Critical Review Cardiolipin–Cytochrome c Complex: Switching Cytochrome c from an Electron-Transfer Shuttle to a Myoglobin- and a Peroxidase-like Heme-protein

Paolo Ascenzi1* Massimo Coletta2,3 Michael T. Wilson4 Laura Fiorucci2 Maria Marino5 Fabio Polticelli5,6 Federica Sinibaldi2 Roberto Santucci2

1

Interdepartmental Laboratory for Electron Microscopy, Roma Tre University, Roma, Italy 2 Department of Clinical Sciences and Translational Medicine, University of Roma “Tor Vergata,” Roma, Italy 3 Interuniversity Consortium for the Research on the Chemistry of Metals in Biological Systems, Bari, Italy 4 School of Biological Sciences, University of Essex, Colchester, UK 5 Department of Science, Roma Tre University, Roma, Italy 6 National Institute of Nuclear Physics, Roma Tre Section, Roma, Italy

Abstract Cytochrome c (cytc) is a small heme-protein located in the space between the inner and the outer membrane of the mitochondrion that transfers electrons from cytc-reductase to cytcoxidase. The hexa-coordinated heme-Fe atom of cytc displays a very low reactivity toward ligands and does not exhibit significant catalytic properties. However, upon cardiolipin (CL) binding, cytc achieves ligand binding and catalytic properties reminiscent of those of myoglobin and peroxidase. In particular, the peroxidase activity of the cardiolipin–cytochrome c complex (CL–cytc) is critical for the redistribution of CL from

the inner to the outer mitochondrial membranes and is essential for the execution and completion of the apoptotic program. On the other hand, the capability of CL–cytc to bind NO and CO and the heme-Fe-based scavenging of reactive nitrogen and oxygen species may affect apoptosis. Here, the ligand binding and catalytic properties of CL–cytc are analyzed in parallel with those of CL-free cytc, myoglobin, and peroxidase to dissect the potential mechanisms of CL in modulating the proC and anti-apoptotic actions of cytc. V 2015 IUBMB Life, 67(2):98–109, 2015

Keywords: cardiolipin–cytochrome c complex; redox properties; nitrite reductase activity; peroxidase activity; peroxynitrite detoxification; apoptosis

Introduction Abbreviations: CL, cardiolipin; CL–cytc, cardiolipin–cytochrome c complex; cytc, cytochrome c; Fe (III), ferric heme-Fe atom; Fe (II), ferrous heme-Fe atom; Mb, myoglobin C 2015 International Union of Biochemistry and Molecular Biology V Volume 67, Number 2, February 2015, Pages 98–109 *Address correspondence to: Paolo Ascenzi, Interdepartmental Laboratory for Electron Microscopy, Roma Tre University, Via della Vasca Navale 79, Roma, I-00146 Italy. Tel.: 139-06-5733-3621. Fax: 139-06-5733–6321. E-mail: [email protected] Received 6 October 2014; Accepted 11 January 2015 DOI 10.1002/iub.1350 Published online 9 April 2015 in Wiley Online Library (wileyonlinelibrary.com)

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Cytochrome c (cytc) is a small heme-protein located in the space between the inner and the outer membrane of the mitochondrion that transfers electrons from cytc-reductase (i.e., the bc1 complex) to cytc-oxidase (i.e., the terminal electron acceptor of the respiratory chain) (1–4). Approximately, 15% of mitochondrial cytc is tightly bound to the cristae of the mitochondrial inner membrane, cardiolipin (CL) representing the preferential interacting partner of cytc. This phospholipid (which represents about 20% of total membrane lipids) possesses a unique structure containing four (instead of two) fatty acid tails (5). The CL-bound cytc (CL–cytc) shows a non-native open tertiary structure and a weak heme-Fe distal ligation,

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cytosol where actively participates in the execution of the programmed cell death by binding to the apoptosis protease activation factor 1 (i.e., APAF-1) with the subsequent activation of procaspase-9 (12,29–37) (Fig. 1). Although nitrosylated and carbonylated cytc have not been yet detected in mitochondria, NO (cellular concentration ranging between subnanomolar and micromolar), CO (cellular concentration ranging between picomolar and nanomolar), peroxynitrite (cellular concentration ranging between subnanomolar and micromolar), and H2O2 (cellular concentration being submicromolar) interact with mitochondrial components, modulating a range of biological responses spanning from the modulation of mitochondrial respiration, to mitochondrial dysfunction and signaling of apoptotic cell death (13–15,17,20,35,38–43). It is worth noting that more than 1,000 mitochondrial proteins, involved and encompassed in biogenesis, proliferation, apoptosis, and autophagy, have been shown to be modulated by NO (44). Here, the reactivity and structural properties of CL–cytc are reviewed to dissect the potential mechanisms of CL in modulating the pro- and antiapoptotic actions of cytc.

The CL–Cytc Complex

FIG 1

Role of H2O2, NO, and CO on CL peroxidation and cytc release from mitochondria. APAF-1, apoptosis protease activation factor 1; Cas-9, caspase-9; CL, cardiolipin, CLox, peroxidized CL; cytc, cytochrome c; MPTP, mitochondrial permeability transition pore.

following the cleavage of the heme-Fe-Met80 distal bond. In particular, several independent studies provided convincing evidence that the heme-Fe(III) distal ligand of the CL–cytc complex is a Lys residue, although the available experimental data do not allow its unequivocal identification, the most likely candidate being Lys79 (6–9). As a consequence of the cleavage of the heme-Fe-Met80 distal bond, CL–cytc displays myoglobin (Mb)-like binding and catalytic properties (10–24). CL-bound cytc displays peroxidase activity, which is critical during the early stages of the apoptotic process generating CL hydroperoxides. CL peroxidation is in fact one of the factors considered responsible for the dissociation of the CL–cytc complex and for the redistribution of CL from the inner to the outer mitochondrial membrane, although the mechanisms governing these events remain unclear. This assumption is based on the observation that CL undergoes oxidative degradation in the p53-induced apoptosis (25), and that CL hydroperoxides show a decreased affinity for cytc with respect to CL itself (26). CL-free cytc, which is believed to reacquire its native-like sixcoordinated ferric conformation (23,27,28), is released in the

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Lipids have been reported to favor the formation of non-native protein conformers; indeed, lipid-bound cytc shows spectroscopic properties typical of the molten globule (35,45). CL binding to cytc has been proposed to occur at two distinct sites, namely A and C, which display different affinities for CL. At the A-site, the interaction, of electrostatic nature, involves positively charged residues of the protein and the negatively charged phosphate groups of CL, whereas at the C-site the interaction is of hydrophobic nature (23,46–48). Three main models have been hypothesized for CL binding to horse heart cytc. The first model proposes that the acyl chain of CL may protrude into the cytc matrix through the hydrophobic channel located close to the Asn52 residue. According to this model, the protonated phosphate group of CL could be H-bonded to the Asn52 residue of cytc (27,49). The second model suggests that CL binds close to the Met80containing loop of cytc, protruding into the protein matrix between the hydrophobic strands formed by residues 67–71 and 82–85. In particular, the deprotonated phosphate group of CL could be anchored to the Lys72 residue of cytc via electrostatic interactions (48). Based on the biphasic character of CL binding to cytc and on bioinformatic considerations, a third model hypothesizes that two adjacent acyl chains of CL could be accommodated in the two hydrophobic channels in the neighborhood of the Asn52 residue and in the region of the Met80-containing loop (Fig. 2) (8,23). In addition, site-directed mutagenesis studies indicate that Lys72 and Lys79 residues are critical for CL–cytc recognition, and the concurrent presence of Lys72, Lys73, and Lys79 residues is required for the peroxidase activity of the CL–cytc complex (8,9,23). In line with this observation, salts induce CL–cytc dissociation,

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the CL–cytc(III) binding reaction being 1.8 s21. This rate constant is independent of the CL concentration, suggesting the existence of a rate-limiting step, represented probably by the Fe-Met80 bond cleavage (23).

Redox, Ligand Binding, and Catalytic Properties of the CL–Cytc Complex Redox Properties of the CL–Cytc Complex

FIG 2

Schematic representation of the hypothetical structural model of the CL–cytc complex. CL would interact with cytc by the insertion of two acyl chains in the vicinity of the Asn52 and Met80 residues, respectively (8). The heme is colored by atom type, the iron atom in red, and the CL molecule in yellow. For the sake of clarity, only the heme iron proximal ligand (His18) and residues discussed in the text are shown in stick representation. The image was produced using UCSF Chimera molecular graphics package (50).

suggesting that the two negative CL head-group charges interact with approximately five positive surface charges of the protein (13). In any case, CL binding to cytc induces a large conformational change(s) of the heme pocket leading to the cleavage or the severe weakening of the Met80-heme-Fe bond which could be replaced by a weaker ligand, likely Lys79 (9), and the transformation of cytc into a globin- and peroxidaselike heme-protein (13,14,21,35,51). Finally, time-resolved fluorescence resonance energy transfer measurements (using dansyl-derivatives of cytc) point to different populations of conformers for the CL–cytc complex, some of which would display a high degree of protein unfolding (52). From the quantitative viewpoint, the values of the apparent dissociation equilibrium constant for CL binding to cytc(II) range between 2.2 3 1026 M at pH 6.5 and 1.4 3 1026 M at pH 8.1; approximately, 30 CL per cytc are required for pentacoordination of the heme-Fe(II) atom and full reactivity toward ligands (19). On the other hand, CL binding to cytc(III) follows a two-step process, values of the apparent dissociation equilibrium constants being 2.0 3 1025 M (first step) and 4.2 3 1025 M (second step), between pH 6.5 and 7.5. The different affinity of CL for ferrous and ferric cytc envisages a redox-linked conformational change of cytc. The CL–cytc(III) complex formation requires approximately six CL per cytc, the rate constant for

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The structural rearrangements occurring in cytc upon CL binding affect the protein redox properties; indeed, the redox potential of CL-bound cytc is significantly lower than that of the wild-type protein, being out of the range required for the cytc physiological role (21,53,54). In addition, CL–cytc contributes to activate the apoptotic cascade. This suggests that CL regulates the cytc functions by switching the cytc role from that of an electron-transfer shuttle to that of an important proapoptotic factor (28,55–57). The CL–cytc complex is an ensemble of subfractions with redox potentials ranging between 250 mV (vs. NHE) and 2200 mV (vs. NHE) (21,53). These values are close to those of Fe(III)-protoporphyrin IX (2107 mV vs. NHE) (58), indicating that CL binding to cytc indeed induces the cleavage of the Fe(III)-Met80 axial bond (9,48). In contrast, an electrochemical investigation on cytc-bound to a CL-modified electrode detected two protein subpopulations (i.e., 1 and 2) showing very different redox potentials (1224 and 1100 mV, respectively, vs. NHE). The redox potential of subpopulation 1 was found to be similar to that of cytc, whereas subpopulation 2 was assumed to correspond to CL–cytc, formed by CL interaction to the cytc hydrophobic site (54). Although the different redox potentials of the CL–cytc complex may reflect the very different experimental conditions, they indicate that CL binding to cytc stabilizes the oxidized form of the heme-protein and favors the formation of CL–cytc subpopulations in solution, most of which lack the Fe(III)-Met80 axial bond present in the CL-free cytc.

Nitrosylation and Carbonylation of Ferrous CL–Cytc Complex Nitrosylation of horse heart CL–cytc(II) is a multistep process involving a fast bimolecular reaction followed by slow monomolecular processes (Scheme 1) (14,20). The hexa- and pentacoordinated species of CL–cytc(II) (A and B, respectively, in Scheme 1) are in rapid equilibrium, and therefore the formation of the reactive penta-coordinated CL–cytc(II) species (B in Scheme 1) does not limit the formation of CL–cytc(II)–NO (C in Scheme 1). The very rapid NO-dependent bimolecular kinetic process (k11 5 2 3 107 M21 s21) is followed by the cleavage of the proximal His18-Fe(II) bond, leading to the formation of the penta-coordinated Fe(II)–NO derivative (D in Scheme 1). This penta-coordinated intermediate evolves toward a hexacoordinated adduct characterized by an unknown trans ligand (i.e., X; E in Scheme 1; k12 5 7 s21). Then, this hexa-coordinated

Cardiolipin Modulates Cytochrome C Actions

SCH 1

NO binding to ferrous CL–cytc. The formation of the reactive penta-coordinated CL–cytc(II) species (B) does not limit the formation of the initial CL–cytc(II)–NO complex (C). The formation of the final CL–cytc(II)–NO complex (G) is limited by the dissociation of the endogenous ligand X from the hexa-ccordinated X–CL–cytc(II)–NO species (E). The values of k11, k21, k12, and k13 are 2 3 107 M21 s21, 5 3 1024 s21, 7 s21, and 0.1 s21, respectively, at pH 7.4 and 22.0  C.

intermediate likely converts to a putative hexa-coordinated bisnitrosylated species (F in Scheme 1; k13 5 0.1 s21). The bis-nitrosylated intermediate (F in Scheme 1) would then evolve rapidly toward the final penta-coordinated species binding NO to the proximal side of the CL–cytc(II) heme pocket (G in Scheme 1). The values of the overall first-order rate constant for NO dissociation from horse heart CL–cytc(II)–NO (i.e., k21) and of the dissociation equilibrium constant for the adduct formation (i.e., K1) are 5 3 1024 s21 and 3310211 M, respectively (14,18,20) (Table 1). In the absence of structural data of the CL–cytc(II) complex, the binding mode of NO to the proximal heme coordination site is difficult to envisage. However, extensive investigations of NO binding to Alcaligenes xylosoxidans cytc’(II) provide some clues. One important factor appears to be the destabilization of NO bound to the distal position by locating a hydrophobic and relatively bulky amino acid residue (i.e., Leu16 in A. xylosoxidans cytc’(II)) above the distal binding site (see ref. 20 and references therein). This residue, while permitting NO binding, imposes some steric restraint on the heme distal site leading to a low NO affinity. The dissociation of the His120 residue from the proximal site of A. xylosoxidans cytc’(II)–NO (trans-effect of NO) allows a further NO molecule to bind and this, in turn, leads to the dissociation of the NO molecule bound to the low-affinity heme distal site. An analogous situation may be envisaged for CL–cytc(II) nitrosylation. In fact, there is circumstantial evidence (8) that one of the hydrocarbon chains of CL lies close to the heme-Fe atom on

the distal side, the Met80 residue being displaced from coordination (Fig. 2). Occluding the heme distal side (cf. the Leu16 residue in A. xylosoxidans cytc’(II)–NO), the Met80 residue could destabilize the heme distal-bound NO and perturb the balance of the affinities between the two heme sides in favor of proximal NO binding (20). In contrast, horse heart CL–cytc(II) binds reversibly CO to the heme distal site by simple bimolecular kinetics and thermodynamics (Scheme 2), the values of k11, k21, and K1 are 1 3 107 M21 s21, 2 3 1021 s21, and 3 3 1028 M, respectively (13) (Table 1). Remarkably, CO binding induces only a weak labilization of the trans-proximal His18–Fe(II) bond at variance with NO (70), this does not permit CO binding to the proximal heme side but it is sufficient to facilitate CO dissociation (13). For comparison, the binding mode of NO and CO to the proximal and distal heme pocket of A. xylosoxidans cytc’(II), respectively, (71) is shown in Fig. 3.

Nitrite Reductase Activity of Ferrous CL–Cytc Complex CL facilitates the conversion of NO22 to NO by horse heart cytc(II) (Scheme 3) in a dose-dependent manner (k11 5 2.6 M21 s21), concomitantly the formation of CL–cytc(III) (C in Scheme 3) takes place (19) (Table 1). As observed for most hemeproteins (see ref. 19 and references therein), the NO22-mediated conversion of (CL–)cytc–Fe(II) to (CL–)cytc–Fe(III) requires one proton (Scheme 3). Accordingly, on increasing the proton concentration by one pH unit, the rate of the NO22-mediated conversion of (CL–)cytc–Fe(II) to (CL–)cytc–Fe(III) increases by one order of magnitude (19).

Reductive Nitrosylation of Ferric CL–Cytc Complex

SCH 2

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CO binding to ferrous CL–cytc. The formation of the reactive penta-coordinated CL–cytc(II) species (B) does not limit the formation of the CL–cytc(II)–CO complex (C). The values of k11 and k21 are 1 3 107 M21 s21 and 1.8 3 1021 s21, respectively, at pH 7.4 and 25.0  C.

At neutral pH, NO binds reversibly to horse heart CL–cytc(III) (Scheme 4), kinetics of complex formation being independent of the NO concentration. This indicates that CL–cytc(III) nitrosylation is limited by a monomolecular process, likely referable to the dissociation rate from the sixth coordination position of the heme-Fe(III) atom of the residue substituting Met80 as the axial ligand in CL–cytc, likely Lys79 (i.e., k21  6 s21; Scheme 4); the value of k11 is 80 s21 (9,17,18). The transient pentacoordinated CL–cytc(III) species (B in Scheme 4) reacts very rapidly with NO (leading to C in Scheme 4), as observed after photolysis (i.e., k12  1 3 106 M21 s21); the value of k22 is

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TABLE 1

Values of kinetic and thermodynamic parameters for ligand binding and (pseudo-)enzymatic properties of horse heart cytc, horse heart CL–cytc, mammalian Mbs, and horseradish peroxidase

Parameter

Horse heart cytc

Horse heart CL–cytc

Mammalian Mb

Horseradish peroxidase

8.3a

2 3 107b

1.6 3 107c

k21 (s21)

2.9 3 1025a

5 3 1024b

1.3 3 1024d

K1 (M)

6.1 3 1026a

3 3 10211e

8.1 3 10212f

6.5 3 1025g

1 3 107h

5.0 3 105i

3.4 3 103j

k21 (s21)

1.8 3 1021h

1.7 3 1022i

1.1 3 1024j

K1 (M)

1.8 3 1028h

3.4 3 1028i

2.9 3 1028j

2.6k

2.9l

Fe(II) nitrosylation k11 (M21 s21)

k12 (s21)

7b

k13 (s21)

1 3 1021b

Fe(II) carbonylation k11 (M21 s21)

Fe(II) nitrite reduction k11 (M21 s21)

7.3 3 1022k

Fe(III) reductive nitrosylation k11 (s21)

80m

k21 (s21)

6m 7.2 3 102n

1 3 106m

6.8 3 104o

k22 (s21)

4.4 3 1022n

5m

5.2o

K2 (M)

6.1 3 1025n

5 3 1026m

1.2 3 1024o

1.8 3 103p

3 3 103q

3.9 3 102r

3.2 3 105s

2.9 3 104t

6.0 3 102u

3.2 3 103u

1.9 3 102v

1.7 3 105u

kcat (s21)

1.4 3 1022u

2.0 3 1022u

1.1 3 1021v

5.6 3 102u

Km (M)

2.3 3 1025u

6.2 3 1026u

5.7 3 1024v

3.3 3 1023u

k12 (M21 s21)

k13 (M21 s21) Fe(III) peroxynitrite isomerization k11 (M21 s21) Fe(III) guaiacol peroxidation kcat/Km (M21 s21)

a

pH 6.5 and room temperature (59). pH 7.4 and 22.0  C (14). c Horse heart Mb; pH 9.2 and 20.0  C (60). d Horse heart Mb; pH 7.0 and 20.0  C (61). e Calculated from ref. 14 (pH 7.4 and 22.0  C). f Horse heart Mb; calculated from ref. 60 (pH 9.2 and 20.0  C) and ref. 61 (pH 7.0 and 20.0  C). g pH 7.4 and 25.0  C, under anaerobic conditions (62). h pH 7.4 and 25.0  C (13). i Horse heart Mb; pH 7.0 and 20.0  C (63). j pH 7.0 and 20.0  C (64). k pH 7.4 and 20.0  C (19). l Horse heart Mb; pH 7.4 and 25.0  C (65). m pH 7.4 and 22.0  C (17). n pH 6.5 and 20.0  C (59). o Horse heart Mb; pH 9.2 and 20.0  C (60). p pH independent value, 20.0  C (59). q pH independent value and 20.0  C (18). r Horse heart Mb; pH independent value, 20.0  C (60). s pH 7.0 and 20.0  C (15). t Horse heart Mb; pH 7.0 and 20.0  C (66). u pH 7.0 and 25.0  C (67,68). v Sperm whale Mb; pH 7.0 and 20.0  C (69). b

FIG 3

The structures of ferrous nitrosylated and carbonylated A. xylosoxidans cytc’(II). In the nitrosylated derivative of A. xylosoxidans cytc’(II) (Ax-cytc’–NO), the displacement of the proximal His120 residue and the flipping of the Arg124 side chain (which stakes against the heme plane and is hydrogen bonded to conformer 2 of NO) occur. In the carbonylated derivative of A. xylosoxidans cytc’(II) (Ax-cytc’–CO), the displacement of the Leu16 side chain takes place, the proximal heme-Fe(II) atom axial ligand is His120. In addition, a second CO molecule is hydrogen bonded to the Nd1 atom of His120 (dashed line). Atomic coordinates have been taken from PDB entries 1E85 and 1E86 (71). The image was produced using UCSF Chimera molecular graphics package (50).

5 s21. In addition, NO saturation of CL–cytc(III) at the lowest ligand concentration explored (5 3 1025 M) suggests that the value of the apparent dissociation equilibrium constant (i.e., K) is 5 3 1026 M (9,17,18) (Table 1). Between pH values of 7.9 and 9.5, the irreversible reductive nitrosylation of horse heart CL–cytc(III) occurs (C–E species; Scheme 4) after initial NO binding. In fact, CL–cytc(III)– NO (C in Scheme 4) is in rapid equilibrium with the CL– cytc(II)–NO1 species (D in Scheme 4), which converts to CL– cytc(II) (E in Scheme 4); this process depends linearly on the OH2 concentration (k13 5 3.0 3 103 M21 s21) (Table 1). Fur-

SCH 3

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Nitrite reductase activity of ferrous CL–cytc.The k11 value is 2.6 M21 s21, at pH 7.4 and 20.0  C. The dashed arrow indicates the occurrence of reaction intermediate(s).

thermore, CL–cytc(II) converts to CL–cytc(II)–NO very rapidly in the presence of NO excess (18).

Peroxynitrite Detoxification by Ferric CL–Cytc Complex Horse heart CL–cytc(III) catalyzes the isomerization of peroxynitrite in a dose-dependent fashion (Scheme 5) (15). The value of the second-order rate constant for the CL–cytc(III)mediated isomerization of peroxynitrite reflects the formation of the CL–cytc(III)–OONO complex (C in Scheme 5; k12 5 3.2 3 105 M21 s21), which converts very rapidly and irreversibly to CL–cytc(III) and NO32 (D in Scheme 5) (15). Apparently, the formation of the CL–cytc(III)–OONO complex is not limited by the cleavage of the Lys79–Fe(III) heme distal bond. In fact, the observed values of the pseudo-first-order rate constant for HOONO binding to CL–cytc(III) (ranging between 2.6 and 12 s21) increase linearly with the CL– cytc(III) concentration (15) and exceed the values of the firstorder rate constant for CL–cytc(III) penta-coordination (k21 ranging between 6 and 9.3 s21; see Scheme 4) (17,18) (Table 1). This suggests that an unknown intermediate is involved in the formation of the CL–cytc(III)–OONO complex (C in Scheme 5). It is worth noting that, although both NO and peroxynitrite are likely to be present in the intermembrane space (40), NO could not inhibit peroxynitrite

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SCH 4

Reductive nitrosylation of ferric CL–cytc. The formation of the CL–cytc(III)–NO complex is limited by the dissociation of the sixth coordination ligand of the heme-Fe(III) atom. At pH  7.9 and 20.0  C, the formation of the CL–cytc(III)–NO complex is followed by the reduction of the heme-Fe(III) atom and the formation of the CL–cytc(II)–NO adduct, in the presence of NO excess. The reductive nitrosylation of the CL–cytc(III) complex is limited by the [OH2]-dependent reaction. The value of the [OH2]-dependent apparent first-order rate constant increases from 4.1 3 1023 s21 (at pH 7.9) to 6.7 3 1022 s21 (at pH 9.5); k13 5 3.0 3 103 M21 s21, at 20.0  C. The values of k11, k21, k12, and k22 are 80 s21, 6 s21, 1 3 106 M21 s21, and 5 s21, respectively, at pH 7.4 and 22.0  C. The dashed arrow indicates the occurrence of reaction intermediate(s).

detoxification by CL–cytc–Fe(III) as it displays a low reactivity toward CL–cytc–Fe(III) (Table 1).

Peroxidase Activity of CL–Cytc Complex In the absence of CL, cytc has a very weak peroxidase activity (21); however, upon binding of CL, the penta-coordination or weak hexa-coordination of the heme-Fe atom confers peroxidase competence on cytc that is specific toward CL peroxidation (51). The generation of CL hydroperoxides is involved in the release of pro-apoptotic factors, including cytc, from the mitochondrial membrane. In addition, as H2O2 may enter in the CL–cytc peroxidase cycle, behaving as an effective source of oxidizing equivalents, CL–cytc eliminates H2O2, contributing to antioxidant cell defenses (11,12,21). The contribution of the CL–cytc complex to the maintenance of the antioxidant balance in mitochondria is not only related to the removal of H2O2. In fact, free fatty acid-hydroperoxides are better substrates for CL–cytc than H2O2, suggesting that the control of fatty acidhydroperoxides may be an additional antioxidant function of cytc in damaged mitochondria (11). In addition, as the production and accumulation of oxidized CL may have a physiological function as apoptotic signal in the process of elimination of injured mitochondria, cytc has been proposed to control the levels of oxygenated fatty acids generated from the hydrolysis of CL by phospholipase A2 (72). Although the catalytic mechanism is not fully understood, the peroxidase activity of horse heart CL–cytc(III) (73–75) has been analyzed in the framework of the classical peroxidase cycle involving compound I (CL–cytc(IV) 5 OAR1•; B in Scheme

SCH 5

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Peroxynitrite detoxification by ferric CL–cytc. The values of k21, k11, and k12 are 6 s21, 80 s21, and 3.2 3 105 M21 s21, respectively, at pH 7.0 and 20.0  C.

6) and II (Fe(IV); C in Scheme 6) (76). In compound I, the second oxidizing equivalent is stored as an organic radical (i.e., R1•), mainly residing on Tyr67 (73–75), which is the likely electron donor (radical acceptor) in the oxygenase halfreaction catalyzed by CL–cytc (73). It is worth noting that Tyr67 is positioned close to the hypothesized binding site of CL, which undergoes peroxidation (73). However, multiple organic radical species have been reported for CL–cytc, and this is consistent with the heterogeneous nature of the CL–cytc ensemble (77,78). Interestingly, the single mutation of trimethyl-Lys72 to Ala in yeast iso-1-cytc induces small but significant conformational changes of the heme crevice that

SCH 6

Classical catalytic peroxidase cycle applied to ferric CL–cytc. Dashed arrows indicate the likely occurrence of (not yet detected) reaction intermediate(s).

Cardiolipin Modulates Cytochrome C Actions

lead to the detachment of Met80 from the heme-Fe atom and the enhancement of the peroxidase activity (79). CL peroxidation by cytc appears to be modulated not only by H2O2 but also by fatty acid-hydroperoxides, including CL hydroperoxides, as a source of oxidizing equivalents feeding the peroxidase cycle. In fact, fatty acid-hydroperoxides can accelerate the CL peroxidation by CL–cytc more than two orders of magnitude (11). Therefore, H2O2 loses its regulatory role in CL peroxidation by CL–cytc with the accumulation of CL hydroperoxides. The peroxidase activity and conformation of CL–cytc are also modulated by ATP which weakens the CL– cytc interactions and allows the heme-protein refolding into its native structure (45,67,68,77,78,80). In addition, the occurrence of Tyr phosphorylation sites in cytc suggested that this type of signaling may regulate the peroxidase activity. In fact, Tyr48 phosphorylation might serve as an anti-apoptotic switch, possibly via its effects on CL peroxidation during apoptosis (81,82). Furthermore, CO effectively competes with H2O2 for the heme-Fe atom inhibiting the peroxidase activity. In fact, upon mixing CL–cytc(II) with H2O2, heme degradation occurs within 1 sec, whereas in the presence of CO, this reaction occurs with a rate of 1024 to 1023 s21. A similar rate was observed for the reaction of H2O2 with hexa-coordinated cytc in the absence of CL (13). Although NO has been postulated to affect the peroxidase activity of CL–cytc (14), cytc–Fe(II)–NO formation seems to represent a less important event in the mechanism of inhibition of the peroxidase activity of CL–cytc when compared with the reduction by NO of compounds I and II as well as of the organic radical species (83).

Cytc Versus the CL–Cytc Complex CL binding to horse heart cytc switches this heme-protein from an electron transfer shuttle to a Mb- and a peroxidaselike heme-protein. The reaction parameters listed in Table 1 have been obtained between pHs of 6.5 and 9.2 and between 20.0 and 25.0  C. The pH range investigated covers the physiological pH interval rendering the reaction parameters biologically relevant. Although the reaction parameters were obtained in a temperature range lower than that of the physiological temperature, this permits the homogeneous comparison with most of the data obtained for monomeric hemeproteins. The inspection of Table 1, showing kinetic and thermodynamic parameters for ligand binding and catalytic properties of cytc, CL–cytc, mammalian Mb, and horseradish peroxidase, allows the following considerations. i. CL facilitates allosterically NO and CO binding to horse heart cytc(II) (13,14,20), which undergoes nitrosylation very slowly (59) and carbonylation not at all (84) in the absence of CL. In addition, CL induces NO binding to the proximal site of the cytc(II) heme pocket (13,14,20,85,86) (Scheme 1). In contrast, CO binds to the heme distal site CL–cytc(III) with a simple mechanism (13) (Scheme 2). It is worth noting that both NO and CO bind to the heme distal site of sperm whale Mb(II) (87,88). Therefore, NO and CO

Ascenzi et al.

ii.

iii.

iv.

v.

vi.

vii.

viii.

binding to the heme proximal and distal sites of CL–cytc, respectively, may be at the root of selective diatomic gasbased signal transduction pathways. CL facilitates the nitrite reductase activity of horse heart cytc(II) (Scheme 3) (19), which becomes very similar to that observed for horse heart Mb(II) (65); in contrast, cytc(II) catalyzes very slowly the conversion of NO22 to NO (19). As reported for CL–cytc(II) nitrosylation and carbonylation (13,14,20), the nitrite reductase activity of CL– cytc(II) is not limited by the hexa-to-penta-coordination transition of the heme-Fe(II) atom (19). At neutral pH, nitrosylation of CL-bound horse heart cytc(III) follows a simple mechanism limited by the dissociation of the endogenous sixth coordination ligand of the heme-Fe(III) atom (17,18) (Scheme 4). However, at alkaline pH, the CL–cytc(III)–NO complex evolves to the CL– cytc(II)–NO species (18), as reported for sperm whale Mb(III) (89). In contrast, NO converts cytc(III) to cytc(II), which, however, does not undergo nitrosylation (59). In the absence of CL, hexa-coordinated horse heart cytc(III) does not catalyze peroxynitrite isomerization (15), although peroxynitrite may induce the very slow nitration of the solvent-exposed Tyr74 residue. This leads to the cleavage of the distal heme-Fe(III)-Met80 bond, which could be substituted by the weak heme-Fe(III)– Lys72 ligation (6). Although CL–cytc(III) (15) and horse heart Mb(III) (66) display similar catalytic parameters for peroxynitrite isomerization, an unknown transient may be involved in the formation of the CL–cytc(III)–OONO adduct (Scheme 5). CL-bound horse heart cytc displays a more favorable kcat/ Km value than that of cytc for guaiacol peroxidation mainly reflecting a lower Km value (67,68). It is worth noting that although the kcat/Km and kcat values for guaiacol peroxidation by horseradish peroxidase are higher than those of CL–cytc, the substrate affinity is lower (68). The CL-induced hexa-to-penta-coordination transition of the heme-Fe atom of cytc depends on its oxidation state, limiting mainly ligand binding and catalytic properties of the CL–cytc(III) derivative (13–15,17–20). Reactions shown in Schemes 1–6 depend on the redox state of the heme-Fe atom of (CL–)cytc. Because of the very high affinity of cytc oxydase, cytc is preferentially in the ferric form under normal conditions (90); therefore, it could be postulated that (CL–)cytc–Fe(III) undergoes preferentially reductive nitrosylation, peroxynitrite isomerization, and substrate peroxidation. On the other hand, under ischemic conditions where the heme-Fe atom is in the ferrous state (90), (CL–)cytc–Fe(II) nitrosylation, carbonylation, and nitrite reduction could take place preferentially. The estimated cellular concentrations of NO, CO, H2O2, and peroxynitrite (39–43) are compatible with kinetic, thermodynamic, and catalytic parameters for the reactions of monomeric heme-proteins.

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Conclusions and Perspectives The capability of triggering either pro- or anti-apoptotic processes renders the CL–cytc complex a crucial element for the regulation of the cell life/death balance (35,57,91). CL–cytc functions as a pro-apoptotic factor catalyzing the peroxidative reduction of H2O2, which leads, among others, to CL peroxidation and reactive oxygen species generation, including even singlet oxygen (91,92). In turn, reactive oxygen species accelerate the formation of a pore in the outer membrane and the release of cytc in the cytosol (91). In this respect, it has been recently reported that the excessive reactive oxygen species production during oxidative stress brings about the oxidation of Met80, inducing the conversion of cytc into a peroxidase able to oxidize the lynoleic acid groups of CL. The cytc peroxidase-like activity reduces then H2O2 to H2O, generating oxidized CL and favoring both CL redistribution into mitochondria and cytc release in the cytosol (93) (Fig. 1). When in the cytoplasm, cytc interacts with APAF-1 leading to a multimeric apoptosome that activates pro-caspase-9 (94); in turn, caspase-9 triggers execution caspase activation inducing apoptosis (12,30–36,91,95,96) (Fig. 1). However, CL–cytc could also exert an anti-apoptotic action facilitating the isomerization of peroxynitrite to nitrate and the detoxification of reactive nitrogen and oxygen species (15,97,98). Furthermore, the reaction of CO, NO, and NO22 with CL–cytc(II) and CL–cytc(III) (13,14,17,19) could impair the heme-protein reactivity including CL peroxidation and therefore the initiation of the cell death program by the release of pro-apoptotic factors (including cytc) in the cytoplasm (10,35,99,100). Thus, the apoptotic cascade could be efficiently stopped by specifically inhibiting CL peroxidation; this might explain the reason why apoptotic reactions can be prevented or diminished by antioxidants (101). Accounting for in vitro kinetic, thermodynamic, and catalytic parameters of CL–cytc listed in Table 1, therapeutic levels of NO could affect only cytc(II) reactivity. In fact, although a direct relationship between plasma nitrate and nitrite concentrations (generally taken as precursors and end-products of NO metabolism) and clinical effects of NO-donors cannot be firmly established, plasma NO levels in the micromolar range may be reached after drug administration (102–104). Endogenous and exogenous compounds impairing neuron apoptosis have been reported to act as inhibitors of CL peroxidation (68,105). Remarkably, owing to the fundamental function of CL–cytc in cell apoptosis, new mitochondria-targeted compounds, including electron acceptors, electron donors, and hydride acceptors, have been reported to regulate CL peroxidation and are potentially exploitable as anti- or pro-apoptotic drugs (106–108). Over the last decade, several chemicals, including dopamine, L-DOPA, minocycline, and nitroxide/gramicidin, have been reported to efficiently decrease the progression of degenerative diseases inhibiting apoptosis by impairing CL– cytc peroxidase activity (68, 105-112). Mitochondria-targeted peptides have also been reported to represent a promising therapeutic approach for treating mitochondrial dysfunction(s). In

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particular, SS-31 (D-Arg-dimethylTyr-Lys-Phe-NH2), which is capable of interacting with CL, turns out to be an antioxidant peptide by preventing CL interaction with cytc and the consequent conversion of cytc into a peroxidase (113). Furthermore, a new group of mitochondria-targeted imidazole-substituted analogs of stearic acid has been proven to impair CL-induced structural rearrangements in cytc, thus preventing the appearance of a peroxidase-like activity (114). In view of the role of CL oxidation in mitochondrial dysfunction, interventions designed to preserve mitochondrial CL integrity and function from reactive oxygen species might be an effective strategy for the therapy of severe disorders, including neurodegenerative diseases, diabetic complications, and ischemia–reperfusion injury.

Acknowledgements The authors thank Dr. Loris Leboffe for helpful discussions. This study was supported by grants from the Ministero dell’Is e della Ricerca of Italy (PRIN truzione, dell’Universita 20109MXHMR_001 to P. A. and FIRB RBNE03PX83 to M. C.),  Roma Tre (CLA 2014 to P. A.). and Universita

References [1] Pettigrew, G. W. and Moore, G. R. (1987) Cytochromes c. Biological aspects. Springer-Verlag, Heidelberg. [2] Meyer, T. E. (1996) Evolution and classification of c-type cytochromes. In: Cytochrome c. A Multidisciplinary Approach. (Scott, R. A. and Mauk, A. G., eds.). Chapter 2, pp. 33–99, University Science Books, Sausalito, CA. [3] Ow, Y.-L. P., Green, D. R., Hao, Z., and Mak, T. W. (2008) Cytochrome c: functions beyond respiration. Nat. Rev. Mol. Cell. Biol. 9, 532–542. [4] Pun, P. B., Lu, J., Kan, E. M., and Moochhala, S. (2010) Gases in the mitochondria. Mitochondrion 10, 83–93. [5] Lewis, R. N. and McElhaney, R. N. (2009) The physicochemical properties of cardiolipin bilayers and cardiolipin-containing lipid membranes. Biochim. Biophys. Acta 1788, 2069–2079. [6] Abriata, L. A., Cassina, A., Tortora, V., Marin, M., Souza, J. M., et al. (2009) Nitration of solvent-exposed tyrosine 74 on cytochrome c triggers heme ironmethionine 80 bond disruption: nuclear magnetic resonance and optical spectroscopy studies. J. Biol. Chem. 284, 17–26. [7] Bradley, J. M., Silkstone, G., Wilson, M. T., Cheesman, M. R., and Butt, J. N. (2011) Probing a complex of cytochrome c and cardiolipin by magnetic circular dichroism spectroscopy: implications for the initial events in apoptosis. J. Am. Chem. Soc. 133, 19676–19679. [8] Sinibaldi, F., Howes, B. D., Piro, M. C., Polticelli, F., Bombelli, C., et al. (2010) Extended cardiolipin anchorage to cytochrome c: a model for proteinmitochondrial membrane binding. J. Biol. Inorg. Chem. 15, 689–700. [9] Sinibaldi, F., Howes, B. D., Droghetti, E., Polticelli, F., Piro, M. C., et al. (2013) Role of lysines in cytochrome c-cardiolipin interaction. Biochemistry 52, 4578–4588. [10] Kagan, V. E., Tyurin, V. A., Jiang, J., Tyurina, Y. Y., Ritov, V. B., et al. (2005) Cytochrome c acts as a cardiolipin oxygenase required for release of proapoptotic factors. Nat. Chem. Biol. 1, 223–232. [11] Belikova, N. A., Tyurina, Y. Y., Borisenko, G., Tyurin, V., Samhan Arias, A. K., et al. (2009) Heterolytic reduction of fatty acid hydroperoxides by cytochrome c/cardiolipin complexes: antioxidant function in mitochondria. J. Am. Chem. Soc. 131, 11288–11289. [12] Kagan, V. E., Bayır, H. A., Belikova, N. A., Kapralov, O., Tyurina, Y. Y., et al. (2009) Cytochrome c/cardiolipin relations in mitochondria: a kiss of death. Free Radic. Biol. Med. 46, 1439–1453.

Cardiolipin Modulates Cytochrome C Actions

[13] Kapetanaki, S. M., Silkstone, G., Husu, I., Liebl, U., Wilson, M. T., et al. (2009) Interaction of carbon monoxide with the apoptosis-inducing cytochrome c-cardiolipin complex. Biochemistry 48, 1613–1619. [14] Silkstone, G., Kapetanaki, S. M., Husu, I., Vos, M. H., and Wilson, M. T. (2010) Nitric oxide binds to the proximal heme coordination site of the ferrocytochrome c/cardiolipin complex: formation mechanism and dynamics. J. Biol. Chem. 285, 19785–19792. [15] Ascenzi, P., Ciaccio, C., Sinibaldi, F., Santucci, R., and Coletta, M. (2011) Cardiolipin modulates allosterically peroxynitrite detoxification by horse heart cytochrome c. Biochem. Biophys. Res. Commun. 404, 190–194. [16] Ascenzi, P., Ciaccio, C., Sinibaldi, F., Santucci, R., and Coletta, M. (2011) Peroxynitrite detoxification by horse heart carboxymethylated cytochrome c is allosterically modulated by cardiolipin. Biochem. Biophys. Res. Commun. 415, 463–467. [17] Silkstone, G., Kapetanaki, S. M., Husu, I., Vos, M. H., and Wilson, M. T. (2012) Nitric oxide binding to the cardiolipin complex of ferric cytochrome c. Biochemistry 51, 6760–6766. [18] Ascenzi, P., Marino, M., Ciaccio, C., Santucci, R., and Coletta, M. (2014) Reductive nitrosylation of the cardiolipin-ferric cytochrome c complex. IUBMB Life 66, 438–447. [19] Ascenzi, P., Marino, M., Polticelli, F., Santucci, R., and Coletta, M. (2014) Cardiolipin modulates allosterically the nitrite reductase activity of horse heart cytochrome c. J. Biol. Inorg. Chem. 19, 1195–1201. [20] Hough, M A., Silkstone G., Worrall, J. A. R., and Wilson, M. T. (2014) NO binding to the proapoptotic cytochrome c-cardiolipin complex. Vitam. Horm. 96, 193–209. [21] Basova, L. V., Kurnikov, I. V., Wang, L., Ritov, V. B., Belikova, N. A., et al. (2007) Cardiolipin switch in mitochondria: shutting off the reduction of cytochrome c and turning on the peroxidase activity. Biochemistry 46, 3423– 3434. [22] Belikova, N. A., Jiang, J., Tyurina, Y. Y., Zhao, Q., Epperly, M. W., et al. (2007) Cardiolipin-specific peroxidase reactions of cytochrome c in mitochondria during irradiation-induced apoptosis. Int. J. Radiat. Oncol. Biol. Phys. 69, 176–186. [23] Sinibaldi, F., Fiorucci, L., Patriarca, A., Lauceri, R., Ferri, T., et al. (2008) Insights into cytochrome c-cardiolipin interaction: role played by ionic strength. Biochemistry 47, 6928–6935. [24] Abe, M., Niibayashi, R., Koubori, S., Moriyama, I., and Miyoshi, H. (2011) Molecular mechanisms for the induction of peroxidase activity of the cytochrome c-cardiolipin complex. Biochemistry 50, 8383–8391. [25] Polyak, K., Xia, Y., Zweier, J. L., Kinzler, K. W., and Vogelstein, B. (1997) A model for p53-induced apoptosis. Nature 389, 300–305. [26] Shidoji, Y., Hayashi, K., Komura, S., Ohishi, N., and Yagi, K. (1999) Loss of molecular interaction between cytochrome c and cardiolipin due to lipid peroxidation. Biochim. Biophys. Res. Commun. 264, 343–347. € maa, M. and Kinnunen, P. K. J. (1995) Reversibility of the binding of [27] Ryto cytochrome c liposomes: implications for lipid-protein interactions. J. Biol. Chem. 270, 3197–3202. [28] Brown, G. C. and Borutaite, V. (2008) Regulation of apoptosis by the redox state of cytochrome c. Biochim. Biophys. Acta 1777, 877–881. [29] Li, P., Nijhawan, D., Budihardjo, I., Srinivasula, S. M., Ahmad, M., et al. (1997) Cytochrome c and dATP-dependent formation of Apaf-1/caspase-9 complex initiates an apoptotic protease cascade. Cell 91, 479–489. [30] Iverson, S. L. and Orrenius, S. (2004) The cardiolipin-cytochrome c interaction and the mitochondrial regulation of apoptosis. Arch. Biochem. Biophys. 423, 37–46. [31] Twiddy, D., Brown, D. G., Adrain, C., Jukes, R., Martin, S. J., et al. (2004) Pro-apoptotic proteins released from the mitochondria regulate the protein composition and caspase-processing activity of the native Apaf-1/caspase-9 apoptosome complex. J. Biol. Chem. 279, 19665–19682. [32] Garrido, C., Galluzzi, L., Brunet, M., Puig, P. E., Didelot, C., et al. (2006) Mechanisms of cytochrome c release from mitochondria. Cell Death Differ. 13, 1423–1433. [33] Riedl, S. J. and Salvesen, G. S. (2007) The apoptosome: signalling platform of cell death. Nat. Rev. Mol. Cell. Biol. 8, 405–413.

Ascenzi et al.

[34] Caroppi, P., Sinibaldi, F., Fiorucci, L., and Santucci, R. (2009) Apoptosis and human diseases: mitochondrion damage and lethal role of cytochrome c as proapoptotic protein. Curr. Med. Chem. 16, 4058–4065. [35] Ascenzi, P., Polticelli, F., Marino, M., Santucci, R., and Coletta, M. (2011c) Cardiolipin drives cytochrome c proapoptotic and antiapoptotic actions. IUBMB Life 63, 160–165. [36] Kulikov, A. V., Shilov, E. S., Mufazalov, I. A., Gogvadze, V., Nedospasov, S. A., et al. (2012) Cytochrome c: the Achilles’ heel in apoptosis. Cell Mol. Life Sci. 69, 1787–1797. [37] Kagan, V. E., Chu, C. T., Tyurina, Y. Y., Cheikhi, A., and Bayir, H. (2014) Cardiolipin asymmetry, oxidation and signaling. Chem. Phys. Lipids 179, 64–69. [38] Orrenius, S., Gogvadze, V., and Zhivotovsky, B. (2007) Mitochondrial oxidative stress: implications for cell death. Annu. Rev. Pharmacol. Toxicol. 47, 143–183. [39] Boveris, A. and Cadenas, E. (2000) Mitochondrial production of hydrogen peroxide regulation by nitric oxide and the role of ubisemiquinone. IUBMB Life 50, 245–250. [40] Radi, R., Cassina, A., and Hodara, R. (2002) Nitric oxide and peroxynitrite interactions with mitochondria. Biol. Chem. 383, 401–409. [41] Colasanti, M. and Suzuki, H. (2000) The dual personality of NO. Trends Pharmacol. Sci. 21, 249–252. [42] Gullotta, F., di Masi, A., Coletta, M., and Ascenzi, P. (2012) CO metabolism, sensing, and signaling. Biofactors 38, 1–13. [43] Ascenzi, P., di Masi, A., Sciorati, C., and Clementi, E. (2010) Peroxynitrite— an ugly biofactor? Biofactors 36, 264–273. [44] Piantadosi, C. A. (2012) Regulation of mitochondrial processes by protein Snitrosylation. Biochim. Biophys. Acta 1820, 712–721. [45] Muenzner, J. and Pletneva, E. V. (2014) Structural transformations of cytochrome c upon interaction with cardiolipin. Chem. Phys. Lipids 179, 57–63. € maa, M. and Kinnunen, P. K. J. (1994) Evidence for two distinct acidic [46] Ryto phospholipids-binding sites in cytochrome c. J. Biol. Chem. 269, 1770–1774. [47] Tuominen, E. K., Zhu, K., Wallace, C. J., Clark-Lewis, I., Craig, D. B., et al. (2001) ATP induces a conformational change in lipid-bound cytochrome c. J. Biol. Chem. 276, 19356–19362. [48] Kalanxhi, E. and Wallace, C. J. A. (2007) Cytochrome c impaled: investigation of the extended lipid anchorage of a soluble protein to mitochondrial membrane models. Biochem. J. 407, 179–187. [49] Tuominen, E. K., Wallace, C. J., and Kinnunen, P. K. J. (2002) Phospholipidcytochrome c interaction: evidence for the extended lipid anchorage. J. Biol. Chem. 277, 8822–8826. [50] Pettersen, E.F., Goddard, T.D., Huang, C.C., Couch, G.S., Greenblatt, D.M., et al. (2004) UCSF Chimera—a visualization system for exploratory research and analysis. J. Comput. Chem. 25, 1605–1612. [51] Belikova, N. A., Vladimirov, Y. A., Osipov, A. N., Kapralov, A. A., Tyurin, V. A., et al. (2006) Peroxidase activity and structural transitions of cytochrome c bound to cardiolipin-containing membranes. Biochemistry 45, 4998–5009. [52] Hanske, J., Toffey, J. R., Morenz, A. M., Bonilla, A. J., Schiavoni, K. H., et al. (2012) Conformational properties of cardiolipin-bound cytochrome c. Proc. Natl. Acad. Sci. USA 109, 125–130. [53] Oellerich, S., Lecomte, S., Paternostre, M., Heimburg, T., and Hidebrandt, P. (2004) Peripheral and integral binding of cytcochrome c to phospholipids vesicles. J. Phys. Chem. B 108, 3871–3878. [54] Perhirin, A., Kraffe, E., Marty, Y., Quentel, F., Elies, P., et al. (2013) Electrochemistry of cytochrome c immobilized on cardiolipin-modified electrodes: a probe for protein-lipid interactions. Biochim. Biophys. Acta 1830, 2798– 2803. [55] Kagan, V. E., Borisenko, G., Tyurina, Y. Y., Tyurin, V. A., Jiang, J., et al. (2004) Oxidative lipidomics of apoptosis: redox catalytic interactions of cytochrome c with cardiolipin and phosphatidylserine. Free Radic. Biol. Med. 37, 1963–1985. [56] Sharonov, G. V., Feofanov, A. V., Bocharova, O. V., Astapova, M. V., Dedukhova, V. I., et al. (2005) Comparative analysis of proapoptotic activity of cytochrome c mutants in living cells. Apoptosis 10, 797–808. [57] Bayir, H., Fadeel, B., Palladino, M. J., Witasp, E., Kurnikov, I. V., et al. (2006) Apoptotic interactions of cytochrome c: redox flirting with anionic

107

IUBMB LIFE

[58]

[59]

[60]

[61]

[62]

[63] [64]

[65]

[66] [67]

[68]

[69]

[70] [71]

[72]

[73]

[74]

[75]

[76]

[77]

[78]

phospholipids within and outside of mitochondria. Biochim. Biophys. Acta 1757, 648–659. Faraoni, A., Santucci, R., Campanella, L., Tranchida, G., and Brunori, M. (1990) Voltammetric studies on the electrochemical behaviour of membrane-entrapped hemes. Biol. Metals 3, 122–124. Hoshino, M., Ozawa, K., Seki, H., and Ford, P. C. (1993) Photochemistry of nitric oxide adducts of water-soluble iron(III) porphyrin and ferrihemoproteins studied by nanosecond laser photolysis. J. Am. Chem. Soc. 115, 9568– 9575. Ascenzi, P., di Masi A., Gullotta, F., Mattu, M., Ciaccio, C., et al. (2010) Reductive nitrosylation of ferric cyanide horse heart myoglobin is limited by cyanide dissociation. Biochem. Biophys. Res. Commun. 393, 196–200. Møller, J. K. and Skibsted, L. H. (2004) Mechanism of nitrosylmyoglobin autoxidation: temperature and oxygen pressure effects on the two consecutive reactions. Chemistry 10, 2291–2300. Qiang, L., Zhu, S., Ma, H., and Zhou, J. (2010) Investigation on binding of nitric oxide to horseradish peroxidase by absorption spectrometry. Spectrochim. Acta A Mol. Biomol. Spectrosc. 75, 417–421. Antonini, E. and Brunori, M. (1971) Hemoglobin and Myoglobin in their Reactions with Ligands. North-Holland Publishing Co., Amsterdam. Coletta, M., Ascoli, F., Brunori, M., and Traylor, T. G. (1986) pH dependence of carbon monoxide binding to ferrous horseradish peroxidase. J. Biol. Chem. 261, 9811–9814. Tiso, M., Tejero, J., Basu, S., Azarov, I., Wang, X., et al. (2011) Human neuroglobin functions as a redox-regulated nitrite reductase. J. Biol. Chem. 286, 18277–18289. Herold, S. and Kalinga, S. (2003) Metmyoglobin and methemoglobin catalyze the isomerization of peroxynitrite to nitrate. Biochemistry 42, 14036–14046. Patriarca, A., Eliseo, T., Sinibaldi, F., Piro, M. C., Melis, R., et al. (2009) ATP acts as a regulatory effector in modulating structural transitions of cytochrome c: implications for apoptotic activity. Biochemistry 48, 3279–3287. Patriarca, A., Polticelli, F., Piro, M. C., Sinibaldi, F., Mei, G., et al. (2012) Conversion of cytochrome c into a peroxidase: inhibitory mechanisms and implication for neurodegenerative diseases. Arch. Biochem. Biophys. 522, 62–69. Matsui, T., Ozaki, S.-i., Liong, E., Phillips, G. N. Jr., and Watanabe, Y. (1999) Effects of the location of distal histidine in the reaction of myoglobin with hydrogen peroxide. J. Biol. Chem. 29, 2838–2844. Traylor, T. G. and Sharma, V. S. (1992) Why NO? Biochemistry 31, 2847– 2849. Lawson, D. M., Stevenson, C. E., Andrew, C. R., and Eady, R. R. (2000) Unprecedented proximal binding of nitric oxide to heme: implications for guanylate cyclase. EMBO J. 19, 5661–5671. Tyurina, Y. Y., Poloyac, S. M., Tyurin, V. A., Kapralov, A. A., Jiang, J., et al. (2014) A mitochondrial pathway for biosynthesis of lipid mediators. Nat. Chem. 6, 542–552. Kapralov, A. A., Yanamala, N., Tyurina, Y. Y., Castro, L., Samhan-Arias, A., et al. (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. Rajagopal, B. S., Silkstone, G. G., Nicholls, P., Wilson, M. T., and Worrall, J. A. (2012) An investigation into a cardiolipin acyl chain insertion site in cytochrome c. Biochim. Biophys. Acta 1817, 780–791. Rajagopal, B. S., Edzuma, A. N., Hough, M. A., Blundell, K. L., Kagan, V. E., et al. (2013) The hydrogen-peroxide-induced radical behaviour in human cytochrome c-phospholipid complexes: implications for the enhanced proapoptotic activity of the G41S mutant. Biochem. J. 456, 441–452. Price, N. C. and Stevens, L. (1999) Fundamentals of Enzymology: The Cell and Molecular Biology of Catalytic Proteins, 3rd edn. Oxford University Press, Oxford. Muenzner, J., Toffey, J. R., Hong, Y., and Pletneva, E. V. (2013) Becoming a peroxidase: cardiolipin-induced unfolding of cytochrome c. J. Phys. Chem. B 117, 12878–12886. Hong, Y., Muenzner, J., Grimm, S. K., and Pletneva, E. V. (2012) Origin of the conformational heterogeneity of cardiolipin-bound cytochrome c. J. Am. Chem. Soc. 134, 18713–18723.

108

[79] McClelland, L. J., Mou, T. C., Jeakins-Cooley, M. E., Sprang, S. R., and Bowler, B. E. (2014) Structure of a mitochondrial cytochrome c conformer competent for peroxidase activity. Proc. Natl. Acad. Sci. USA 111, 6648– 6653. [80] Sinibaldi, F., Mei, G., Polticelli, F., Piro, M.C., Howes, B.D., et al. (2005) ATP specifically drives refolding of non-native conformations of cytochrome c. Protein Sci. 14, 1049–1058. [81] Pecina, P., Borisenko, G. G., Belikova, N. A., Tyurina, Y. Y., Pecinova, A., et al. (2010) Phosphomimetic substitution of cytochrome c tyrosine 48 decreases respiration and binding to cardiolipin and abolishes ability to trigger downstream caspase activation. Biochemistry 49, 6705–6714. [82] H€ uttemann, M., Lee, I., Grossman, L. I., Doan, J. W., and Sanderson, T. H. (2012) Phosphorylation of mammalian cytochrome c and cytochrome c oxidase in the regulation of cell destiny: respiration, apoptosis, and human disease. Adv. Exp. Med. Biol. 748, 237–264. [83] Vlasova, I. I., Tyurin, V. A., Kapralov, A. A., Kurnikov, I. V., Osipov, A. N., et al. (2006) Nitric oxide inhibits peroxidase activity of cytochrome c-cardiolipin complex and blocks cardiolipin oxidation. J. Biol. Chem. 281, 14554– 14562. [84] Wilson, M. T., Brunori, M., Rotilio, G. C., and Antonini, E. (1973) Properties of modified cytochromes. II. Ligand binding to reduced carboxymethyl cytochrome c. J. Biol. Chem. 248, 8162–8169. [85] Ascenzi, P., Santucci, R., Coletta, M., and Polticelli, F. (2010) Cytochromes: reactivity of the “dark side” of the heme. Biophys. Chem. 152, 21–27. [86] Ascenzi, P., Leboffe, L., and Polticelli, F. (2013) Reactivity of the human hemoglobin “dark side”. IUBMB Life 65, 121–126. [87] Yang, F., Phillips, G. N. Jr. (1996) Crystal structures of CO-, deoxy- and metmyoglobins at various pH values. J. Mol. Biol. 256, 762–774. [88] Brucker, E. A., Olson, J. S., Ikeda-Saito, M., and Phillips, G. N. Jr. (1998) Nitric oxide myoglobin: crystal structure and analysis of ligand geometry. Proteins 30, 352–356. [89] Hoshino, M., Maeda, M., Konishi, R., Seki, H., and Ford, P. C. (1996) Studies on the reaction mechanism for reductive nitrosylation of ferrihemoproteins in buffer solutions. J. Am. Chem. Soc. 118, 5702–5707. [90] Brunori, M., Giuffre`, A., and Sarti, P. (2005) Cytochrome c oxidase, ligands and electrons. J. Inorg. Biochem. 99, 324–336. [91] Dibrova, D. V., Cherepanov, D. A., Galperin, M. Y., Skulachev, V. P., and Mulkidjanian, A. Y. (2013) Evolution of cytochrome bc complexes: from membrane-anchored dehydrogenases of ancient bacteria to triggers of apoptosis in vertebrates. Biochim. Biophys. Acta 1827, 1407–1427. [92] Kapralov, A. A., Kurnikov, I. V., Vlasova, I. I., Belikova, N. A., Tyurin, V. A., et al. (2007) The hierarchy of structural transitions induced in cytochrome c by anionic phospholipids determines its peroxidase activation and selective peroxidation during apoptosis in cells. Biochemistry 46, 14232–14244. [93] Aluri, H. S., Simpson, D. C., Allegood, J. C., Hu, Y., Szczepanek, K., et al. (2014) Electron flow into cytochrome c coupled with reactive oxygen species from the electron transport chain converts cytochrome c to a cardiolipin peroxidase: role during ischemia-reperfusion. Biochim. Biophys. Acta 1840, 3199–3207. [94] Zou, H., Li, Y., Liu, X., and Wang, X. (1999) An APAF-1
cytochrome c multimeric complex is a functional apoptosome that activates procaspase-9. J. Biol. Chem. 274, 11549–11556. € ttemann, M., Pecina, P., Rainbolt, M., Sanderson, T. H., Kagan, V. E., [95] Hu et al. (2011) 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. [96] Santucci, R., Sinibaldi, F., Polticelli, F., and Fiorucci, L. (2014) Role of cardiolipin in mitochondrial diseases and apoptosis. Curr. Med. Chem. 21, 2702–2714. [97] Zhao, Y., Wang, Z. B., and Xu, J. X. (2003) Effect of cytochrome c on the generation and elimination of O2•2 and H2O2 in mitochondria. J. Biol. Chem. 278, 2356–2360. [98] Min, L. and Jian-Xing, X. (2007) Detoxifying function of cytochrome c against oxygen toxicity. Mitochondrion 7, 13–16. ~ oz-Pinedo, C., Castro, L., Cardaci, S., Schonhoff, C. M., [99] Godoy, L. C., Mun et al. (2009) Disruption of the M80-Fe ligation stimulates the translocation of

Cardiolipin Modulates Cytochrome C Actions

cytochrome c to the cytoplasm and nucleus in nonapoptotic cells. Proc. Natl. Acad. Sci. USA 106, 2653–2658. [100] Yin, H. and Zhu, M. (2012) Free radical oxidation of cardiolipin: chemical mechanisms, detection and implication in apoptosis, mitochondrial dysfunction and human diseases. Free Radic. Res. 46, 959–974. [101] Zhao, K., Zhao, G. M., Wu, D., Soong, Y., Birk, A. V., et al. (2004) Cell-permeable peptide antioxidants targeted to inner mitochondrial membrane inhibit mitochondrial swelling, oxidative cell death, and reperfusion injury. J. Biol. Chem. 279, 34682–34690. [102] Thadani, U. and Whitsett, T. (1988) Relationship of pharmacokinetic and pharmacodynamics properties of the organic nitrates. Clin. Pharmacokinet. 15, 32–43. [103] Ghafourifar, P., Parihar, M. S., Nazarewicz, R., Zenebe, W. J., and Parihar, A. (2008) Detection assays for determination of mitochondrial nitric oxide synthase activity: advantages and limitations. Methods Enzymol. 440, 317–334. [104] Lundberg, J. O., Weitzberg, E., and Gladwin, M. T. (2008) The nitratenitrite-nitric oxide pathway in physiology and therapeutics. Nat. Rev. Drug Discov. 7, 156–167. [105] Everse, J. and Coates, P. W. (2009) Neurodegeneration and peroxidases. Neurobiol. Aging 30, 1011–1025. [106] Hoye, A. T., Davoren, J. E., Wipf, P., Fink, M. P., and Kagan, V. E. (2008) Targeting mitochondria. Acc. Chem. Res. 41, 87–97. [107] Szeto, H. H. (2014) First-in-class cardiolipin-protective compound as a therapeutic agent to restore mitochondrial bioenergetics. Br. J. Pharmacol. 171, 2029–2050.

Ascenzi et al.

[108] Szeto, H. H., James, L. P., and Atkinson, A. J. (2014) Mitochondrial pharmacology: its future is now. Clin. Pharmacol. Ther. 96, 629–633. [109] Prasad, S., Maiti, N. C., Mazumdar, S., and Mitra, S. (2002) Reaction of hydrogen peroxide and peroxidase activity in carboxymethylated cytochrome c: spectroscopic and kinetic studies. Biochim. Biophys. Acta 1596, 63–75. [110] Roncone, R., Monzani, E., Murtas, M., Battaini, G., Pennati, A., et al. (2004) Engineering peroxidase activity in myoglobin: the haem cavity structure and peroxide activation in the T67R/S92D mutant and its derivative reconstituted with protohaemin-l-histidine. Biochem J. 377, 717– 724. [111] Santucci, R., Sinibaldi, F., Patriarca, A., Santucci, D., and Fiorucci, L. (2010) Misfolded proteins and neurodegeneration: role of non-native cytochrome c in cell death. Expert Rev. Proteomics 7, 507–517. [112] Ghavami, S., Shojaei, S., Yeganeh, B., Ande, S. R., Jangamreddy, J. R., et al. (2014) Autophagy and apoptosis dysfunction in neurodegenerative disorders. Prog. Neurobiol. 112, 24–49. [113] Birk, A. V., Chao, W. M., Bracken, C., Warren, J. D., and Szeto, H. H. (2014) Targeting mitochondrial cardiolipin and the cytochrome c/cardiolipin complex to promote electron transport and optimize mitochondrial ATP synthesis. Br. J. Pharmacol. 171, 2017–2028. [114] Jiang, J., Bakan, A., Kapralov, A. A., Ishara Silva, K., Huang, Z., et al. (2014) Designing inhibitors of cytochrome c/cardiolipin peroxidase complexes: mitochondria-targeted imidazole-substituted fatty acids. Free Radic. Biol. Med. 71, 221–230.

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