the respiratory complexes and suggest the likely role of oxidative damage in respiratory chain dysfunction in chagasic murine hearts. To identify the specific subunits of the respiratory complexes that may sustain oxidative damage during the course of T. cruzi infection and cardiac disease development and contribute to catalytic inactivation of respiratory chain, we performed Western blot analysis. Mitochondria, isolated from normal and infected murine hearts, were subjected to BN-PAGE to resolve the CI– CV complexes. Individual complexes from the firstdimension gel were then excised, in-gel denatured and

Fig. 2. Protein carbonylation of cardiac mitochondria in T. cruziinfected mice. Infected mice were sacrificed at various intervals postinfection. Carbonylated proteins in the cardiac mitochondria were derivatized with DNPH, and the DNP-reactive proteins identified by Western blot analysis using DNP-specific antibody (A). Coomassie staining of the cardiac mitochondrial proteins (B).

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Fig. 3. (A) Separation of murine heart mitochondria respiratory chain complexes. Cardiac mitochondria were solubilized and the respiratory chain complexes separated on a blue-native, first-dimension gel (lane 1). Immunoblot analysis with complex-specific antibodies confirmed the identity and position of each complex on the blue-native gels. Shown is the immunodetection of CI (lane 2), CV (lane 3), CIII (lane 4), CIV (lane 5), and CII (lane 6) respiratory complexes of normal murine hearts. (B,C). In vitro inactivation of respiratory complexes by oxidative adduct formation. The CI–CV complexes in normal murine heart mitochondria preparation were resolved on BN-gel. BN-gels were incubated with HNE (256 AM) for 3 h. The BN gels were either stained with Coomassie blue (B) or gel slices containing individual complexes were subjected to enzymatic colorimetric reactions (C), as described under Materials and Methods.

derivatized with DNP, and run on the second-dimension denaturing gel. The proteins from the second-dimension gel were transferred to membranes, and DNP-derivatized carbonylated proteins detected by immunoblot analysis with anti-DNP antibody (Figs. 4–8). All polypeptides that are oxidatively modified were identified on the basis of their molecular weight. Among the N42 subunits of CI,

Fig. 4. Identification of oxidatively damaged proteins of CI complex in T. cruzi-infected murine hearts. Myocardial mitochondria from normal and infected mice were solubilized and the respiratory chain complexes separated on a blue-native, first-dimension gel. The gel slice containing the CI complex was excised, and the subunits were separated by second-dimension SDS–PAGE. The oxidatively modified proteins were detected by immunoblotting with anti-DNP antibody (A). Coomassie blue G-250 stain of the membrane used for immunoblot analysis (B). Identification of each numbered band is summarized in Table 1.

several polypeptides were oxidatively modified in the cardiac mitochondria of infected mice (Fig. 4A). We observed oxidative adduct formation with 75, 51/49, and 23/21 kDa CI subunits in both the acute (21 dpi) and chronic (105 dpi) stages of infection and disease development. Carbonylation of the 30 kDa subunit of the CI complex was observed in acute hearts only (Fig. 4A). Among the 11 components of CIII, the ~49/47, 28, 13.5, and 11/9.5 kDa polypeptides were found to be

Fig. 5. Identification of oxidatively modified proteins of the CIII complex in T. cruzi-infected murine hearts. The respiratory chain complexes of murine heart mitochondria were separated by BN gel electrophoresis. The gel slice containing the CIII complex was excised, and the subunits were resolved by second-dimension SDS–PAGE. The oxidatively modified proteins were detected by immunoblotting with anti-DNP antibody (A). Coomassie blue G-250 stain of the membrane used for immunoblot analysis (B).

Mitochondrial oxidative damage during T. cruzi infection

Fig. 6. Identification of oxidatively damaged proteins of the CIV complex in T. cruzi-infected murine hearts. The murine heart respiratory complexes were resolved on BN gels. CIV-containing gel slices were derivatized using DNPH, and protein subunits resolved by SDS–PAGE. Shown are immunoblot analysis with anti-DNP antibody (A) and Coomassie blue G-250 staining of the same membrane after immunoblotting (B).

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in acute hearts (Fig. 6A), carbonylation of CV components was substantially increased during the disease phase (Fig. 7A). The extent of oxidative modification of the mitochondrial proteins and the subunits of the respiratory complexes was not substantially altered in the myocardium of 70 and 154 day old normal mice (controls for 21 and 105 dpi, respectively) (Fig. 8A), thus confirming that the observed oxidative modifications in infected murine hearts are not related to aging. Coomassie blue staining of the membranes used for immunoblot analysis with anti-DNP antibody confirmed equal loading of the protein samples (Figs. 4B–8B). To validate the identity of the oxidatively modified proteins in infected murine hearts, we performed Nterminal protein sequencing by the Edman degradation method (Table 1). The sequencing of the CI subunits identified oxidative modification of the Fe–S protein 2 (NDUFS2, 49 kDa), Fe–S protein 4 (NDUFS4, 23.2 kDa), and h subcomplex subunit 8 (NDUFB8, 21 kDa). The 75 and 51 kDa protein bands submitted for

oxidatively modified in infected murine hearts at all stages of infection and disease development. The extent of carbonylation of 11 and 9.5 kDa subunits of CIII was more pronounced at 21 dpi than at 105 dpi (Fig. 5A). Among the 13 subunits of CIV and 16 subunits of CV, we noted carbonylation mainly of the 50 and 35 kDa polypeptides (Figs. 6, 7). While the extent of oxidative modification of CIV polypeptides was more pronounced

Fig. 7. Identification of oxidatively damaged proteins of the CV complex in T. cruzi-infected murine hearts. The murine heart respiratory complexes were resolved on BN gels. CV-containing gel slices were derivatized using DNPH and protein subunits resolved by SDS–PAGE. Shown are the immunoblot analysis with anti-DNP antibody (A) and Coomassie blue G-250 staining of the same membrane after immunoblotting (B).

Fig. 8. Protein carbonylation of mitochondrial proteins in normal murine hearts is not increased with age. Oxidatively modified mitochondrial (mt) proteins and the subunits of the respiratory complexes (CI, CIII, and CV) in 70 and 154 day old murine hearts were determined by immunoblotting with anti-DNP antibody (A). Coomassie staining of the membranes used for immunoblotting (B).

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Table 1. Carbonylated Subunits of the Respiratory Complexes in Cardiac Mitochondria of T. cruzi-Infected Mice, Identified by N-Terminal Sequencing Molecular mass (kDa)

Protein ID

Z score

NCBI accession No.

Mitochondrial localization

75.0 51.0 49.0

NDUFS1 NDUFV1 NDUFS2

NADH ubiquinone oxidoreductase, Complex I NP_663493 Inner membrane NDa ND NP_598427 Inner membrane 167.7 NP_694704 Inner membrane

23.2

NDUFS4

177.3

NP_035017

Inner membrane

21.0

NDUFB8

186.0

NP_080337

Inner membrane

49.0

UQCRC1

47.0

UQCRC2

28.0

CYC1

11.0

UQCRH

28

P99028

Inner membrane

9.5

UQCRQ

175.4

Q9CQ69

Inner membrane

56.4

MTCO1

169.9

50

ATP5A1 and/or ATPB1

ND

35

ATP5C1

149.9

a

Core protein Core protein Fe-S protein 2, component of the iron–sulfur subcomplex, binds 2Fe–2S and 4Fe–4S iron–sulfur clusters Fe-S protein 4, AQDQ subunit of the iron–sulfur subcomplex, binds 4Fe–4S iron–sulfur cluster h subcomplex subunit 8

Ubiquinol cytochrome c reductase, Complex III 157.0 Q9CZ13 Inner membrane, matrix side ND Q9DB77 Inner membrane, matrix side 159.7 Q9D0M3 Inner membrane, outer side

Cytochrome c oxidase, Complex IV P00397 Inner membrane

F 1 F 0 ATP synthase, Complex V X , AAH62202 Inner membrane Q91VR2

Inner membrane

Function

Formation of complex between cytochrome c and c1 Required for assembly of the CIII complex cytochrome c1 heme-containing redox component, accepts electrons from rieske protein and transfers to cytochrome c Hinge protein, formation of complex between cyt c and c1 Core-associated protein, subunit VII Catalytic subunit 1, forms functional core of CIV with subunits 2 and 3 a and h chains of F1, constitute the catalytically active site of F1 ATP synthase g chain of F1, regulates proton flow and ATP synthesis at F1

ND, could not be sequenced due to N-terminal blockage.

sequencing failed to provide any information due to a blocked N terminus; however, comparison with protein profiles of bovine and human CI suggested that the 75 and 51 kDa bands would most likely encode the NDUFS1 and NDUFV1 core proteins [24]. The sequencing of the CIII subunits identified oxidative modifications of core protein I (UQCRC1, 49 kDa), cytochrome c1 (CYC1, 28 kDa), subunit VII (UQCRQ, core-associated protein, 9.5 kDa), and subunit VIII (UQCRH, hinge protein, 11 kDa). The 47 kDa protein band submitted for sequencing failed to provide any sequence information; however, comparison of the CIII protein profile in our study with those in the literature suggested that it would most likely encode core protein 2 (UQCRQ2, 46.5 kDa) [26]. The CIV ~50 kDa oxidatively modified subunit was identified to be catalytic subunit 1 (MTCO1, 56.4 kDa). One of the CVcarbonylated polypeptides exhibited a substantial homology with the g chain (ATP5C1, 35 kDa) of the F1 ATPase sub complex of F1F0 ATP synthase. The ~50 kDa band of

CV consisted of more than one polypeptide and failed to provide useful sequence information. Following comparison of the CV protein profile in our study with those in the literature, it appeared that the ~50 kDa band would most likely consist of the a or h subunit of the F1 ATPase subcomplex [29]. DISCUSSION

The present study demonstrates an increase in TBARS and PCO content in mitochondria and enhanced carbonyl adduct formation with CI, CIII, and CV respiratory complexes in the myocardium of T. cruzi-infected mice. Our data show a direct correlation between the extent of protein carbonylation of specific subunits of the respiratory complexes and a loss in their catalytic activities during the course of infection and CCM development. Findings from the present study suggest that the impaired catalytic activity of the mitochondrial respiratory chain

Mitochondrial oxidative damage during T. cruzi infection

might be attributable to oxidative modification of the specific subunits of the respiratory complexes and HNE/ MDA generated from lipid peroxidation within the mitochondrial membranes. We have used the measurement of lipid peroxides and protein carbonyls to serve as an indicator of mitochondrial oxidative stress. Our data show a gradual increase in the mitochondrial levels of HNE/MDA and LHPO, the major products of lipid peroxidation [20]; and the consistent formation of PCO derivatives during the course of T. cruzi infection and disease development (Figs. 1, 2). Protein modifications elicited by direct oxidative attack on Lys, Arg, Pro, or Thr [30] or by a secondary reaction of Cys, His, or Lys residues with reactive compounds (e.g., 4-HNE, MDA) [31] can all lead to PCO accumulation. Considering that lipid peroxidation is preceded by PCO formation in cardiac mitochondria of infected mice, it is likely that the modification of protein residues by HNE/MDA-mediated crosslinking and Michael adduct formation contributes to an elevated PCO level in cardiac mitochondria. Concerning the mechanism responsible for the observed increase in mitochondrial oxidative stress, it is noted that substantial alterations in the activities of the mitochondrial respiratory complexes (CI, CIII, and CV) are observed in the myocardium of T. cruzi-infected mice during the course of disease progression [12]. CI and CIII are the prime sites of electron leakage to molecular oxygen and ROS generation in mitochondria [6,13–15]. It is, therefore, logical to postulate that consistent free radical generation due to deficiencies in respiratory complex activities contribute to sustained oxidative stress in the cardiac mitochondria of infected mice. Further, a decline in the enzymatic activity of manganese superoxide dismutase (MnSOD) is reported during the course of infection and disease development in chagasic myocardium [3]. MnSOD is the major oxygen radical scavenger in the mitochondrial matrix and acts as a first line of defense against the superoxide produced as a byproduct of oxidative phosphorylation [32]. We surmise that increased ROS release due to respiratory chain defects, along with limited MnSOD-mediated antioxidant defense capability, result in increased oxidative stressinduced modifications in cardiac mitochondria of infected mice. The major goal of this study was to determine whether oxidative modifications might be a mechanism underlying the decreased activity of the respiratory complexes in infected murine myocardium. The oxidative modifications may affect the processing and/or folding of the proteins [33], resulting in rapid turnover by proteolytic enzymes in mitochondria [34,35]. A decline in catalytic staining of the HNE-treated respiratory complexes (Fig. 3B) implies that the assembled complexes are also

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susceptible to oxidative damage. Our data show that the specific susceptibility of the subunits of the respiratory complexes, and not generalized protein oxidative damage, contributes to the respiratory dysfunction in chagasic hearts. For example, of the N42 subunits of CI, carbonyl adducts were detected with seven polypeptides in the cardiac mitochondria of T. cruzi-infected murine hearts (Fig. 4A). Of these, NDUFS1, NDUFS2, and NDUFV1 are the core subunits considered essential for electron transfer from NADH to ubiquinone and for the generation of protonmotive force [24, 36]. NDUFS4, also oxidatively modified in infected murine hearts, though not a core subunit, plays an important role in regulating the enzymatic efficiency of CI [36]. Genetic mutations leading to slight alterations in the encoded proteins, e.g., NDUFS1 [37], NDUFS2 [38], NDUFS4 [39], NDUFV1 [40], NDUFS7 [41], and NDUFV2 [42], are linked to CI deficiency in patients. Others have demonstrated oxidation/nitration of NDUFS2 and NDUFS8 subunits in association with a decline in CI activity in the mitochondria of human and bovine hearts [43]. The deficiency in CI activity by mutation/oxidation of CI subunits that were noted to be oxidatively modified in cardiac mitochondria of infected mice provides support for our hypothesis. We surmise that oxidatively modified structural subunits contribute to the inactivation of the assembled CI complex in chagasic hearts. Among the 11 components of CIII, we noted consistent carbonylation of CYC1 and core protein 1 (UQCRC1), and most likely core protein 2 (UQCRC2), in the cardiac mitochondria of infected mice. It is likely that oxidative modifications of CYC1 and core proteins contribute to a loss in CIII activity in the cardiac mitochondria of infected murine hearts, albeit through different mechanisms. CYC1, along with Reiske [2Fe– 2S] protein (ISP) and cytochrome b (CYB), forms the intermembrane-associated, central catalytic domain of CIII [44] that is involved in energy conservation and electron transfer from ubiquinol to cytochrome c (CYC). CYC1 accepts an electron from ISP and transfers it to soluble CYC [26]. The inability to accept or transfer electrons by oxidatively modified CYC1 would disrupt the electron flow-coupled proton translocation by the protonmotive Q cycle and thus directly affect CIII activity. Oxidative modifications of UQCRH (hinge protein) in acute hearts may further enhance the loss in CIII activity as it is speculated to facilitate the CYC1 and cytochrome c interaction during electron transfer [45]. Conversely, the core 1 and core 2 proteins, along with a small core-associated protein (UQCRQ), constitute the matrix portion of the CIII complex. The core proteins have high sequence similarity with soluble matrixprocessing peptidases (MPPs) [46] and are thought to be involved in the cleavage of targeting presequences

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after import of proteins in mitochondria [47]. Recently, structural studies have shown that peptidase activity of the core subunits may also be important in cleavage and processing of the signal sequence of the ISP protein [48]. The oxidatively modified core proteins may not process the presequence of ISP and other proteins, resulting in the accumulation of unprocessed proteins. If such is the case in chagasic hearts, incorporation of the misfolded ISP in CIII is likely to result in misassembly of the catalytic site and inhibition of the enzymatic activity of CIII complex. Of the two functionally distinct parts of F1F0 ATP synthase, the F1 subcomplex performs ATP synthesis and hydrolysis reactions, while the transmembrane F0 portion mediates proton transport [28]. The F1 catalytic moiety is an assembly of five subunits with the stoichiometry of a3h3gyq [28]. X-Ray crystallographic studies of the bovine F1 subcomplex have led to the suggestion that the g subunit rotates in the central cavity of the spherical a3h3 catalytic hexamer and is extended to the F0 moiety as a central stalk [49]. During catalysis, the rotation of the g subunit driven by protonmotive force through the F0 base determines the rate of ATP synthesis. It is suggested that the energy of the g chain rotation drives the release of ATP and increased affinity of ADP/P i to the three catalytic sites of the h subunit of the F1 moiety [50]. In our model system, carbonylation of a ~50 kDa band (a and/or h subunit), though detectable at all stages of infection and disease development, was enhanced in chronic hearts. A substantial oxidative modification of the g subunit was detected only in chronic hearts when ATP synthase activity and, subsequently, the ATP synthesis capacity of the cardiac mitochondria were also decreased [12]. Considering the direct correlation between the oxidative modification of the g subunit and the loss in CV activity in chagasic hearts, we surmise that oxidative damage of the g chain was disruptive and contributed to a loss in the enzymatic activity of F1F0ATP synthase in cardiac mitochondria of infected mice. To summarize, the results presented in this study enhance our understanding of the mechanisms involved in respiratory chain dysfunction in CCM. A depletion of mitochondrial DNA and repression of mitochondrial function-related transcripts has previously been associated with a decline in the level of the respiratory complexes in a murine model of T. cruzi infection [12,51]. The data presented in this study show that oxidative adduct formation, resulting in catalytic inactivation of the assembled complexes, may further impair the mitochondrial function and oxidative phosphorylation capacity of the myocardium during the course of T. cruzi infection and CCM progression. In addition, the oxidative damage of CI and CIII may potentiate

oxidative stress in the mitochondria, as a decline in the activities of these complexes is likely to result in the increased release of electrons to molecular oxygen and ROS formation. Acknowledgments — This work was supported in part by grants from American Heart Association (0160074Y) and National Institutes of Health (AI053098-01). Our thanks go to Dr. Istvan Boldogh for constructive discussions, Dr. John Papaconstantinou and Mr. Choksi Kashyap for providing the antibodies to mitochondrial subunits, and Ms. Mardelle Susman for editing and proofreading the manuscript. REFERENCES [1] World Health Organization. Chagas disease: tropical diseases progress in research, 1997–1998. WHO Tech. Rep. Ser. 1:1; 1999. [2] Higuchi Mde, L.; Benvenuti, L. A.; Martins Reis, M.; Metzger, M. Pathophysiology of the heart in Chagas’ disease: current status and new developments. Cardiovasc. Res. 60:96 – 107; 2003. [3] Wen, J.-J.; Vyatkina, G.; Garg, N. Oxidative damage during chagasic cardiomyopathy development: role of mitochondrial oxidant release and inefficient antioxidant defense. Free Radic. Biol. Med. (in press). [4] Tanowitz, H. B.; Gumprecht, J. P.; Spurr, D.; Calderon, T. M.; Ventura, M. C.; Raventos-Suarez, C.; Kellie, S.; Factor, S. M.; Hatcher, V. B.; Wittner, M., et al. Cytokine gene expression of endothelial cells infected with Trypanosoma cruzi. J. Infect. Dis. 166:598 – 603; 1992. [5] Martins, G. A.; Cardoso, M. A.; Aliberti, J. C.; Silva, J. S. Nitric oxide-induced apoptotic cell death in the acute phase of Trypanosoma cruzi infection in mice. Immunol. Lett. 63: 113 – 120; 1998. [6] Sadek, H. A.; Humphries, K. M.; Szweda, P. A.; Szweda, L. I. Selective inactivation of redox-sensitive mitochondrial enzymes during cardiac reperfusion. Arch. Biochem. Biophys. 406:222 – 228; 2002. [7] Piper, H. M.; Noll, T.; Siegmund, B. Mitochondrial function in the oxygen depleted and reoxygenated myocardial cell. Cardiovasc. Res. 28:1 – 15; 1994. [8] Cardoso, S. M.; Pereira, C.; Oliveira, R. Mitochondrial function is differentially affected upon oxidative stress. Free Radic. Biol. Med. 26:3 – 13; 1999. [9] Carrasco Guerra, H. A.; Palacios-Pru, E.; Dagert de Scorza, C.; Molina, C.; Inglessis, G.; Mendoza, R. V. Clinical, histochemical, and ultrastructural correlation in septal endomyocardial biopsies from chronic chagasic patients: detection of early myocardial damage. Am. J. Heart 113:716 – 724; 1987. [10] Uyemura, S. A.; Albuquerque, S.; Curti, C. Energetics of heart mitochondria during acute phase of Trypanosoma cruzi infection in rats. Int. J. Biochem. Cell Biol. 27:1183 – 1189; 1995. [11] Garg, N.; Popov, V. L.; Papaconstantinou, J. Profiling gene transcription reveals a deficiency of mitochondrial oxidative phosphorylation in Trypanosoma cruzi-infected murine hearts: implications in chagasic myocarditis development. Biochim. Biophys. Acta 1638:106 – 120; 2003. [12] Vyatkina, G.; Bhatia, V.; Gerstner, A.; Papaconstantinou, J.; Garg, N. Impaired mitochondrial respiratory chain and bioenergetics during chagasic cardiomyopathy development. Biochim. Biophys. Acta 1689:162 – 173; 2004. [13] Ide, T.; Tsutsui, H.; Kinugawa, S.; Utsumi, H.; Kang, D.; Hattori, N.; Uchida, K.; Arimura, K.; Egashira, K.; Takeshita, A. Mitochondrial electron transport complex I is a potential source of oxygen free radicals in the failing myocardium. Circ. Res. 85:357 – 363; 1999. [14] Lesnefsky, E. J.; Gudz, T. I.; Migita, C. T.; Ikeda-Saito, M.; Hassan, M. O.; Turkaly, P. J.; Hoppel, C. L. Ischemic injury to mitochondrial electron transport in the aging heart: damage to the iron-sulfur protein subunit of electron transport complex III. Arch. Biochem. Biophys. 385:117 – 128; 2001.

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