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• W2020188627 abstract "Article1 November 2007free access A role for cytochrome c and cytochrome c peroxidase in electron shuttling from Erv1 Deepa V Dabir Deepa V Dabir Department of Chemistry and Biochemistry, University of California at Los Angeles, Los Angeles, CA, USA Search for more papers by this author Edward P Leverich Edward P Leverich Department of Chemistry and Biochemistry, University of California at Los Angeles, Los Angeles, CA, USA Search for more papers by this author Sung-Kun Kim Sung-Kun Kim Department of Chemistry and Biochemistry, Center for Biotechnology and Genomics, Texas Tech University, Lubbock, TX, USA Department of Chemistry and Biochemistry, Baylor University, Waco, TX, USA Search for more papers by this author Frederick D Tsai Frederick D Tsai Department of Chemistry and Biochemistry, University of California at Los Angeles, Los Angeles, CA, USA Search for more papers by this author Masakazu Hirasawa Masakazu Hirasawa Department of Chemistry and Biochemistry, Center for Biotechnology and Genomics, Texas Tech University, Lubbock, TX, USA Search for more papers by this author David B Knaff David B Knaff Department of Chemistry and Biochemistry, Center for Biotechnology and Genomics, Texas Tech University, Lubbock, TX, USA Search for more papers by this author Carla M Koehler Corresponding Author Carla M Koehler Department of Chemistry and Biochemistry, University of California at Los Angeles, Los Angeles, CA, USA Molecular Biology Institute, University of California at Los Angeles, Los Angeles, CA, USA Jonsson Comprehensive Cancer Center, University of California at Los Angeles, Los Angeles, CA, USA Search for more papers by this author Deepa V Dabir Deepa V Dabir Department of Chemistry and Biochemistry, University of California at Los Angeles, Los Angeles, CA, USA Search for more papers by this author Edward P Leverich Edward P Leverich Department of Chemistry and Biochemistry, University of California at Los Angeles, Los Angeles, CA, USA Search for more papers by this author Sung-Kun Kim Sung-Kun Kim Department of Chemistry and Biochemistry, Center for Biotechnology and Genomics, Texas Tech University, Lubbock, TX, USA Department of Chemistry and Biochemistry, Baylor University, Waco, TX, USA Search for more papers by this author Frederick D Tsai Frederick D Tsai Department of Chemistry and Biochemistry, University of California at Los Angeles, Los Angeles, CA, USA Search for more papers by this author Masakazu Hirasawa Masakazu Hirasawa Department of Chemistry and Biochemistry, Center for Biotechnology and Genomics, Texas Tech University, Lubbock, TX, USA Search for more papers by this author David B Knaff David B Knaff Department of Chemistry and Biochemistry, Center for Biotechnology and Genomics, Texas Tech University, Lubbock, TX, USA Search for more papers by this author Carla M Koehler Corresponding Author Carla M Koehler Department of Chemistry and Biochemistry, University of California at Los Angeles, Los Angeles, CA, USA Molecular Biology Institute, University of California at Los Angeles, Los Angeles, CA, USA Jonsson Comprehensive Cancer Center, University of California at Los Angeles, Los Angeles, CA, USA Search for more papers by this author Author Information Deepa V Dabir1,‡, Edward P Leverich1,‡, Sung-Kun Kim2,5, Frederick D Tsai1, Masakazu Hirasawa2, David B Knaff2 and Carla M Koehler 1,3,4 1Department of Chemistry and Biochemistry, University of California at Los Angeles, Los Angeles, CA, USA 2Department of Chemistry and Biochemistry, Center for Biotechnology and Genomics, Texas Tech University, Lubbock, TX, USA 3Molecular Biology Institute, University of California at Los Angeles, Los Angeles, CA, USA 4Jonsson Comprehensive Cancer Center, University of California at Los Angeles, Los Angeles, CA, USA 5Department of Chemistry and Biochemistry, Baylor University, Waco, TX, USA ‡These authors contributed equally to this work *Corresponding author. Department of Chemistry and Biochemistry, Molecular Biology Institute, Jonsson Comprehensive Cancer Center, Box 951569, University of California at Los Angeles, Los Angeles, CA 90095, USA. Tel.: +1 310 794 4834; Fax: +1 310 206 4038; E-mail: [email protected] The EMBO Journal (2007)26:4801-4811https://doi.org/10.1038/sj.emboj.7601909 PDFDownload PDF of article text and main figures. ToolsAdd to favoritesDownload CitationsTrack CitationsPermissions ShareFacebookTwitterLinked InMendeleyWechatReddit Figures & Info Erv1 is a flavin-dependent sulfhydryl oxidase in the mitochondrial intermembrane space (IMS) that functions in the import of cysteine-rich proteins. Redox titrations of recombinant Erv1 showed that it contains three distinct couples with midpoint potentials of −320, −215, and −150 mV. Like all redox-active enzymes, Erv1 requires one or more electron acceptors. We have generated strains with erv1 conditional alleles and employed biochemical and genetic strategies to facilitate identifying redox pathways involving Erv1. Here, we report that Erv1 forms a 1:1 complex with cytochrome c and a reduced Erv1 can transfer electrons directly to the ferric form of the cytochrome. Erv1 also utilized molecular oxygen as an electron acceptor to generate hydrogen peroxide, which is subsequently reduced to water by cytochrome c peroxidase (Ccp1). Oxidized Ccp1 was in turn reduced by the Erv1-reduced cytochrome c. By coupling these pathways, cytochrome c and Ccp1 function efficiently as Erv1-dependent electron acceptors. Thus, we propose that Erv1 utilizes diverse pathways for electron shuttling in the IMS. Introduction The mitochondrial intermembrane space (IMS) is conducive to disulfide bond formation (Koehler et al, 2006; Wiedemann et al, 2006; Herrmann and Kohl, 2007), suggesting that the redox environment is oxidizing, similar to that of the bacterial periplasm (Kadokura et al, 2003). Thiol-trapping assays indicated that the small Tim proteins with the twin CX3C motif (Tim8, Tim9, Tim10, and Tim13) form disulfide bonds (Curran et al, 2002a, 2002b; Koehler, 2004). Structural and mutational studies confirmed that the small Tim proteins contain juxtapositional disulfide linkages (Allen et al, 2003; Lu et al, 2004; Webb et al, 2006). Subsequently, a new import pathway was identified for the small Tim proteins and other substrates with a twin CX9C motif (Chacinska et al, 2004; Mesecke et al, 2005; Rissler et al, 2005; Terziyska et al, 2005; Gabriel et al, 2007). Specifically, Mia40 functions as a tethering component in the IMS by forming a transient disulfide bond with the incoming substrate, trapping the precursor in the IMS. The substrates are released from Mia40 by a mechanism that may involve rearrangement of disulfide bonds. It has been proposed that the sulfhydryl oxidase Erv1 participates in recycling Mia40 by reoxidizing Mia40's cysteine residues (Mesecke et al, 2005). Thus, electrons may be transferred from the imported proteins to Erv1 via Mia40. Erv1 also functions in the maturation of cytosolic FeS centers (Lange et al, 2001), suggesting its diverse functions in the IMS. Both Mia40 and Erv1 are required for viability (Lisowsky, 1992; Chacinska et al, 2004). Erv1 was first identified by Lisowsky and co-workers (Lisowsky, 1992) and is a member of the flavin-dependent sulfhydryl oxidase family (Coppock and Thorpe, 2006). Erv1 shares homology with Erv2, a protein that functions in the endoplasmic reticulum (Sevier et al, 2001), with Quiescin-sulfhydryl oxidases that function in the extracellular matrix (Hoober et al, 1996), and with the E10R protein of the pox virus that inserts disulfide linkages into the virion's coat proteins (Senkevich et al, 2002). These sulfhydryl oxidases are predicted to use oxygen (O2) as an electron acceptor for the oxidation of cysteine residues to protein disulfides, generating hydrogen peroxide (H2O2) in the process (Coppock and Thorpe, 2006). Erv1 has three potential redox centers, two pairs of redox-active cysteines (C30–C33 and C130–C133) and a noncovalently bound FAD (Hofhaus et al, 2003; Farrell and Thorpe, 2005). The C30–C33 pair is important for the formation of an intermolecular disulfide bond between two Erv1 monomers and is required for in vivo function (Lee et al, 2000; Hofhaus et al, 2003). The C30–C33 pair, however, is not required for sulfhydryl oxidase activity in vitro, suggesting that the C30–C33 pair is not required for Erv1 enzymatic activity, but instead serves a structural role. In contrast, the C130–C133 pair, which is positioned near the likely flavin-binding site, is important for the redox activity of Erv1 (Hofhaus et al, 2003; Farrell and Thorpe, 2005), as mutagenesis of either cysteine to a serine or alanine abolishes sulfhydryl oxidase activity. Farrell and Thorpe (2005) have estimated an Em value of −178 mV for the midpoint potential of the flavin in ALR, but were not able to reduce the disulfide near ALR's flavin-binding site (i.e., the disulfide equivalent to C130–C133 in Erv1), raising the possibility that this putative redox-active cysteine pair in ALR may have a considerably more negative midpoint potential. Bacteria utilize a wide array of terminal electron acceptors for the oxidative formation of protein disulfide bonds (Bader et al, 1999). Erv1 may also display a level of flexibility with regard to electron acceptors, because yeast cells lacking the mitochondrial genome and respiratory pathway still import mitochondrial proteins. In this study, we have used genetic and biochemical approaches to characterize Erv1 and investigate potential pathways in which Erv1 might be oxidized by different terminal electron acceptors. We show that Erv1 forms a complex with cytochrome c (cyt c) in organello and in vitro. With in vitro assays, we demonstrate that Erv1 shuttles electrons to both O2 and then cytochrome c peroxidase (Ccp1) and oxidized cyt c. These studies suggest that, like the prokaryotic disulfide catalytic system, Erv1 utilizes several terminal electron acceptor pathways. Results To begin characterization of the electron acceptor pathways utilized by the sulfhydryl oxidase Erv1, we generated temperature-sensitive (ts) mutants using error-prone PCR (Koehler et al, 1998). Two strains harboring alleles erv1-101 and erv1-12 on centromeric plasmids as well as the tim9-3 control (Leuenberger et al, 2003) grew at 25°C but arrested growth at 37°C on rich glucose (YPD) and ethanol-glycerol (YPEG) media (Figure 1A). We subsequently analyzed the steady-state levels of mitochondrial proteins using α-ketoglutarate dehydrogenase (Kdh) as a loading control (Figure 1B). The abundance of outer membrane markers porin and Tom40 was not affected (data not shown); however, the steady-state levels of the substrates of the MIA40 pathway (Erv1, Tim9, Tim10, Tim12, and Tim13) and Mia40 were markedly reduced in mitochondria that were shifted to 37°C. In addition, Tim22, Tim23, and Tim54 levels were reduced. The reduction in the TIM22 import components and Tim23 likely results from a decreased import of the small Tim proteins via the MIA import pathway. Interestingly, Ccp1 migrated aberrantly in mitochondria shifted to 37°C, and cyt c abundance was not obviously affected at 37°C. Taken together, Erv1 dysfunction affects proteins primarily imported by the MIA40 import pathway and the TIM22 import pathway. Figure 1.Steady-state levels of IMS proteins are reduced in mitochondria lacking functional Erv1. (A) Cells were grown to mid-log phase at 25°C and then serially diluted by a factor of 3 onto rich glucose (YPD) and ethanol-glycerol (YPEG) media before incubation at 25°C and 37°C. Strains included the parent (WT), tim9-3 mutant, and erv1 mutants harboring alleles erv1-101 and erv1-12. Plates were photographed after 3–4 days. (B) Steady-state levels of mitochondrial proteins (50 and 100 μg) were investigated by immunoblot analyses with antibodies against mitochondrial proteins indicated to the left. Mitochondria were purified from the parent (WT) and erv1-101 and erv1-12 strains grown either at the permissive temperature or shifted to the restrictive temperature of 37°C for 7 h. Download figure Download PowerPoint Cyt c partners with Erv1 in organello To identify Erv1 partner proteins, we placed 10X-His tags on the C-termini of both Erv1 and Ccp1 and substituted the His-tagged versions on multicopy plasmids for the endogenous genes (Figure 2A and B). The Erv1-His and Ccp1-His strains grew like wild type (WT, data not shown), indicating that the His-tagged proteins were functional. Initial attempts to express Erv1-His from a centromeric plasmid yielded a marked decrease in the Erv1-His protein levels. Therefore, Erv1-His expression from the multicopy plasmid resulted in increased expression of Erv1-His relative to WT mitochondria, whereas cyt c and Ccp1 abundance were comparable with that of WT mitochondria (Supplementary Figure 1). Mitochondria were solubilized in 1.0% digitonin (T) and incubated with Ni2+ agarose. After washing to remove non-bound proteins (S), the bound proteins (B) were eluted in sample buffer and separated by SDS–PAGE. Immunoblotting analysis showed that cyt c and a fraction of the Mia40 co-purified with Erv1-His, whereas Ccp1 did not co-purify. Note that in independent purifications, the fraction of cyt c that co-purified with Erv1 varied from approximately 70 to 100% (data not shown), most likely caused by the increased abundance of Erv1-His. With Ccp1-His, a fraction of the cyt c but not Erv1 or Mia40 co-eluted. In a control reaction, WT mitochondria were treated identically and the tested proteins showed no affinity for the Ni2+ agarose (Figure 2C). Therefore, cyt c is a partner protein with both Erv1 and Ccp1, but Ccp1 seemingly does not form a complex with Erv1. Figure 2.Erv1 and cyt c form a complex in the IMS. (A) Mitochondria from a strain expressing a C-terminal histidine-tagged Erv1 (Erv1-His) were solubilized at 5 mg/ml in 1% digitonin. As a control, 100 μg of extract was withdrawn (T), and 500 μg lysate was incubated with Ni2+-agarose beads. The beads were washed, and bound proteins (B) were eluted with SDS–PAGE sample buffer. To assess the effectiveness of binding, 100 μg of the unbound protein fraction (S) was also included. Proteins were analyzed by immunoblotting with polyclonal antibodies against Mia40, Ccp1, Erv1, and cyt c. (B) Similar to (A), except that a C-terminal histidine-tagged Ccp1 was utilized. (C) The control reaction in which WT mitochondria have been treated identically. (D) Strains (WT, Δtim54, Δccp1, Δcyc3, and erv1 mutants, erv1-101 and erv1-12) were serially diluted on YPD medium in the presence and absence of ethidium bromide (EtBr) and incubated in aerobic (+O2) or anaerobic (−O2) conditions at 25°C. Plates were photographed after 3–5 days. Petite-negative Δtim54 was included as a control. (E) A cross between erv1-101 and Δccp1 that yielded tetratype segregation was analyzed for growth as described in (D). Download figure Download PowerPoint Terminal electron acceptor pathways are required in both aerobic and anaerobic growth conditions, and different pathways are utilized in prokaryotes depending on the oxidative state (Bader et al, 1999). First, we tested whether the erv1 mutants grew under anaerobic conditions and in the presence of ethidium bromide, which induces loss of mitochondrial DNA and respiratory ability (Figure 2D). The erv1 mutants grew, albeit more slowly, when the mitochondrial genome was absent (+EtBr) under aerobic conditions (+O2). The erv1-101 also showed a slower growth rate under anaerobic conditions (−O2), whether mitochondrial DNA was present or absent (Figure 2D). In contrast, the negative control Δtim54, which is petite negative, failed to grow under the conditions tested. Because yeast express two different cyt c genes (CYC1 and CYC7) under different O2 concentrations (Burke et al, 1997), we used a Δcyc3 strain to generate a cyt c null strain, as Cyc3 is the heme lyase for assembly of holocytochrome c (Dumont et al, 1988). The Δcyc3 and Δccp1 strains grew under anaerobic and aerobic conditions in the presence and absence of the mitochondrial genome (Figure 2D). We tested CYC3 and ERV1 for synthetic lethality at 25°C (Figure 2E). Tetrad analysis showed that the strain deleted for cyc3 in the erv1-101 mutant background was viable in aerobic conditions at 25°C. However, the Δcyc3 erv1-101 mutant failed to grow in anaerobic conditions. Thus, CYC3 and ERV1 are synthetic lethal in anaerobic conditions, indicating that cyt c and Erv1 interact genetically as well as physically. Erv1 has three different midpoint potentials corresponding to the two redox-active cysteine pairs (C30–C33 and C130–C133) and the FAD moiety Previous studies have suggested that C30–C33 plays a structural role in the formation of Erv1 dimers, whereas the C130–C133 pair has sulfhydryl oxidase activity and is situated near the FAD (Lee et al, 2000; Hofhaus et al, 2003). We measured the disulfide/dithiol redox properties of Erv1 at pH 7.0 using GSH/oxidized glutathione (GSSG) redox buffers to poise samples at defined redox potentials (Eh) values ranging from −100 to −240 mV and reduced dithiothreitol (DTT)/oxidized (DTTox) redox buffers to poise samples at Eh values ranging from −220 to −380 mV (Figure 3; Krimm et al, 1998; Hirasawa et al, 1999). Recombinant Erv1 was incubated in the indicated redox potential buffer and then treated with mBBr to form fluorescent covalent adducts of Erv1 thiols (mBBr does not react with disulfides). The titration results were independent of the redox equilibration time, over the range from 1.5 to 3.5 h, and were also independent of the concentration of the redox buffer, over the range from 1 to 5 mM, indicating the likelihood of good redox equilibration between Erv1 and the ambient potential imposed by the redox buffers. Additional support for the establishment of good redox equilibration during these titrations comes from the fact that all of the titrations gave excellent fits to the Nernst equation for a two-electron redox couple. Figure 3A shows the results of a typical titration and reveals the presence of two two-electron components. The average values (based on three titrations) for the redox midpoint potentials (Em) of the two components are −330 and −150 mV, respectively (the use of the average deviation as an approximate measure of the experimental uncertainty suggests that the experimental uncertainty does not exceed±10 mV). The Em value and the n=2 character of the titration curve, as well as the known specificity of mBBr for chemical modification of thiols (Krimm et al, 1998; Hirasawa et al, 1999), indicate that the redox titration results are most readily interpreted in terms of the presence of two separate disulfide/dithiol redox couples in Erv1. Figure 3.Oxidation–reduction titrations of Erv1 reveal three midpoint redox potentials. (A, B) Redox titrations of dithiol/disulfide couples in Erv1 were carried out using DTT or glutathione redox buffers. Redox equilibration was performed for 2 h at pH 7.0 with total redox buffer concentrations of 2.0 mM. Data in all titrations were fitted to the Nernst equation for a two-electron carrier. In (A), the titration was tested using the mBBr method. The best fit to the mBBr fluorescence magnitude versus Eh value was obtained for the presence of two separate (n=2) components, with Em values of −330 and −150 mV, respectively. In (B), the titration was performed using intrinsic tryptophan fluorescence to monitor the redox state of Erv1. The best fit to the data was for a single (n=2) component with Em=−315 mV. (C) The redox midpoint potential of the flavin group was determined by electrochemical titration of Erv1 (10.0 μM) at 10°C in 50 mM KPO4, pH 7.0, in the presence of 10 μM benzyl viologen, 10 μM anthraquinone-2-sulfonate, and 10 μM 2-hydroxy-1,4-napthoquinone as redox mediators. The difference in absorbance at 465 nm minus that at 550 nm was used to monitor the extent of FAD reduction. The value of this absorbance difference for the fully oxidized FAD was set as 1.0 and the value for the fully reduced FAD was set at 0.0, to define the relative absorbance difference scale. The open circles represent the measured relative absorbance and the solid line is the computer best fit of these data to the Nernst Equation for a two-electron redox couple with Em=−215 mV. Representative individual spectra taken at different defined Eh values are shown in Supplementary Figure 3. Download figure Download PowerPoint Added support for the notion of two separate thiol-based redox couples comes from DTNB assays of the thiol content of Erv1 samples poised at defined redox potentials (See Table I). In particular, the difference in thiol content of Erv1 samples poised at a potential where the Em=−150 mV transition would be either completely reduced or completely oxidized is 2.0±0.3 mol of thiol per mol of Erv1, exactly what would be predicted for the reduction of one intramolecular disulfide per Erv1 monomer. Similarly, the difference in thiol contents of Erv1 samples poised at potential where the Em=−320 mV transition would be either completely reduced or completely oxidized is 0.9±0.3 mol of thiol per mol of Erv1, exactly what would be predicted for the reduction of one intermolecular disulfide between two Erv1 monomers. Table 1. DNTB addition assay to Erv1 equilibrated at different Eh values (based on calculated Em potentials) Redox potential at which Erv1 is equilibrated Mole of free thiol per mole of Erv1 Fully reduced, denatured 6.0±0.1 Fully reduced (−400 mV) 3.0±0.1 Half reduced (−240 mV) 2.1±0.2 Fully oxidized (−90 mV) 0.1±0.1 An attempt was made to confirm the assignment, based on DTNB thiol assays of samples poised at defined Eh values, of the Em=−330 mV to an intermolecular disulfide formed between two Erv1 monomers, using non-reducing SDS–PAGE gels of Erv1 samples that had been poised at defined Eh values centered around −330 mV before SDS–PAGE analysis. The amount of dimer increases as the Eh is increased over the range from −380 to −220 mV (Supplementary Figure 2), consistent with an Em value of approximately −330 mV for an intermolecular disulfide. Figure 3B shows the results of a redox titration, achieved under conditions identical to those of Figure 3A (except that mBBr was not added), but with tryptophan fluorescence used to monitor redox-dependent changes in Erv1. The data are most readily interpreted in terms of a two-electron redox couple, with Em=−315±10 mV at pH 7.0, in which the microenvironment of at least one Erv1 tryptophan residue differs in the oxidized and reduced forms of the protein. The fact that the Em value of the two-electron couple detected in tryptophan fluorescence is indistinguishable, within the combined experimental uncertainties of the measurements, from the Em value of the more negative couple seen in the mBBr titrations, leads us to conclude that it is the monomer/dimer transition that alters the microenvironment of at least one Erv1 tryptophan. Flavin analysis of Erv1 indicated the presence of 1.0±0.1 mole of FAD per mole of recombinant Erv1 (no evidence was found for any FMN in the protein). Figure 3C shows the results of a typical titration of the FAD group of Erv1 at pH 7.0, using the absorbance in the visible region to monitor the oxidation state of FAD. The data showed an excellent fit to the Nernst equation for a single n=2 component with an Em value of −215 mV. Representative spectra generated during the potentiometric titration of the flavin group are included (Supplementary Figure 3). The absence of well-defined isosbestic points in these spectra arises from a slow drift in the instrument baseline during the approximately 2 h required for the full titration. Under the conditions employed, no red anionic or neutral blue flavin semiquinone could be detected, in contrast to the results reported for ALR (Farrell and Thorpe, 2005). It should also be pointed out that while the flavin in ALR has been suggested to have a more positive redox potential than the disulfide/dithiol centers present in ALR (Farrell and Thorpe, 2005), this is not the case for Erv1 where the FAD has an Em value that is significantly more negative than that of the Em=−150 mV intramolecular disulfide. Erv1 and cyt c interact in a 1:1 complex and Erv1 can directly reduce cyt c Farrell and Thorpe (2005) have previously suggested that cyt c is the physiological oxidant of Erv1. In Figure 2A, we demonstrated that cyt c partners with Erv1 in vivo. To further probe the stoichiometry of the Erv1:cyt c complex, we examined the interactions of our characterized recombinant Erv1 and yeast cyt c, using a spectral perturbation technique to monitor protein–protein interactions. This technique has been used to characterize a large number of protein complexes, including the one formed between cyt c and Ccp1 (Erman and Vitello, 2002). The solid line in Figure 4A shows the difference spectrum arising from the interaction of Erv1 with cyt c in low ionic strength buffer (i.e., it represents the spectrum of a 1:1 mixture of the two proteins from which has been subtracted the sum of the spectra of the two separate proteins). This difference spectrum was not observed if either BSA or Ccp1 replaced cyt c, suggesting that the difference spectrum of Figure 4A does not arise from some nonspecific artifact. Figure 4A also shows (see the dashed line) that no spectral perturbations are observed at high ionic strength (solid NaCl was added to the sample used for the low ionic strength spectrum to yield an approximate concentration of 250 mM and the spectrum was re-recorded). The simplest explanation for this observation is that electrostatic forces, which weaken at high ionic strength, make an important contribution to stabilizing the Erv1:cyt c complex. Figure 4B shows the results of an experiment, carried out at low ionic strength, in which the magnitude of spectral changes arising from complex formation were plotted against varying concentrations of cyt c at a fixed concentration of Erv1. The data give a good fit to the theoretical curve for the presence of a 1:1 Erv1:cyt c complex of very high affinity (computer simulations suggest that the Kd for complex formation under these conditions is probably ⩽100 nM). As protein–protein complex formation can alter the Em value of protein prosthetic groups (Ondrias et al, 1985), the FAD group of Erv1 and the heme of cyt c were titrated in a 1:1 mixture of the proteins at the low ionic strength that favors complex formation between the proteins. The Em value of the Erv1 FAD was unaffected by the presence of cyt c and the Em value of cyt c was found to be +250 mV, regardless of whether Erv1 was present or absent (data not shown). The value obtained in the absence of Erv1 is in good agreement with the published literature (Wallace, 1984). Figure 4.Erv1 and cyt c form a 1:1 complex, in which Erv1 reduces cyt c directly. (A) A representative run showing the difference spectra resulting from complex formation between a 1:1 M ratio of Erv1 and cyt c. The solid line represents the spectrum arising from the complex under low ionic buffer (10 mM KPO4, pH 7.0); dashed line represents the spectrum from the complex in high ionic buffer (250 mM NaCl, pH 7.0). (B) Binding isotherms for complex formation at pH 7.0 between Erv1 and cyt c were monitored by changes in the UV/visible region of the absorbance spectrum. Increasing concentrations of cyt c as indicated were added to Erv1. The spectrum was not perturbed further when cyt c was added at an Erv1:cyt c molar ratio of greater than 1:1, suggesting a 1:1 complex. (C) The reduction of cyt c by Erv1 was measured at 550 nm as a function of time. The concentration of cyt c was fixed at 10 μM; Erv1 and DTT concentrations were 2.3 μM and 0.4 mM, respectively. Download figure Download PowerPoint That Erv1 and cyt c form a complex suggests that Erv1 might shuttle electrons directly to cyt c. This possibility was tested directly, under anaerobic conditions, by observing changes in absorbance at 550 nm that arise from reduction of the cyt c heme when DTT-reduced Erv1 is added to ferric cyt c (Figure 4C). Erv1, at the concentration used in these experiments, does not contribute significantly to the absorbance at this wavelength. As shown in Figure 4C, the addition of reduced Erv1 indeed cause rapid reduction of cyt c, denoted by the marked increase in absorbance after Erv1 addition. In control experiments, in which DTT alone was added to cyt c in the absence of Erv1, a much slower rate of heme reduction was observed, indicating that DTT is a kinetically poor reductant for cyt c and confirming that Erv1 can reduce the heme of cyt c directly in vitro. Ccp1 and cyt c compete with O2 in Erv1-mediated oxidation of DTT Ccp1 serves as a general catalyst to remove H2O2 formed in mitochondria during aerobic metabolism (Boveris, 1976). However, despite extensive in vitro characterization of Ccp1 (Erman and Vitello, 2002), few studies have clarified its specific physiological role in yeast. We tested CCP1 and ERV1 for synthetic lethality at 25°C (Figure 5A). Tetrad analysis showed that strains deleted for ccp1 in the erv1 mutant background were viable at 25°C when the mitochondrial genome was absent. In addition, the erv1-101 mutant grew, albeit slowly, in anaerobic conditions, similar to that in Figure 2D, regardless if CCP1 was present. Thus, CCP1 and ERV1 are not synthetic lethal, indicating that CCP1 is not strictly required for growth under the conditions investigated. Figure 5.(A) A cross between erv1-101 and Δccp1 that yielded tetratype segregation was analyzed for growth as described in Figure 2D. (B) The e" @default.
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• W2020188627 date "2007-11-01" @default.
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• W2020188627 title "A role for cytochrome c and cytochrome c peroxidase in electron shuttling from Erv1" @default.
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