Cytochrome c signalosome in mitochondria

1 Post-print of: Eur Biophys J (2011) 40:1301–1315 Cytochrome c signalosome in mitochondria Irene Díaz-Moreno (1), José M. García-Heredia (1), Antonio Díaz-Quintana (1) and Miguel A. De la Rosa (1) (1) Instituto de Bioquímica Vegetal y Fotosíntesis, Centro de Investigaciones Científicas Isla de la Cartuja (cicCartuja), Universidad de Sevilla-CSIC, Avda. Américo Vespucio 49, 41092 Sevilla, Spain Abstract Cytochrome c delicately tilts the balance between cell life (respiration) and cell death (apoptosis). Whereas cell life is governed by transient electron transfer interactions of cytochrome c inside the mitochondria, the cytoplasmic adducts of cytochrome c that lead to cell death are amazingly stable. Interestingly, the contacts of cytochrome c with its counterparts shift from the area surrounding the heme crevice for the redox complexes to the opposite molecule side when the electron flow is not necessary. The cytochrome c signalosome shows a higher level of regulation by post-translational modifications—nitration and phosphorylation—of the hemeprotein. Understanding protein interfaces, as well as protein modifications, would puzzle the mitochondrial cytochrome c-controlled pathways out and enable the design of novel drugs to silence the action of pro-survival and pro-apoptotic partners of cytochrome c. Keywords Biointeractome, Cytochrome c, Transient complex, Electron transfer, Mitochondria, Signalosome Abbreviations Adx Adrenodoxin AdxR NADPH-dependent adrenodoxin reductase bc 1 Cytochrome bc 1 complex CB Cytochrome binding 2 Cb 5 Cytochrome b 5 Cb 5R NADH-dependent cytochrome b 5 reductase Cc Cytochrome c Cc 552 Cytochrome c 552 CcO Cytochrome c oxidase CcP Cytochrome c peroxidase CH1 Collagen homologous 1 CH2 Collagen homologous 2 CL CardioLipin ET Electron transfer GALDH l-GAlactono-1,4-Lactone DeHydrogenase IMM Inner mitochondrial membrane IMS Intermembrane mitochondrial space n-Cc 3 Nitrated cytochrome c NMR Nuclear magnetic resonance OMM Outer mitochondrial membrane p-Cc Phosphorylated cytochrome c PCD Programmed cell death PKCβ Protein kinase C β PKCδ Protein kinase C δ PRE Paramagnetic relaxation enhancement PTB PhosphoTyrosine binding R(N)OS Reactive (nitrogen)oxygen species Sco Synthesis of cytochrome c oxidase SH2 Src homology 2 WT Wild-type Cytochrome c: a multitasking post-translationally modified protein 4 Cells must be considered as a crowded system, in which any particular protein may be in contact with lots of other proteins, nucleic acids, metabolites, etc. It thus requires a way of recognition that allows the specific interaction with only a few of them. Such recognition mechanisms between biomolecules occur in a wide range of time scales. On one hand, stable complexes, with a lifetime ranging from minutes to days, involve high affinity and high specificity binding. On the other, weak complexes, with a lifetime within the s–μs range, are formed when a fine balance between specificity of binding and high turnover rate is sought, resulting in adducts with equilibrium dissociation constants in the μM or even mM range (Ubbink 2009; Bashir et al. 2011; Díaz-Moreno and De la Rosa 2011a, b). Intriguingly, contrary to what one might think, these molecular recognition mechanisms are not uncommon, being crucial in electron transfer (ET) chains—such as respiration, peroxidation and steroid hormone biosynthesis. Both types of complexes—stable versus weak—meet on cytochrome c (Cc), which is an excellent model: Cc is not only able to form protein adducts with different lifetimes, but is also a highly conserved protein along evolution. Cc is a small soluble metalloprotein of around 12.5 kDa located at the intermembrane mitochondrial space (IMS). It folds in four α-helices and two extended loops, which sandwich on the heme group and provide its two axial ligands, His18 and Met80 (Louie and Brayer 1990; Reincke et al. 2001; Jeng et al. 2002). The porphyrin ring is covalently bound to the cysteine residues of the CXXCH motif and partially exposed to solvent, a feature that is essential for Cc to carry out most of its functions. Under physiological, non-stressed conditions, Cc plays a key role in energy metabolism by a controlled redox interaction with its counterparts in the mitochondrial respiratory chain (Moore and Pettigrew 1990). Shuttling electrons between the two membrane-bound protein complexes cytochrome bc 1 (bc 1) and cytochrome c oxidase (CcO) requires rapid adduct formation and rapid protein dissociation, as well as a proper and efficient orientation of the two proteins of the transient complex to optimize the ET. Upon an apoptotic signalling stimulus due to DNA damage or an excess in Reactive (Nitrogen)Oxygen Species (R(N)OS), the cells may undergo disturbances of their regulatory pathways that lead to the release of mitochondrial Cc. Although Cc is preferably reduced in the cytosol, the interaction with Apaf-1 is independent of its redox state. The Cc/Apaf-1 complex forms the apoptosome, which is the enzymatic machinery of apoptosis (Cai et al. 1998; Orrenius 2007). Thus, the apoptosome is the result of stable and long-lived interactions of Cc with other protein partners (Acehan et al. 2002). Cell life is governed by transient interactions of Cc inside the mitochondria, but the cytoplasmic adducts of Cc that lead to cell death are amazingly stable (Fig. 1; Table 1). There are only two exceptions. The first one is the long-lived complex between Cc and CardioLipin (CL), a lipid allocated at the inner mitochondrial membrane (IMM). Such a complex triggers CL peroxidation and further Cc release at the beginning of apoptosis. The last step before Cc release is the highly dynamic interaction between Cc and Bcl-x2 at the outer mitochondrial membrane (OMM). It is even more interesting that (1) all IMS-protein contacts performed by Cc show a high turnover and (2) that they are all involved in ET reactions—for instance, binding to bc 1, CcO, cytochrome b 5 (Cb 5), cytochrome c peroxidase (CcP), Erv1 and p66Shc. In 5 contrast, the interactions of Cc at the cytosol—upon Bcl-x2 and Apaf-1 binding—are not redox, regardless of whether they are stable or transient (Fig. 1; Table 1). Within such a frame, this review is focused on the complex regulatory network of transient intermolecular contacts hovering on Cc, a moonlighting hemeprotein performing a high number of functions in the IMS and OMM (Fig. 2). As for other proteins, the multitasks ascribed to Cc can be regulated by post-translational modifications and, in particular, by nitration or phosphorylation of tyrosine residues. Both modifications, mutually exclusive, can affect the way that Cc interacts with its physiological partners—either in mitochondria or in cytosol—but such effects are themselves highly dependent on which tyrosine is modified. Nitration of Cc is caused by the excess of mitochondrial R(N)OS, which can diffuse from extramitochondrial compartments into mitochondria or can be generated accidentally by the activity of the mitochondrial respiratory chain (Chance et al. 1979; Chen et al. 2003). Respiration is drastically impaired by nitration, no matter which tyrosine is nitrated (RodríguezRoldán et al. 2008). In contrast, only nitrated Tyr46, Tyr48 and Tyr74 block the apoptotic reaction (García-Heredia et al. 2010, unpublished data). Whereas Tyr48 and Tyr74 are highly conserved along evolution, Tyr46 is present in Cc from humans but not from other sources. An example of gain-of-function modification is the increase in peroxidase activity of Cc upon nitration (Cassina et al. 2000; Batthyány et al. 2005; García-Heredia et al. 2010), mainly of the nitrated Cc (n-Cc) species that behave as high-spin proteins (Díaz-Moreno et al. 2011c). Cytochrome c phosphorylation, whose specific phosphorylating kinase is still unknown, has been shown to inhibit ET between Cc and CcO (Lee et al. 2006; Yu et al. 2008). However, only phosphorylation of Cc-Tyr48 disrupts apoptosome activation (Pecina et al. 2010; GarcíaHeredia et al. 2011). Besides the tyrosines susceptible to phosphorylation, there are two other phosphorylation residues on human Cc, namely Thr28 and Ser47 (Zhao et al. 2011; Hüttemann et al. 2011a, b). Nevertheless, their functional consequences remain unknown. The role of Cc in mitochondrial respiration chain The cytochrome bc 1 and cytochrome c complex The mitochondrial respiratory chain couples ET from reduction equivalents to molecular oxygen, with vectorial proton translocation across the lipid membrane. The generated electrochemical proton gradient drives ATP synthesis. Four multisubunit enzymes (complexes I–IV) are embedded in the IMM. The soluble protein Cc, located in the IMS, shuttles electrons between cytochrome bc 1 (complex III or ubiquinol/cytochrome c oxidoreductase) and CcO (complex IV) (Saraste 1999). These interactions are highly transient, enabling high turnover rates, which are essential for the continuous electron flow through the different components of the respiratory chain (Fig. 3). Cytochrome bc 1 is a 500-kDa homodimeric multisubunit integral membrane protein complex. The catalytic core comprises cytochrome b, with two noncovalently attached heme groups; the so-called Rieske protein, with an iron-sulfur cluster; and cytochrome c 1 (Cc 1), with a covalently attached heme c group (Berry et al. 2000). The enzyme catalyzes the ET from 6 ubiquinol to Cc coupled to the net translocation of protons over the mitochondrial membrane (Berry et al. 2000; Fig. 3). The crystal structure of the mitochondrial bc 1–Cc complex reveals that there is a small nonpolar contact area (ca. 957 Ǻ2), including a cation–π interaction with the heme cofactors in the center surrounded by charged residues whose contribution to the interaction is mainly electrostatic (Lange and Hunte 2002). This is consistent with the two-step model of ET complex formation, in which the final complex first entails a primary unspecific recognition via electrostatic steering as an encounter that can be transiently stabilized to yield a productive and specific complex as an outcome. On one hand, the electrostatic component between Cc 1 and Cc accelerates protein association by limiting diffusion space, despite keeping the pairs of complementary charged residues far enough to avoid forming salt bridges. On the other, hydrophobic and cation–π contact pairs define an area around the core of the bc 1–Cc interface defined by the heme cleft with their pyrrole C rings pointing toward each other, which allows ET to occur directly from c 1 heme to Cc heme (Lange and Hunte 2002). This has also been inferred not only from the orientation and close proximity of the heme groups in the bc 1–Cc crystallographic structure, but also from the estimated ET rates (Saraste 1999) and stopped-flow measurements revised by Yu et al. (2002). Such ET rates perfectly match those calculated by laser flash photolysis using ruthenium-labeled Cc derivatives (Tian et al. 2000; Engstrom et al. 2003). The rates are ionic strength-dependent. The first-order rate constant does not change as the ionic strength increases from 10 to 50 mM, but diminishes significantly with increasing ionic strength. At high ionic strength, the rate constant becomes Cc 1 concentration-dependent, which is indicative of a second-order kinetics (Yu et al. 2002, for revision). Interestingly, the ET between Cc 1 and Cc is fully reversible, consistent with the fact that the reduction potentials of both cytochromes are nearly the same. More recently, a higher resolution bc 1–Cc structure was resolved, showing a substantially hydrated interface in which the relatively low surface complementary between the two hemeproteins provides space for hydration (Solmaz and Hunte 2008; Nyola and Hunte 2008). Interestingly, most of the water molecules are stabilized by interactions with Cc 1 and not with Cc. The hydration pattern of Cc 1 rearranges significantly upon Cc binding, resulting in a single water-molecule-mediated intermolecular hydrogen bond at the Cc 1–Cc interface (Solmaz and Hunte 2008). In contrast, comparable ET complexes such as CcP–Cc show three interface water molecules that establish hydrogen bonds between both proteins (see below; Pelletier and Kraut 1992). The lack of salt bridges and hydrogen bonds, along with the high solvation of the interface, make the Cc 1–Cc interaction specifically transient and the lifetime of the complex relatively short. This correlates with the mobility mismatch of the positively charged interacting side chains of Cc, which may further contribute to the undocking process (Solmaz and Hunte 2008). Of interest is the 1:1 binding stoichiometry of bc 1–Cc complex. Cc binds specifically only to one of the two possible recognition sites of the dimeric bc 1 (Lange and Hunte 2002). This indicates that bc 1 might be able to reduce Cc with the second functional unit not being active, thereby supporting a sequential or independent mode. Recently, it has been demonstrated that electrons move freely within and between monomers of bc 1, acting as a molecular-scale bus bar that increases the effective diffusion for Cc (Świerczek et al. 2010). 7 Post-translational modifications of tyrosines from Cc can modulate the binding to Cc 1. R(N)OS promotes tyrosine nitration of proteins, with Cc being the main target in mitochondria. The nitration of two out of five tyrosine residues—at positions 46 and 48—turns Cc into a high-spin species without significant changes in its secondary structure (Díaz-Moreno et al. 2011c), a finding that may explain the drop of ca. 100 mV in the midpoint reduction potential value of nCc forms (Rodríguez-Roldán et al. 2008). Thus, cellular respiration is partially disrupted by nitration because Cc is no longer isopotential with Cc 1, and it becomes unable to accept electrons from the cytochrome bc 1 complex. Under (nitro)oxidative stress, the excess in R(N)OS yielded from the first complexes of the respiratory chain could lead to a positive nitration-driven feedback cycle, with cytochrome bc 1 promoting the increase in R(N)OS and nCc levels. On the other hand, phosphorylation of Cc-Tyr48 induces significant modifications in the heme environment without major structural change, namely an 80-mV drop in the midpoint reduction potential value and inhibition of the electron flux between complexes III and IV (Yu et al. 2008; Pecina et al. 2010; García-Heredia et al. 2011). The cytochrome c and cytochrome c oxidase complex Cytochrome c oxidase is the last electron acceptor of the mitochondrial respiratory chain and catalyzes the reduction of molecular oxygen to water, coupling the free energy of water formation to proton translocation across the membrane (Papa et al. 2004; Fig. 3). Eukaryotic CcO contains 13 subunits, each different from the other, the catalytic core of the enzyme being formed by the three largest subunits: Cox1, Cox2 and Cox3. Cox1 contains one copper ion (termed CuB), whereas a binuclear copper binding site, named CuA, is located in Cox2 (Tsukihara et al. 1996). The delivery of copper to the CuA site during the process of mitochondrial CcO assembly is carried out by the Sco (Synthesis of cytochrome c oxidase) protein, which, in turn, receives copper from the chaperone Cox17 (Banci et al. 2011a). In eukaryotic organisms, in particular, the Sco protein develops additional functions to the CcO assembly, including mitochondrial signaling and regulation of copper homeostasis. Structural information for the eukaryotic Cc–CcO complex has been recently reported (Sakamoto et al. 2011). One of the most interesting features is that the adduct is mainly stabilized by hydrophobic interactions between partners, which are mediated by the hydrophobic heme periphery and adjacent hydrophobic amino acid residues of Cc. Such interactions place the two redox centers of Cc and CcO in close proximity. In addition, charged residues near the hydrophobic core refine the orientation of Cc with respect to CcO in order to facilitate the strictly controlled ET reaction. Unexpectedly, negatively charged residues, along with Lys residues, are included in the interaction surface with the exception of Glu16 and Lys72 whose nuclear magnetic resonance (NMR) signals show no substantial perturbations although they surround the hydrophobic core and seem to be involved in the interaction with CcO (Roberts and Pique 1999). The CuA site of CcO, which acts as an electron entry site (Yoshikawa et al. 1998), is surrounded by an exposed hydrophobic cluster containing aromatic residues—Trp104, Tyr105, Tyr121 and Phe206. The analysis of the intermolecular contacts found in Cc points out that the exposed hydrophobic Cc heme edge, as well as its adjacent hydrophobic residues, can interact with the 8 hydrophobic cluster at the CuA site to form an ensemble that is competent for ET. Needless to say that the adducts among ET partners, as in the Cc–CcO complex, are usually dynamic and transient to guarantee a high turnover (Prudêncio and Ubbink 2004). This dynamic model is compatible with the two-step docking mechanism, previously proposed for the Cc 1–Cc interaction. Cc and CcO are brought in close proximity and pre-oriented by long-range electrostatic forces responsible for the initial encounter complex in which different orientations have approximately the same energy. Hydrophobic interactions, acting at short distances, account for the formation of the more specific active complex upon rearrangement of the two partners (Prudêncio and Ubbink 2004). In the prokaryotic counterpart between cytochrome c 552 (Cc 552) and the CuA subunit of CcO from Paracoccus denitrificants, hydrophobic contacts are limited to few residues from both proteins (Bertini et al. 2005), which can explain that the adduct remains in a highly dynamic ensemble of different orientations rather than in a productive single orientation. Interestingly, some of these conformations are active for ET, but others are not. The binding affinity of Cc and CcO is essentially independent of the oxidation state of Cc. However, modifications in the polarity of the Cc heme environment due to the relevant conformational change of Cc Lys13 towards its oxidized heme upon CcO binding decrease its affinity for CcO to be replaced later by the reduced Cc (Sakamoto et al. 2011). Cc experiences multiple encounters with its two respiratory partners: bc 1 and CcO. However, only few of them are productive. The number of encounters, and hence the ET rate, may be increased by the formation of a bc 1–Cc–CcO supercomplex (Heinemeyer et al. 2007) and/or by increasing the concentration of the individual partners in the IMS. Nevertheless, recent cryoelectron tomography data on the respirasome—multisubunit supercomplex of the respiratory chain containing complexes I, III and IV—reveals that the Cc binding sites in bc 1 and CcO are placed further apart (Dudkina et al. 2011). Tyrosine nitration significantly decreases the reactivity of Cc towards CcO, impairs the respiratory ET process and enhances R(N)OS production by mitochondria (Rodríguez-Roldán et al. 2008). It is remarkable that the dependence of the ET rate constant upon CcO concentration reaches a saturation plateau with wild-type (WT) Cc, but linearly increases with n-Cc. This suggests that the mechanism of Cc–CcO binding includes the formation of a transient complex before ET, whereas the interaction between n-Cc and CcO follows a collisional mechanism (Rodríguez-Roldán et al. 2008). The drastically diminished reactivity of Cc upon nitration is just contrary to what should be expected from the higher driving force of the reaction resulting from the lowering in reduction potential of n-Cc. However, such a decrease is a specific effect of nitration on the interaction with CcO, since the effect on the reactivity of the n-Cc toward other electron acceptors, such as flavins, is negligible (RodríguezRoldán et al. 2008). Cc tyrosine phosphorylation also leads to partial inhibition of the reaction with CcO. While Tyr97-phosphorylated Cc yields sigmoidal kinetics of CcO reduction, hyperbolic responses are obtained with Tyr48-phosphorylated and WT Cc species (Lee et al. 2006; Yu et al. 2008). However, the highest turnover is reduced to half when Cc becomes phosphorylated at Tyr48 (Yu et al. 2008). 9 Cytochrome c acts as an electron shuttle between the outer and inner mitochondrial membranes: the cytochrome b 5 and cytochrome c complex Cytochrome b 5 is located at the inner surface of the OMM and transfers electrons from the cytosolic NADH—using the mitochondrial outer membrane-bound form of NADH-dependent cytochrome b 5 Reductase (Cb 5R)—to Cc in the IMS (Bernardi and Azzone 1981; Fig. 2). Thus, reduced Cc interferes with the respiratory chain, delivering electrons to CcO, as corroborated by the complete inhibition of KCN-mediated CcO (Matlib and O’Brien 1976; Bernardi and Azzone 1981). The demonstration that Cc communicates OMM and IMM implies that the oxidation– reduction state of the cytosolic NADH system and mitochondrial oxidation–reduction reactions are closely dependent on each other. The Cb 5–Cc docking reveals an ensemble of orientations assumed by the hemeproteins rather than a single, well-defined conformation (Banci et al. 2003; Volkov et al. 2005; Deep et al. 2005). This is based on the magnitude of several NMR parameters, such as the chemical-shift perturbation for Cb 5 and Cc, which average to zero when proteins adopt a multitude of transient orientations within the complex. These orientations have nearly equal energies and are in fast exchange on the NMR time scale, thus resulting in an average of the chemical-shift perturbations over all orientations. In combination with the absence of close contacts and extensive desolvation, this explains the observed small chemical-shift perturbations due binding within the complex. In addition, the complex is electrostatically guided to guarantee the recognition and preorientation of the two proteins, as inferred from site-directed mutagenesis studies (Sun et al. 1999, 2001) and ionic strength titrations (Volkov et al. 2005). Whereas Cc uses a single surface formed by the positively charged lysine residues surrounding the heme crevice for interaction (Fig. 1), Cb 5 mainly requires its negatively charged region around the solvent-exposed heme edge. Controversial data regarding the stoichiometry for this complex vary depending on the source of Cc. In the complex between yeast Cc and bovine Cb 5, the concept of a ternary Cb 5–(Cc)2 complex, with an stoichiometry of 2:1, is supported by the two Cb 5 surfaces involved in the interaction (Banci et al. 2003; Volkov et al. 2005). In one area Cc binds to the front side of Cb 5 (“head-on” orientation), in the other Cc interacts at the left side of Cb 5 (“side-on” orientation). Both docking solutions satisfy all experimental restraints in accordance with the dynamic nature of the complex. Among these adducts, only a few of them are ET active, even though they are not the most favorable for binding. In fact, the docked structures with the “head-on” orientation might represent ET-productive complexes, since this is the conformation in which the heme-to-heme distance is optimum for ET. In contrast, those Cb 5–Cc complexes formed by horse Cc occur in 1:1 stoichiometry and predominantly exist in a favored orientation (Shao et al. 2003; Deep et al. 2005). It is worth mentioning that the binding surface used by Cc to interact with Cb 5 is independent of the source where it comes from (Deep et al. 2005). The driving force for the Cb 5–Cc ET reaction may be significantly blocked upon nitration or phosphorylation of Cc as they both make the reduction potential drop. However, some 16 Exclusively in plants, l-GAlactono-1,4-Lactone DeHydrogenase (GALDH) is another integral membrane-bound target of Cc from which electrons shuttle into the ET chain via Cc (Leferink et al. 2008). Recent data do suggest that GALDH catalyzes the l-ascorbate production with the concomitant reduction of Cc (Leferink et al. 2009). Neither the GALDH–Cc complex structure nor the effects of post-translationally modified Cc have been elucidated yet. The non-physiological complexes of Cc Non-physiological complexes between redox proteins have also been studied. They exhibit fast ET and affinity constants, within the same range as the physiological partners (Fig. 1; Table 1). Among the non-physiological complexes of Cc reported in the literature, the one formed by adrenodoxin (Adx) and Cc has been widely analyzed because Cc serves as a model for cytochrome P450 (Fig. 2). Actually, Adx is involved in steroid hormone biosynthesis shuttling electrons between NADPH-dependent adrenodoxin reductase (AdR) and several cytochromes P450 inside the mitochondria (Fig. 3). As the one-electron transfer from (AdR) to Adx is the slowest, rate-limiting step (Lambeth and Kamin 1979), the fact that Adx and Cc show fast ET suggests that a complex is formed. The Adx–Cc complex is entirely dynamic and can be considered as a pure encounter complex dominated by electrostatics—between four acidic patches in Adx and the positively charged area surrounding the heme group of Cc—and, nevertheless, active in ET (Worrall et al. 2003; Xu et al. 2008). The apparent lack of surface matching on both partners allows more than one orientation to be sampled (Fig. 1). Authors propose that these orientations most likely have similar energies and are in fast exchange in the NMR scale, thus resulting in averaging of several NMR parameters over all orientations. The absence of close contacts and extensive desolvation would explain the small signal perturbations upon complex formation, which suggests that the two metalloproteins sample other surfaces areas away from the predominant binding sites. Interestingly, the dipole moment of Adx does not seem to be involved in electrostatic steering among the proteins in the redox chain, including the nonphysiological target Cc, in contrast to what has been widely accepted (Hannemann et al. 2009). Mitochondrial scaffold of Cc: the homeostatic-to-apoptotic transition The mitochondrial signalosome of Cc is constituted by metabolic pathways that are closely linked to each other (Fig. 3). Under homeostasis, Cc carries out the ET flux of the mitochondrial respiration in a controlled manner and ensures healthy mitochondrial membrane potentials. Eventually, R(N)OS can be accidentally generated by the own activity of the mitochondrial respiratory chain. Such radical species can efficiently be neutralized by enzymes like manganese superoxide dismutase or by the Cc–CcP complex, which functions as a R(N)OS scavenger. Simultaneously, Cc can serve as the nexus between the cytosolic NADH-generating system and respiratory chain at the level of CcO. Indeed, the inactivation state of p66Shc is accompanied by the overexpression of Bcl-xL, which sequesters Cc from the cytosol to avoid PCD in the absence of stress. Under apoptotic conditions, Cc allows the respiration chain to work at the maximum rate up to yield an excess 17 in R(N)OS. This promotes oxidation and nitration of Cc, which, in turn, might accelerate R(N)OS generation by transient ET to p66Shc, previously activated upon apoptosis. It is thus plausible to propose that the reduced Cc pool resulting from the Cb 5, Erv1 and Sco pathways could also contribute to the increase in R(N)OS production, although the ET fluxes may be disrupted by post-translationally modified Cc, while CcP is not efficient enough to neutralize extra R(N)OS. R(N)OSs are indeed responsible for hyperpolarization of the mitochondrial membranes by opening the transmembrane pore. At this stage, the peroxidation of CL catalyzed by Cc— especially the nitrated Cc species—is essential for the collapse of the mitochondrial membrane. Then, Cc is released into the cytosol where it stably assembles the apoptosome because of the lack of Bcl-xL so as to amplify the PCD signal by activating the caspases cascade. Conclusions and outlook Puzzling the Cc-controlled complex network out in the cellular environment requires knowledge about contacts between partners and, eventually, the discovery of new targets. Figure 1 shows the Cc residues forming the interface with the Cc targets for which structural information is available. The interaction surface offered by Cc shifts from the area surrounding the heme crevice for those ET-active Cc complexes (Bertini et al. 2011b) to the opposite molecule side when the ET is not necessary. Therefore, the Cc mappings that comprise mitochondrial contacts are almost identical, since they include residues close to the heme edge. Still post-translational modifications, such as nitration and phosphorylation, of Cc need to be explored in the context of the hemeprotein and its interaction network with mitochondrial targets. Because of the central role of Cc in lifespan regulation and cell fate by controlling both respiration and apoptosis, a deeper understanding of the signaling mechanisms involving Cc, as well as their regulation by post-translational modifications of Cc—phosphorylation and nitration—is essential. In addition, the identification of interfaces between Cc and its counterparts will enable the development and design of novel therapies. Other promising approaches are aimed at identifying drugs that inhibits the action of pro-survival—potential oncogenes—and pro-apoptotic—aging-related diseases—partners of Cc. Acknowledgments The authors wish to thank Jonathan Martínez-Fábregas for helpful advice and critical reading of the manuscript. This work was funded by the Spanish Ministry of Science and Innovation (BFU2009-07190) and the Andalusian Government (BIO198 and P08-CVI-3876). 18 References Acehan D, Jiang X, Morgan DG, Heuser JE, Wang X, Akey CW (2002) Three-dimensional structure of the apoptosome: implications for assembly, procaspase-9 binding, and activation. Mol Cell 9:423–432 Acin-Perez R, Hoyos B, Gong J, Vinogradov V, Fischman DA, Leitges M, Borhan B, Starkov A, Manfredi G, Hammerling U (2010) Regulation of intermediary metabolism by the PKCδ signalosome in mitochondria. FASEB J 24:5033–5042 Banci I, Bertini I, Felli IC, Krippahl L, Kubicek K, Moura JJG, Rosato A (2003) A further investigation of the cytochrome b 5–cytochrome c complex. J Biol Inorg Chem 8:777–786 Banci L, Bertini I, Cavallaro G, Ciofi-Baffoni S (2011a) Seeking the determinants of the elusive functions of Sco proteins. FEBS J 278:2244–2262 Banci L, Bertini I, Ciofi-Baffoni S, Kozyreva T, Mori M, Wang S (2011b) Sco proteins are involved in electron transfer processes. J Biol Inorg Chem 16:391–403 Bashir Q, Volkov AN, Ullmann GM, Ubbink M (2010) Visualization of the encounter ensemble of the transient electron transfer complex of cytochrome c and cytochrome c peroxidase. J Am Chem Soc 132:241–247 Bashir Q, Scanu S, Ubbink M (2011) Dynamics in electron transfer protein complexes. FEBS J 278:1391–1400 Batthyány C, Souza JM, Durán R, Cassina A, Cerveñansky C, Radi R (2005) Time course and site(s) of cytochrome c tyrosine nitration by peroxynitrite. Biochemistry 44:8038–8046 Bayir H, Fadeel B, Palladino MJ, Witasp E, Kurnikov IV, Tyurina YY, Tyurin VA, Amoscato AA, Jiang J, Kochanek PM, DeKosky ST, Greenberger JS, Shvedova AA, Kagan VE (2006) Apoptotic interactions of cytochrome c: redox flirting with anionic phospholipids within and outside of mitochondria. Biochim Biophys Acta 1757:648–659 Belikova NA, Vladimirov YA, Osipov AN, Kapralov AA, Tyurin VA, Potapovich MV, Basova LV, Peterson J, Kurnikov IV, Kagan VE (2006) Peroxidase activity and structural transitions of cytochrome c bound to cardiolipin-containing membranes. Biochemistry 45:4998–5009 Bernardi P, Azzone GF (1981) Cytochrome c as an electron shuttle between the outer and inner mitochondrial membrane. J Biol Chem 256:7187–7192 Berry EA, Guergova-Kuras M, Huang LS, Crofts AR (2000) Structure and function of cytochrome bc complexes. Annu Rev Biochem 69:1005–1075 Bertini I, Cavallaro G, Rosato R (2005) A structural model for the adduct between cytochrome c and cytochrome c oxidase. J Biol Inorg Chem 10:613–624 Bertini I, Chevance S, Del Conte R, Lalli D, Turano P (2011a) The anti-apoptotic Bcl-xL protein, a new piece in the puzzle of cytochrome c interactome. PLos One. doi:10.1371/journal.pone.0018329 19 Bertini I, Cavallaro G, Rosato A (2011b) Principles and patterns in the interaction between mono-heme cytochrome c and its partners in electron transfer processes. Metallomics. doi:10.1039/c0mt00108b Bihlmaier K, Mesecke N, Terzyiska N, Bien M, Hell K, Herrmann JM (2007) The disulfide relay system of mitochondria is connected to the respiratory chain. J Cell Biol 179:389–395 Cai J, Yang J, Jones DP (1998) Mitochondrial control of apoptosis: the role of cytochrome c. Biochim Biophys Acta 1366:139–149 Cassina AM, Hodara R, Souza JM, Thomson L, Castro L, Ischiropoulos H, Freeman BA, Radi R (2000) Cytochrome c nitration by peroxynitrite. J Biol Chem 275:21409–21415 Chance B, Sies H, Boveris A (1979) Hydroperoxide metabolism in mammalian organs. Physiol Rev 59:527–605 Chen Q, Vazquez EJ, Moghaddas S, Hoppel CL, Lesnefsky EJ (2003) Production of reactive oxygen species by mitochondria: central role of complex III. J Biol Chem 278:36027–36031 Chen YR, Chen CL, Chen W, Zweier JL, Augusto O, Radi R, Mason RP (2004) Formation of protein tyrosine ortho-semiquinone radical and nitrotyrosine from cytochrome c-derived tyrosyl radical. J Biol Chem 279:18054–18062 Cortese J, Voglino AL, Hackenbrock CR (1995) Persistence of cytochrome c binding to membranes at physiological mitochondrial intermembrane space ionic strength. Biochim Biophys Acta 1228:216–228 Dabir DV, Leverich EP, Kim S-K, Tsai FD, Hirasawa M, Knaff DB, Koehler CM (2007) A role for cytochrome c and cytochrome c peroxidase in electron shuttling from Erv1. EMBO J 26:4801– 4811 Daum G (1985) Lipids of mitochondria. Biochim Biophys Acta 822:1–42 Deep S, Im S-C, Zuiderweg ERP, Waskell L (2005) Characterization and calculation of a cytochrome c–cytochrome b 5 complex using NMR data. Biochemistry 44:10654–10668 Demel RA, Jordi W, Lambrechts H, van Damme H, Hovius R, de Kruijff B (1989) Differential interactions of apoand holocytochrome c with acidic membrane lipids in model systems and the implications for their import into mitochondria. J Biol Chem 264:3988–3997 Díaz-Moreno I, De La Rosa MA (2011a) Transient interactions between biomolecules. Eur Biophys J. doi:10.1007/s00249-011-0728-x Díaz-Moreno I, De La Rosa MA (2011b) Transient interactions in metalloproteins. FEBS J. doi:10.1111/j.1742-4658.2011.08065.x Díaz-Moreno I, García-Heredia JM, Díaz-Quintana A, Teixeira M, De La Rosa MA (2011c) Nitration of tyrosines 46 and 48 induces the specific degradation of cytochrome c upon change of the heme iron state to high-spin. Biochem Biophys Acta Bioenerg. doi:10.1016/j.bbabio.2011.09.012 20 Dudkina NV, Kudryashev M, Stahlberg H, Boekema EJ (2011) Interaction of complexes I, III, and IV within the bovine respirasome by single particle cryoelectron tomography. Proc Natl Acad Sci USA 108:15196–15200 Durham B, Fairris JL, McLean M, Millett F, Scott JR, Sligar SG, Willie A (1995) Electron transfer from cytochrome b 5 to cytochrome c. J Bioenerg Biomembr 27:331–340 Eble KS, Coleman WB, Hantgan RR, Cunningham CC (1990) Tightly associated cardiolipin in the bovine heart mitochondrial ATP synthase as analyzed by 31P nuclear magnetic resonance spectroscopy. J Biol Chem 265:19434–19440 Engstrom G, Rajagukguk R, Saunders AJ, Patel CN, Rajagukguk S, Merbitz-Zahradnik T, Xiao K, Pielak GJ, Trumpower B, Yu C-A, Yu L, Durham B, Millet F (2003) Design of a ruthenium-labeled cytochrome c derivative to study electron transfer with the cytochrome bc 1 complex. Biochemistry 42:2816–2824 Fry M, Green DE (1981) Cardiolipin requirement for electron transfer in complex I and III of the mitochondrial respiratory chain. J Biol Chem 256:1874–1880 García-Heredia JM, Díaz-Moreno I, Nieto PM, Orzáez M, Kocanis S, Teixeira M, Pérez-Payá E, Díaz-Quintana A, De la Rosa MA (2010) Nitration of tyrosine 74 prevents human cytochrome c to play a key role in apoptosis signaling by blocking caspase-9 activation. Biochem Biophys Acta Bioenerg 1797:981–993 García-Heredia JM, Díaz-Quintana A, Salzano M, Orzáez M, Pérez-Payá E, Teixeira M, De la Rosa MA, Díaz-Moreno I (2011) Tyrosine phosphorylation turns alkaline transition into a biologically relevant process and makes human cytochrome c behave as an anti-apoptotic switch. J Biol Inorg Chem. doi:10.1007/s00775-011-0804-9 Giles SS, Perfect JR, Cox GM (2005) Cytochrome c peroxidase contributes to the antioxidant defense of Cryptococcus neoformans. Fungal Genet Biol 42:20–29 Giorgio M, Migliaccio E, Orsini F, Paolucci D, Moroni M, Contursi C, Pelliccia G, Luzi L, Minucci S, Marcaccio M, Pinton P, Rizzuto R, Bernardi P, Paolucci F, Pelicci PG (2005) Electron transfer between cytochrome c and p66Shc generates reactive oxygen species that trigger mitochondrial apoptosis. Cell 122:221–233 Gomez B Jr, Robinson NC (1999) Phospholipase digestion of bound cardiolipin reversibly inactivates bovine cytochrome bc 1. Biochemistry 38:9031–9038 Haines TH, Dencher NA (2002) Cardiolipin: a proton trap for oxidative phosphorylation. FEBS Lett 528:35–39 Hannemann F, Guyot A, Zöllner A, Müller JJ, Heinemann U, Bernhardt R (2009) The dipole moment of the electron carrier adrenodoxin is not critical for redox partner interaction and electron transfer. J Inorg Biochem 103:997–1004 Heinemeyer J, Braun H-P, Boekema EJ, Kouřil R (2007) A structural model of the cytochrome c reductase/oxidase supercomplex from yeast mitochondria. J Biol Chem 282:12240–12248 21 Hüttemann M, Pecina P, Rainbolt M, Sanderson TH, Kagan VE, Samavati L, Doan JW, Lee I (2011a) The multiple functions of cytochrome c and their regulation in life and death decisions of the mammalian cell: from respiration to apoptosis. Mitochondrion 11:369–381 Hüttemann M, Helling S, Sanderson TH, Sinkler C, Samavati L, Mahapatra G, Varughese A, Lu G, Liu J, Ramzan R, Vogt S, Grossman LI, Doan JW, Marcus K, Lee I (2011b) Regulation of mitochondrial respiration and apoptosis through cell signaling: cytochrome c oxidase and cytochrome c in ischemia/reperfusion injury and inflammation. Biochim Biophys Acta. doi:10.1016/j.bbabio.2011.07.001 Jeng WY, Chen CY, Chang HC, Chuang WJ (2002) Expression and characterization of recombinant human cytochrome c in E. coli. J Bioenerg Biomembr 34:423–431 Jiang H, English AM (2006) Phenotypic analysis of the ccp1Δ and ccp1Δ-ccp1 W191F mutant strains of Saccharomyces cerevisiae indicates that cytochrome c peroxidase functions in oxidative-stress signaling. J Inorg Biochem 100:1996–2008 Kagan VE, Borisenko GG, Tyurina YY, Tyurin VA, Jiang J, Potapovich AI, Kini V, Amoscato AA, Fujii Y (2004) Oxidative lipidomics of apoptosis: redox catalytic interactions of cytochrome c with cardiolipin and phosphatidylserine. Free Rad Biol Med 37:1963–1985 Kagan VE, Tyurina YY, Batir H, Chu CT, Kapralov AA, Vlasova II, Belikova NA, Tyurin VA, Amoscato A, Epperly M, Greenberger J, DeKosky S, Shvedova AA, Jiang J (2006) The “proapoptotic genies” get out of mitochondria: oxidative lipidomics and redox activity of cytochrome c/cardiolipin complexes. Chem Biochem Interact 163:15–28 Kalanxhi E, Wallace CJ (2007) Cytochrome c impaled: investigation of the extended lipid anchorage of a soluble protein to mitochondrial membrane models. Biochem J 407:179–187 Kapralov AA, Yanamala N, Tyurina YY, Castro L, Samhan-Arias AK, Vladimirov YA, Maeda A, Mylnikov D, Demicheli V, Tortora V, Klein-Seetharaman J, Radi R, Kagan VE (2011) Topography of tyrosine residues and their involvement in peroxidation of polyunsaturated cardiolipin in cytochrome c/cardiolipin peroxidase complexes. Biochim Biophys Acta 1808:2147–2155 Kong SK, Yim MB, Stadtman ER, Chock PB (1996) Peroxynitrite disables the tyrosine phosphorylation regulatory mechanism: lymphocyte-specific tyrosine kinase fails to phosphorylate nitrated cdc2(6–20)NH2 peptide. Proc Natl Acad Sci USA 93:33777–33782 Lambeth JD, Kamin H (1979) Adrenodoxin reductase: adrenodoxin complex flavin to iron-sulfur transfer as the rate-limiting step in the NADPH-cytochrome c reductase reaction. J Biol Chem 254:2766–2774 Lange C, Hunte C (2002) Crystal structure of the yeast cytochrome bc 1 complex with its bound substrate cytochrome c. Proc Natl Acad Sci USA 99:2800–2805 Lee I, Salomon AR, Yu K, Doan JW, Grossman LI, Hüttemann M (2006) New prospects for an old enzyme: mammalian cytochrome c is tyrosine-phosphorylated in vivo. Biochemistry 47:9121– 9128 22 Leesch VW, Bujous J, Mauk AG, Hoffman BM (2000) Cytochrome c peroxidase-cytochrome c complex: locating the second binding domain on cytochrome c peroxidase with site-directed mutagenesis. Biochemistry 39:10132–10139 Leferink NGH, van der Berg WAM, van Berkel WJH (2008) l-Galactono-γ-lactone dehydrogenase from Arabidopsis thaliana, a flavoprotein involved in vitamin C biosynthesis. FEBS J 275:713–726 Leferink NGH, Fraaije MW, Joosten H-J, Schaap PJ, Mattevi A, van Berkel WJH (2009) Identification of a gatekeeper residue that prevents dehydrogenases from acting as oxidases. J Biol Chem 284:4392–4397 Louie GV, Brayer GD (1990) High-resolution refinement of yeast iso-1-cytochrome c and comparisons with other eukaryotic cytochromes c. J Mol Biol 214:527–555 Matlib MA, O’Brien PJ (1976) Properties of rat liver mitochondria with intermembrane cytochrome c. Arch Biochem Biophys 173:27–33 Mei H, Geren L, Miller MA, Durham B, Millett F (2002) Role of the low-affinity binding site in electron transfer from cytochrome c to cytochrome c peroxidase. Biochemistry 41:3968–3976 Migliaccio E, Mele S, Salcini AE, Pelicci G, Lai K-MV, Superti-Furga G, Pawson T, Di Fiore PP, Lanfrancone L, Pelicci PG (1997) Opposite effects of the p52Shc/p46Shc and p66Shc splicing isoforms on the EGF receptor-MAP kinase-fos signalling pathway. EMBO J 16:706–716 Migliaccio E, Giorgio M, Mele S, Pelicci G, Reboldi P, Pandolfi PP, Lanfrancone L, Pelicci PG (1999) The p66Shc adaptor protein controls oxidative stress response and life span in mammals. Nature 402:309–313 Millett F, Miller MA, Geren L, Durham B (1995) Electron transfer between cytochrome c and cytochrome c peroxidase. J Bioenerg Biomembr 27:341–351 Moore GR, Pettigrew GW (1990) Cytochromes c. Evolutionary, structural, and physicochemical aspects. Springer, New York Mustonen P, Virtanen JA, Somerharju PJ, Kinnunen PKJ (1987) Binding of cytochrome c to liposomes as revealed by the quenching of fluorescence from pyrene-labeled phospholipids. Biochemistry 26:2991–2997 Nyola A, Hunte C (2008) A structural analysis of the transient interaction between the cytochrome bc 1 complex and its substrate cytochrome c. Biochem Soc Trans 36:981–985 Orrenius S (2007) Reactive oxygen species in mitochondria-mediated cell death. Drugs Metab Res 39:443–455 Orsini F, Migliaccio E, Moroni M, Contursi C, Raker VA, Piccini D, Martin-Padura I, Pelliccia G, Trinei M, Bono M, Puri C, Tacchetti C, Ferrini M, Mannucci R, Nicoletti I, Lanfrancone L, Giorgio M, Pelicci PG (2004) The life span determinant p66Shc localizes to mitochondria where it associates with mitochondrial heat shock protein 70 and regulates trans-membrane potential. J Biol Chem 279:25689–25695 23 Ott M, Robertson J, Gogvadze V, Zhivotovsky B, Orrenius S (2002) Cytochrome c release from mitochondria proceeds by a two-step process. Proc Natl Acad Sci USA 99:1259–1263 Oursler MJ, Bradley EW, Elfering SL, Giulivi C (2005) Native, not nitrated, cytochrome c and mitochondria-derived hydrogen peroxide drive osteoclast apoptosis. Am J Physiol Cell Physiol 288:156–168 Papa S, Capitanio N, Capitanio G (2004) A cooperative model for proton pumping in cytochrome c oxidase. Biochem Biophys Acta 1655:353–364 Pearl NM, Jacobson T, Arisa M, Vitello LB, Erman JE (2007) Effect of single-site charge-reversal mutations on the catalytic properties of yeast cytochrome c peroxidase: mutations near the high-affinity cytochrome c binding site. Biochemistry 46:8263–8272 Pearl NM, Jacobson T, Meyen C, Clementz AG, Ok EY, Choi E, Wilson K, Vitello LB, Erman JE (2008) Effect of single-site charge-reversal mutations on the catalytic properties of yeast cytochrome c peroxidase: evidence for a single, catalytically active, cytochrome c binding domain. Biochemistry 47:2766–2775 Pecina P, Borisenko GG, Belikova NA, Tyurina Y, Pecinova A, Lee I, Samhan-Arias AK, Przyklenk K, Kagan VE, Huttemann M (2010) Phosphomimetic substitution of cytochrome c tyrosine 48 decreases respiration and binding to cardiolipin and abolishes ability to trigger downstream caspase activation. Biochemistry 49:6705–6710 Pelicci G, Lanfrancone L, Grignani F, McGlade J, Cavallo F, Forni G, Nicoletti I, Grignani F, Pawson T, Pelicci PG (1992) A novel transforming protein (SHC) with an SH2 domain is implicated in mitogenic signal transduction. Cell 70:93–104 Pellegrini M, Pacini S, Baldari CT (2005) p66Shc: the apoptotic side of Shc proteins. Apoptosis 10:13–18 Pelletier H, Kraut J (1992) Crystal structure of a complex between electron transfer partners, cytochrome c peroxidase and cytochrome c. Science 258:1748–1755 Pereverzev MO, Vygodina TV, Konstantinov AA, Skulachev VP (2003) Cytochrome c, an ideal antioxidant. Biochem Soc Trans 31:1312–1315 Pettersen EF, Goddard TD, Huang CC, Couch GS, Greenblatt DM, Meng EC, Ferrin TE (2004) UCSF Chimera—a visualization system for exploratory research and analysis. J Comput Chem 25:1605–1612 Pettigrew GW, Prazeres S, Costa C, Palma N, Krippahl L, Moura I, Moura JJG (1999) The structure of an electron transfer complex containing a cytochrome c and a peroxidase. J Biol Chem 274:11383–11389 Pinton P, Rimessi A, Marchi S, Orsini F, Migliaccio E, Giorgio M, Contursi C, Minucci S, Mantovani F, Wieckowski MR, Del Sal G, Pelicci PG, Rizzuto R (2007) Protein kinase C β and prolyl isomerase 1 regulate mitochondrial effects of the life-span determinant p66Shc. Science 315:659–663 24 Poulos TL, Freer ST, Alden RA, Edwards SJ, Skoglands U, Takio K, Eriksson B, Xuong N-H, Yonetani T, Kraut J (1980) The crystal structure of cytochrome c peroxidase. J Biol Chem 255:575–580 Prudêncio M, Ubbink M (2004) Transient complexes of redox proteins: structural and dynamic details from NMR studies. J Mol Recognit 17:524–539 Reincke B, Perez C, Pristovsek P, Lucke C, Ludwig C, Lohr F, Rogov VV, Ludwig B, Ruterjans H (2001) Solution structure and dynamics of the functional domain of Paracoccus denitrificans cytochrome c(552) in both redox states. Biochemistry 40:12312–12320 Riemer J, Fischer M, Hermann JM (2011) Oxidation-driven protein import into mitochondria: insights and blind spots. Biochim Biophys Acta 1808:981–989 Roberts VA, Pique ME (1999) Definition of the interaction domain for cytochrome c on cytochrome c oxidase. J Biol Chem 274:38051–38060 Robinson NC (1993) Functional binding of cardiolipin to cytochrome c oxidase. J Bioenerg Biomembr 25:153–163 Rödiger A, Baudisch B, Langner U, Klösgen RB (2011) Dual targeting of a mitochondrial protein: the case study of cytochrome c 1. Mol Plant 4:679–687 Rodríguez-Roldán V, García-Heredia JM, Navarro JA, De la Rosa MA, Hervás M (2008) Effect of nitration on the physicochemical and kinetic features of wild-type and mono-tyrosine mutants of human respiratory cytochrome c. Biochemistry 47:12371–12379 Rytömaa M, Mustonen P, Kinnunen PK (1992) Reversible, nonionic, and pH-dependent association of cytochrome c with cardiolipin-phosphatidylcholine liposomes. J Biol Chem 267:22243–22248 Sakamoto K, Kamiya M, Imai M, Shinzawa-Itoh K, Uchida T, Kawano K, Yoshikawa S, Ishimori K (2011) NMR basis for interprotein electron transfer gating between cytochrome c and cytochrome c oxidase. Proc Natl Acad Sci USA 108:12271–12276 Saraste M (1999) Oxidative phosphorylation at the fin de siecle. Science 283:1488–1493 Schug ZT, Gottlieb E (2009) Cardiolipin acts as a mitochondrial signaling platform to launch apoptosis. Biochim Biophys Acta 1788:2022–2031 Scorolli L, Meduri A, Morara M, Scalinci SZ, Meduri RA (2007) Effect of cytochrome c peroxidase on the corneal epithelial healing process after excimer laser photoablation in transgenic mice. Eur Surg Res 39:82–87 Shao W, Im S-C, Zuiderweg ERP, Waskell L (2003) Mapping the binding interface of the cytochrome b 5–cytochrome c complex by nuclear magnetic resonance. Biochemistry 42:14774–14784 25 Shidoji Y, Hayashi K, Komura S, Ohishi N, Yagi K (1999) Loss of molecular interaction between cytochrome c and cardiolipin due to lipid peroxidation. Biochem Biophys Res Commun 264:343–347 Sinibaldi F, Fiorucci L, Patriarca A, Lauceri R, Ferri T, Coletta M, Santucci R (2008) Insights into cytochrome c-cardiolipin interaction. Role played by ionic strength. Biochemistry 47:6928– 6935 Sinibaldi F, Howes BD, Piro MC, Polticelli F, Bombelli C, Ferri T, Coletta M, Smulevich G, Santucci R (2010) Extended cardiolipin anchorage to cytochrome c: a model for proteinmitochondrial membrane binding. J Biol Inorg Chem 15:689–700 Solmaz SRN, Hunte C (2008) Structure of complex III with bound cytochrome c in reduced state and definition of a minimal core interface for electron transfer. J Biol Chem 283:17542–17549 Stemp EDA, Hoffman BM (1993) Cytochrome c peroxidase binds two molecules of cytochrome c: evidence for a low-affinity, electron transfer-active site on cytochrome c peroxidase. Biochemistry 32:10848–10865 Stepanov G, Gnedenko O, Mol’nar A, Ivanov A, Vladimirov Y, Osipov A (2009) Evaluation of cytochrome c affinity to anionic phospholipids by means of surface plasmon resonance. FEBS Lett 583:97–100 Sun Y-L, Wang Y-H, Yan M-M, Sun B-Y, Xie Y, Huang Z-X, Jiang S-K, Wu H-M (1999) Structure, interaction and electron transfer between cytochrome b 5, its E44A and/or E56A mutants and cytochrome c. J Mol Biol 285:347–359 Sun Y-L, Wang Y-H, Qian C, Lu J, Li E, Wang W, Lu J, Xie Y, Wang J, Zhu D, Huang Z-X, Tang W (2001) Solution structure of cytochrome b 5 mutant (E44/48/56A/D60A) and its interaction with cytochrome c. Eur J Biochem 268:1620–1630 Świerczek M, Cieluch E, Sarewicz M, Borek A, Moser CC, Dutton PL, Osyczka A (2010) An electronic bus bar lies in the core of cytochrome bc 1. Science 329:451–454 Tian H, Sadoski R, Zhang L, Yu C-A, Yu L, Durham B, Millet F (2000) Definition of the interaction domain for cytochrome c on the cytochrome bc(1) complex. Steady-state and rapid kinetic analysis of electron transfer between cytochrome c and Rhodobacter sphaeroides cytochrome bc(1) surface mutants. J Biol Chem 275:9587–9595 Trinei M, Giorgio M, Cicalese A, Barozzi S, Ventura A, Migliaccio E, Milia E, Martin Padura I, Raker VA, Maccarana M, Petronilli V, Minucci S, Bernardi P, Lanfrancone L, Pelicci PG (2002) A p53-p66Shc signalling pathway controls intracellular redox status, levels of oxidation-damaged DNA and oxidative stress-induced apoptosis. Oncogene 21:3872–3878 Tsukihara T, Aoyama H, Yamashita E, Tomizaki T, Yamaguchi H, Shinzawa-Itoh K, Nakashima R, Yaono R, Yoshikawa S (1996) The whole structure of the 13-subunit oxidized cytochrome c oxidase at 2.8 Ǻ. Science 272:1136–1144 32 Figure 3