Báo cáo khoa học: Probing the access of protons to the K pathway in the Paracoccus denitrificans cytochrome c oxidase pdf

Probing the access of protons to the K pathway in the Paracoccus denitrificans cytochrome c oxidase Oliver-M. H. Richter 1 , Katharina L. Du ¨ rr 1 , Aimo Kannt 2 , Bernd Ludwig 1 , Francesca M. Scandurra 3 , Alessandro Giuffre ` 3 , Paolo Sarti 3 and Petra Hellwig 4 1 Institut fu ¨ r Biochemie, Abteilung Molekulare Genetik, Johann Wolfgang Goethe-Universita ¨ t, Frankfurt-am-Main, Germany 2 Max-Planck-Institut fu ¨ r Biophysik, Abteilung Molekulare Membranbiologie, Frankfurt-am-Main, Germany 3 Department of Biochemical Sciences and CNR Institute of Molecular Biology and Pathology, University of Rome ‘La Sapienza’, Rome, Italy 4 Institut fu ¨ r Biophysik, Johann Wolfgang Goethe-Universita ¨ t, Frankfurt-am-Main, Germany Cytochrome c oxidase of Paracoccus denitrificans [1–4] resides in the cytoplasmic membrane of this soil bac- terium and reduces molecular oxygen to water. In a process still poorly understood, the free energy of the redox reaction is exploited to translocate protons across the membrane. Electrons donated by cyto- chrome c are first transferred to a homobinuclear cop- per center (Cu A ) located in the periplasmic domain of subunit II. Subsequently heme a and another binuclear center consisting of heme a 3 and Cu B , all constituents of subunit I, become reduced. Protons needed for water formation and those to be translocated across the membrane are taken up from the inner side by two different pathways that were identified initially by site- directed mutagenesis experiments [5–7] and were con- firmed in the X-ray structures resolved so far [3,8–11]. A considerable line of evidence suggests that not only those protons that are pumped across the membrane, but also most of the protons required for the forma- tion of water are delivered via the so called D channel (named after a functionally critical aspartate residue at its entrance, D124 in Paracoccus numbering) [12–14]. The separate K channel is indispensable for proton transfer linked to the reduction of the binuclear site and, consistently, mutations in this channel give rise to a drastically retarded reduction of heme a 3 [12,15–17]. In contrast to the aspartate of the D channel, the cru- cial lysine residue (K354 in Paracoccus) is located well within the membrane dielectric [3]. Mutation of resi- dues close to the K channel entrance do not elicit the Keywords cytochrome c oxidase; FTIR; proton channel; electron transfer; site-directed mutagenesis Correspondence O M. H. Richter, Institut fu ¨ r Biochemie, Abteilung Molekulare Genetik, Johann Wolfgang Goethe-Universita ¨ t, Germany Fax: +49 79 829244 Tel: +49 79 829240 E-mail: O.M.Richter@em.uni-frankfurt.de (Received 23 August 2004, revised 1 November 2004, accepted 12 November 2004) doi:10.1111/j.1742-4658.2004.04480.x In recent studies on heme-copper oxidases a particular glutamate residue in subunit II has been suggested to constitute the entry point of the so-called K pathway. In contrast, mutations of this residue (E78 II ) in the Paracoccus denitrificans cytochrome c oxidase do not affect its catalytic activity at all (E78 II Q) or reduce it to about 50% (E78 II A); in the latter case, the mutation causes no drastic decrease in heme a 3 reduction kinetics under anaerobic con- ditions, when compared to typical K pathway mutants. Moreover, both mutant enzymes retain full proton-pumping competence. While oxidized- minus-reduced Fourier-transform infrared difference spectroscopy demon- strates that E78 II is indeed addressed by the redox state of the enzyme, absence of variations in the spectral range characteristic for protonated aspartic and glutamic acids at  1760 to 1710 cm )1 excludes the protonation of E78 II in the course of the redox reaction in the studied pH range, although shifts of vibrational modes at 1570 and 1400 cm )1 reflect the reorganization of its deprotonated side chain at pH values greater than 4.8. We therefore conclude that protons do not enter the K channel via E78 II in the Paracoccus enzyme. Abbreviations SHE¢, standard hydrogen electrode (at pH 7); SVD, singular value decomposition; VIS, visible. 404 FEBS Journal 272 (2005) 404–412 ª 2004 FEBS expected inhibition of the enzyme (for example S291 in Paracoccus [7], see also [18]). A possible explanation was given by electrostatic calculations indicating that the redox state of the binuclear center influences the protonation state of a glutamate residue positioned at the cytoplasmic end of the second helix of subunit II (E78 in Paracoccus) [19], subsequently considered a potential K channel entrance residue (see below). Experiments were first performed with mutants in the corresponding position of the Escherichia coli bo 3 quinol oxidase (E89 II Q ⁄ A ⁄ D) where a considerable drop in oxidase activity was observed (to 10, 43 and 60%, respectively), and enzymes with the A or Q mutation could not support aerobic growth of strains devoid of other terminal oxidases on nonfermentable substrates [20]. It is not clear on the other hand whe- ther this inability to sustain growth reflects an impaired proton pump or is a consequence of the reduced enzymatic activity. In the E. coli bo 3 oxidase, mutation of E89 II to Q has been proposed to block the reduction of the heme-copper binuclear center, lending support to the idea that E89 II is actually at the entrance to the K channel or at least critically import- ant for its functionality. An even broader range of substitutions were introduced at the corresponding glu- tamate (E101) into the cytochrome c oxidase of Rho- dobacter sphaeroides [18,21]. There, oxidase activity is reduced even more drastically in all mutant enzymes (14, 8 and 16% residual activity for E101Q ⁄ A ⁄ D, respectively), with the highest activity of about 30% retained in E101H. In addition, a correlation was observed between the diminished steady-state activity and the slower rate of heme a 3 reduction under anaer- obic conditions, discussed as an impairment of proton delivery through the K channel. As mutations of E101 fulfill criteria that have also been observed with bona fide K channel mutants (for example K362 in Rhodo- bacter), E101 has been considered to be the dominant entry point for protons going into the K channel [21]. Electrochemically induced FTIR difference spectros- copy is a tool to study the reorganization of a protein upon electron transfer and concomitant proton trans- fer. The spectra reveal the contributions of residues addressed by the redox reaction and are specific for their protonation state. For the cytochrome c oxidase from P. denitrificans the protonation state of E278 [22], of individual heme propionates [23,24] and of Y280 [25] was previously studied employing this tech- nique in combination with site-directed mutants and isotopic labeling. Here mutants of E78 II in the Paracoccus cyto- chrome c oxidase are characterized in terms of cata- lytic and proton pumping activity, kinetics of heme a 3 reduction and electrochemically induced FTIR differ- ence spectroscopy. Based on the modest effects caused by the mutations, it is concluded that in the Paracoc- cus oxidase E78 II does not play the critical role documented in the case of the E. coli and the R. sphaero-ides oxidases. Results Enzymatic turnover and electron transfer Mutations of position E78 in subunit II of the Para- coccus cytochrome c oxidase were introduced to probe the relevance of this residue for the catalytic properties of this enzyme in general, and to monitor spectros- copic changes that result from the substitution of this residue to either glutamine or alanine. Mutant oxidases were purified and characterized by VIS redox spectroscopy and SDS gel electrophoresis. Essentially no differences were observed in compar- ison with wild-type oxidase with respect to heme and subunit composition (not shown). The cytochrome c oxidase activity of E78 II Q matched that of wild-type, while the activity of the E78 II A mutant was lowered to about 50% (Table 1). Given the lower activity of the E78 II A mutant, the kinetics of reduction of this mutant were investigated by stopped-flow spectrophotometry by anaerobically mixing a degassed sample of the mutant in the oxidized state with a large excess of ascorbate and ruthenium hexamine (20 and 1 mm after mixing, respectively). Within a few milliseconds after mixing a fast reduction of heme a occurs (data not shown), similarly to what was reported for the wild-type enzyme and the K354M mutant under identical experimental conditions [26]. On a longer time scale (from 20 ms to 20 s), heme a 3 becomes reduced and the corresponding typical absorp- tion changes are consistently observed (Fig. 1, top panel). Table 1. Enzymatic activity and proton-pumping capacity of wild- type and mutant oxidases isolated from Paracoccus denitrificans. Reconstituted oxidases were measured either with a stopped-flow apparatus or potentiometrically (for details see Experimental proce- dures). Enzymatic activity has been determined with samples from the same oxidase preparations. 100% activity corresponds to a turn over of 325 electrons s )1 . Oxidase-type Enzymatic activity (%) H + ⁄ e – WT 100 1.04 a ⁄ 1.0 b E78 II A 50 1.10 a E78 II Q921.0 b a Stopped-flow apparatus; b potentiometric method. O M. H. Richter et al. Proton access to Paracoccus cytochrome c oxidase FEBS Journal 272 (2005) 404–412 ª 2004 FEBS 405 The time course of heme a 3 reduction in the E78 II A mutant as obtained by global fitting analysis of the observed absorption changes is depicted in Fig. 1 (bot- tom panel). By comparison with the wild-type enzyme, it is evident that, unlike the K354M mutation, the E78 II A mutation only slightly affects the rate of heme a 3 reduction. The observed kinetic effect is remarkably smaller than the one reported for the same mutation at the corresponding residue in the R. sph- aeroides oxidase [21]. In all Paracoccus samples heme a 3 reduction is biphasic, similarly to what was previ- ously observed with the beef heart enzyme [27]. The existence of two kinetic phases associated to identical absorption changes demands further investigation, although it can be tentatively assigned to two enzyme subpopulations with different kinetic properties. In the case of the Paracoccus wild-type enzyme, the observed fitted rate constant relative to the major kinetic phase ( 19 s )1 corresponding to  70% of the reaction amplitude) is about four-fold lower than the turnover number for O 2 consumption ( 80 mol O 2 Æmol 6 enzy- me )1 Æs )1 ). As previously shown for the Paracoccus enzyme [26], this is expected given the unfavorable redox equilibrium between heme a and heme a 3 ,ifitis taken into account that our measurements were carried out in the absence of NO acting as a trapping ligand for reduced heme a 3 [27]. Proton-pumping activity To complement these results on electron transfer with proton-pumping measurements, purified mutant oxidases were reconstituted into phospholipid vesicles by the cholate dialysis method (see Experimental pro- cedures). Both the E78 II Q and the E78 II A mutant oxidases show unimpaired proton-pumping (Table 1). The H + ⁄ e – ratio for E78 II Q was found to be around 1.0 when determined by the reductant pulse potentio- metric method [28]. The E78 II A mutant was tested in a stopped-flow approach monitoring the absorbance change of the pH-sensitive dye phenol red [29]. Experi- ments in the absence and presence of the uncoupler CCCP gave a H + ⁄ e – ratio of 0.9 very similar to that of the wild-type enzyme (Fig. 2). Due to slight varia- tions during oxidase reconstitution, proteoliposomes with incorporated E78 II A oxidase result in a faster proton ejection than those with reconstituted wild- type, although activity measurements clearly show a diminished activity for this mutant (Table 1). Electrochemically induced FTIR difference spectroscopy As the above experiments do not provide any informa- tion about structural details of the E78 II A and E78 II Q mutant enzymes, electrochemically induced FTIR difference spectra of the corresponding cytochrome c oxidases were recorded to detect molecular changes concomitant with the redox reaction. This approach allows monitoring of conformational changes or Fig. 1. Kinetics of heme a 3 reduction of wild-type and mutant P. denitrificans oxidase. Degassed samples of the oxidized enzyme (wild-type, E78 II A and K354M) were anaerobically mixed with ascor- bate and ruthenium hexamine (20 and 1 m M after mixing, respect- ively) at 20 °C. Under these conditions the reduction of heme a is complete within a few milliseconds, followed by the reduction of heme a 3. (Top panel) Absorption changes collected from 20 ms to 20 s after mixing the E78 II A with the reductants (baseline: endpoint spectrum acquired at 20 s). Within the experimental error singular value decomposition (SVD) analysis of this spectra set yields only one significant U-column, corresponding to the ox-red spectrum of heme a 3 (inset). (Bottom panel) Time courses of heme a 3 reduction as obtained by SVD analysis. Fitted rate constants with relative amplitudes in brackets: wild-type enzyme, k 1 ¼ 18.7 s )1 (70%), k 2 ¼ 1.1 s )1 (30%); E78 II A mutant, k 1 ¼ 14.5 s )1 (35%), k 2 ¼ 0.38 s )1 (65%); K354M mutant, k 1 ¼ 0.09 s )1 (30%), k 2 ¼ 0.005 s )1 (70%). Data on the K354M mutant are from [34]. Proton access to Paracoccus cytochrome c oxidase O M. H. Richter et al. 406 FEBS Journal 272 (2005) 404–412 ª 2004 FEBS charge redistributions at the cofactor sites, reflecting the reorganization of the hemes, of the polypeptide backbone and of the amino acid side chains upon electron transfer to ⁄ from the redox active centers (hemes a ⁄ a 3 ,Cu B or Cu A ). Additionally proton reac- tions concomitant with electron transfer are expected to contribute to the spectra. The electrochemically induced FTIR difference spectra of wild-type cyto- chrome c oxidase were previously published and dis- cussed in detail [30,31]. Figure 3 shows the oxidized-minus-reduced FTIR difference spectra of the E78 II Q (A) and E78 II A (B) mutant enzymes (full line) in direct comparison to wild-type (dotted line) for a potential step from )0.29 to 0.71 V at pH 7. A clear decrease of the negative mode concomitant with the reduced form at 1546 cm )1 can be seen for both mutants in direct comparison to wild-type. Small shifts are present at 1720, 1676, 1638, 1619, 1554 and 1390 cm )1 . No major variations, how- ever, are present, demonstrating that the overall struc- ture of the proteins is not affected upon mutation. In order to distinguish the shifts, double difference spec- tra have been obtained via subtraction of the differ- ence spectra for the mutant enzymes from wild-type (C). The glutamine of E78 II Q may contribute upon electron transfer: contributions of the m(C¼O) vibra- tional mode of glutamines can be expected at 1668 to 1687 cm )1 and of the d(NH 2 ) at 1585 to 1611 cm )1 Fig. 3. FTIR difference spectroscopy of E78 II Q and E78 II A variants. Oxidized-minus-reduced FTIR difference spectra of the E78 II Q(A) and E78 II A (B) mutant enzymes (line), each in comparison to wild- type (dotted line) for a potential step from )0.29 to 0.71 V at pH 7. The double difference spectra were obtained via subtraction of the difference spectra for the E78 II Q (C, line) and E78 II A (C, dotted line) mutant enzymes from wild-type (C). For experimental details see Experimental procedures. Fig. 2. Proton translocation of reconstituted E78 II A (A) and wild- type (B) Paracoccus oxidase. Reconstituted enzyme was mixed aero- bically in the presence and absence of the uncoupler CCCP, using the indicator dye phenol red to monitor pH changes in the cuvette. Negative excursion denotes an acidification of the exter- nal medium. For details see Experimental procedures. O M. H. Richter et al. Proton access to Paracoccus cytochrome c oxidase FEBS Journal 272 (2005) 404–412 ª 2004 FEBS 407 [32]. The increase of a difference signal at 1676 and at 1608 cm )1 , however, is also observed for the E78 II A mutant and rather seems to reflect a small structural variation induced by each amino acid substitution (see below). The decrease of the negative mode at 1546 cm )1 and of the positive mode at 1554 cm )1 that can be seen comparing the difference spectra for mutant and wild- type enzyme, occurs in the spectral region where the m(COO – ) as from deprotonated glutamic acids are expected. The corresponding m(COO – ) s modes are usu- ally significantly smaller and may be involved in the variations around 1390 cm )1 . The changes of these dif- ferential signals upon mutation indicate that E78 II is deprotonated at the given pH conditions (phosphate buffer, pH 7) and reorganizes upon redox reaction. We note that these modes are present in a complex spec- tral range, where also the amide II mode of secondary structure elements contribute. An effect on the m(C¼C) vibrations of the hemes, which are also included in that spectral region, however, seems unlikely on the basis of the distance of the mutation from the heme porphyrin rings. Protonated acidic residues characteristically contri- bute above 1710 cm )1 . Only a very small and broad variation is seen at 1720 cm )1 , which would be typ- ical for a residue at the surface due to conformation- al flexibility of the carboxylic side chain. The intensity of this variation is significantly smaller than what would be expected for a protonation reaction. To assess whether this variation reflects partial pro- tonation of the E78 II side chain at pH 7, electro- chemically induced FTIR difference spectra were obtained for the E78 II Q mutant enzyme, equilibrated at pH 4.8 in cacodylate buffer. Figure 4 shows the electrochemically induced FTIR difference spectra at pH 7 (solid line) and 4.8 (dotted line) of wild-type (A) and compares it to the E78 II Q mutant enzyme equilibrated at the same pH values (B) as well as the direct comparison of wild-type (dotted line) with the E78 II Q mutant (solid line) at pH 4.8 (C). Interestingly a broad positive mode at 1718 cm )1 and strong negative modes at 1556 and 1402 cm )1 show the largest deviation between each pair of spectra, for both wild-type and mutant. The signal at 1718 cm )1 most likely arises from m(C¼O) modes of the protonat- ed form of carboxylic groups and the modes at 1556 and 1402 cm )1 from the m(COO – ) s ⁄ as vibrational modes of the corresponding deprotonated form. The relation of the extinction coefficients of these modes are close to model compound studies on proto- nation ⁄ deprotonation of isolated acidic amino acids in solution [32]. We attribute these modes to the proto- nation of carboxylic groups upon oxidation at pH 4.8, the group being deprotonated above pH 5. The band width of the contribution at  1718 cm )1 indicates that residues close to the surface, or in the vicinity of sev- eral water molecules are involved here. Comparing the spectra of the E78 II Q mutant and wild-type for both pH values (Fig. 4), no additional pH-dependent varia- tions can be seen, which are not present in the wild- type spectra as well, excluding E78 II to be this residue. This conclusion is supported by the clear decrease at 1546 cm )1 that is not changed for the low pH value (Fig. 4C). Our experimental data show that the pK A Fig. 4. pH dependence of the infrared signals for the E78 II Q vari- ant. Oxidized-minus-reduced FTIR difference spectra at pH 7 (full line) and 4.8 (dotted line) of wild-type (A) and E78 II Q (B) cyto- chrome c oxidase for a potential step from )0.29 to 0.71 V as well as the direct comparison between wild-type (dotted line) and E78 II Q mutant at pH 4.8 (C). Proton access to Paracoccus cytochrome c oxidase O M. H. Richter et al. 408 FEBS Journal 272 (2005) 404–412 ª 2004 FEBS value of E78 is thus below 5 and a protonation of the residue is not expected at pH values above 5. Discussion Electrostatic calculations identified a glutamate (E78) as a redox-responsive residue in subunit II of the P. denitrificans cytochrome c oxidase [19]. It is located at the cytoplasmic side of helix II at a distance of 9 A ˚ to the lysine of the K channel (K354) [3]. Subsequently this residue gained considerable attention due to a potential importance for access of protons into the K channel of heme-copper oxidases. Glutamate residues in equivalent positions of other oxidases (E89 in the E. coli bo 3 ubiquinol oxidase [20] and E101 in the R. sphaeroides aa 3 cytochrome c oxid- ase [21], were mutated, amongst others, to A and Q, revealing clear defects in their catalytic properties. The activity in the A and Q mutants was diminished con- siderably, namely to 43 and 10% (E. coli bo 3 ) and to 8 and 14% (R. sphaeroides aa 3 ), respectively. Both E. coli mutants failed to complement aerobic growth on nonfermentable substrates (in the absence of other oxidases) which unfortunately cannot be correlated directly with an impaired proton pump without addi- tional information. No corresponding information is available for the Rhodobacter mutants. The observed mutational effects were assigned to a lowered rate of reduction of the binuclear center caused by an impaired proton transfer through the K channel. Our results obtained with the corresponding muta- tions in the P. denitrificans cytochrome c oxidase clearly contradict those summarized above. Although we observe a modest reduction of the catalytic activity in E78 II A to about 50% of the wild-type level, the E78 II Q mutation shows unchanged catalytic compet- ence. More importantly, proton-pumping is essentially unaffected in both mutant enzymes. It therefore seems that E78 II lacks a direct role in the overall enzymatic reaction of the Paracoccus enzyme in contrast even to the closely related R. sphaeroides aa 3 oxidase [21]. Interestingly, mutation of E78 II to A causes only a slight effect on the kinetics of heme a 3 reduction, much smaller than that caused by mutation of other residues widely accepted as belonging to the K channel, like K354 [26]. Our measurements were carried out under anaerobic conditions at high reductant concentration, i.e. under conditions in which the very fast reduction of heme a does not limit the overall reduction rate of the enzyme [26,27]. Given that a drastically lowered rate of heme a 3 reduction is considered as diagnostic for mutated residues in the K channel, we conclude that in the P. denitrificans aa 3 cytochrome c oxidase E78 II does not represent the dominant entry point for protons into the K-channel. We used redox-induced FTIR difference spectroscopy to monitor whether E78 II is addressed by the redox reaction at all. At given pH conditions, signals characteristic for a deprotonated carboxylic group were identified. Both mutants lead to similar changes of nearby residues revealing small structural variations induced in the enzyme. Based on their full proton-pumping activity and the moderate effect on enzymatic turnover activit- ies, as well as the reduction kinetics for E78 II A, we conclude that this reorganization of E78 carboxylate upon the redox reaction is, however, without direct implications on the catalytic cycle in the cytochrome c oxidase from P. denitrificans. It is difficult to reconcile the discrepancy observed with mutations of the particular glutamate residue, especially between the closely related aa 3 cytochrome c oxidases of Paracoccus and Rhodobacter. S291 as an alternative for the entrance to the K channel of the Paracoccus oxidase as deduced from the X-ray struc- ture [3], has been addressed by mutation before [33], however, without effect on the overall enzymatic reac- tion. It therefore seems that funneling of protons into the K channel is just one of several examples of differences between even closely related terminal oxidases that operate on a common mechanistic ground but allow for a certain degree of flexibility in the design of indi- vidual mechanistic steps. Experimental procedures Mutagenesis and cloning Site-directed mutations of subunit II were introduced with the Altered Sites Ò system employing pAlter-1 (Promega), and subsequently confirmed by sequencing. The mutated subunit II gene together with the rest of the cta operon [34] was cloned as a XhoI ⁄ HindIII fragment into appropriately cut pUP39 [33] that allows for replication in Paracoccus strain ST4 [35] where most of the cta operon from ctaCto ctaE had been replaced by a kanamycin resistance gene. Cell growth and protein purification Growth of the Paracoccus strains and purification of the oxidases with a tagged F v antibody fragment was per- formed as described [33,36]. For electrochemistry the pro- tein samples were further concentrated to  0.5 mm aa 3 using Microcon ultrafiltration cells (Millipore) and 200 mm phosphate (pH 7), or 200 mm cacodylate buffer (pH 4.8), both containing 100 mm KCl and 0.05% n dodecyl-b- O M. H. Richter et al. Proton access to Paracoccus cytochrome c oxidase FEBS Journal 272 (2005) 404–412 ª 2004 FEBS 409 d-maltopyranoside. For proton-pumping experiments the oxidases were bound to 2 mL columns of Q-sepharose (Amersham), washed with 10 mm Hepes pH 7.3, 50 mm KCl, 0.015% dodecylmaltoside and eluted with 500 mm KCl in the same buffer. Cytochrome c oxidase activity Enzymatic activity was determined at room temperature with 20 lm reduced horse heart cytochrome c (Sigma) at 550 nm with a Hitachi U-3000 spectrophotometer. The reaction buffer contained 20 mm Tris ⁄ HCl pH 7.5, 20 mm KCl, 1 mm EDTA and 0.02% dodecylmaltoside. Electron transfer measurements The kinetics of reduction of the enzyme were investigated at 20 °C by using a stopped-flow apparatus (DX.17MV, Applied Photophysics, Leatherhead, UK), equipped with a photodiode-array (light path ¼ 1 cm). Absorption spectra were collected with an acquisition time of 2.56 ms up to 20 s after mixing. Buffer: 20 mm phosphate pH 7.0, 50 mm NaCl, 0.1% dodecylmaltoside. Anaerobic conditions were obtained by extensive N 2 -equilibration and contaminant oxygen was further scavenged by addition of glucose (2 mm), glucose oxidase (8 unitsÆmL )1 ) and catalase (260 unitsÆmL )1 ), immediately before the experiment. Data were analyzed by using the singular value decomposition (SVD) algorithm implemented in the software matlab (Math- Works, Natick, MA, USA). Time courses were fitted to the sum of two exponentials. Reconstitution of purified oxidase into liposomes Asolectin (40 mgÆmL )1 , Sigma, type IV-S) and 2% cholate, both purified according to [37], were dissolved in 100 mm Hepes pH 7.3, 10 mm KCl and, after stirring for 1 h, were sonified (both steps under a nitrogen atmosphere) at inter- vals of 30 s (Branson Sonifier II 250, output 5, 50% duty cycle) until clarification of the suspension. After a brief centrifugation (6000 g, 15 min, 4 °C) to remove particulate material, purified oxidase (either wild-type or mutant enzyme, see above) was added to a final concentration of 4 lm and the solution subjected to dialysis essentially as described [28]. Protein aggregates were removed by centrifu- gation (6000 g, 15 min, 4 °C) and the resulting liposome suspension stored at 4 °C. Proton translocation of reconstituted oxidase A suspension containing 0.4 lm E78 II A or wild-type oxidase proteoliposomes, 60 l m phenol red and 10 lm valinomycin was prepared in the last dialysis buffer (see above, the pH was readjusted to 7.3, if necessary), and filled into a 2.5-mL syringe of the stopped-flow apparatus (Hi-Tech Scientific, SF-61). For determination of the decoupled rates, 10 lm of CCCP was added. Reduced horse heart cytochrome c was brought to 200 lm with the last dialysis buffer (see above). After adjusting the pH to 7.3 the solution was filled in a 0.25- mL syringe. The 10 : 1 ratio of syringes was chosen to avoid mixing artifacts [28,29]. The absorbance change of phenol red was monitored at 25 °C and 555.6 nm, which was deter- mined to be the isosbestic point for cytochrome c under the experimental conditions. E78 II Q proton translocation was measured potentiometrically as described [28]. Electrochemistry An ultra-thin layer spectroelectrochemical cell for the VIS and IR was used as described previously [38]. Sufficient transmission in the 1800–1000 cm )1 range, even in the region of strong water absorbance around 1645 cm )1 , was achieved with the cell path-length set to 6–8 lm. The gold grid work- ing electrode was chemically modified with a 2-mm cysteam- ine solution and different mediators were added as reported before [22] to a final concentration of 45 lm each (leaving out N-methyl- and N-ethyl-phenazoniumsulfate, but adding neutral red; E m : )307 mV) to accelerate the redox reaction. At this concentration, and with the cell pathlength below 10 lm, no spectral contributions from the mediators in the visible and infrared range could be detected in control experi- ments with samples lacking the protein, except for the PO modes of the phosphate buffer between 1200 and 1000 cm )1 . Potentials were measured with a Ag ⁄ AgCl ⁄ 3M KCl reference electrode and are quoted in reference to SHE¢ (pH 7). Optical spectroscopy FTIR and VIS difference spectra as a function of the applied potential were obtained simultaneously from the same sample with a setup combining an IR beam from the interferometer (modified IFS 25, Bruker, Germany) for the 4000 to 1000 cm )1 range and a dispersive spectro- meter for the 400 to 900 nm range. Electrochemically induced difference spectra were recorded and processed as previously described [22]. Acknowledgements We are indebted to E. Bamberg (Max-Planck-Institut fu ¨ r Biophysik, Frankfurt) for kindly providing techni- cal facilities, to C. Bamann for assistance with the stopped-flow equipment in Frankfurt and to A. Lu ¨ ck and H. Mu ¨ ller for excellent technical assistance. We wish to thank M. Brunori (Rome, Italy) for exten- ded discussions. P. H. thanks W. Ma ¨ ntele (Institut fu ¨ r Biophysik, Frankfurt) for continuous support. Proton access to Paracoccus cytochrome c oxidase O M. H. Richter et al. 410 FEBS Journal 272 (2005) 404–412 ª 2004 FEBS This work was supported by DFG (SFB 472) and by MIUR of Italy (PRIN ‘Bioenergetica: genomica funzionale, meccanismi molecolari ed aspetti fisiopato- logici’ and Fondo per gli Investimenti della Ricerca di Base RBAU01F2BJ to P.S.). References 1 Ludwig B & Schatz G (1980) A two-subunit cytochrome c oxidase (cytochrome aa 3 ) from Paracoccus denitrifi- cans. Proc Natl Acad Sci USA 77, 196–200. 2 Haltia T, Puustinen A & Finel M (1988) The Paracoc- cus denitrificans cytochrome aa 3 has a third subunit. Eur J Biochem 172, 543–546. 3 Iwata S, Ostermeier C, Ludwig B & Michel H (1995) Structure at 2.8 A ˚ resolution of cytochrome c oxidase from Paracoccus denitrificans. Nature 376 , 660–669. 4 Baker SC, Ferguson SJ, Ludwig B, Page MD, Richter O-MH & van Spanning RJM (1998) Molecular genetics of the genus Paracoccus – metabolically versatile bac- teria with bioenergetic flexibility. Microbiol Mol Rev 62, 1046–1078. 5 Thomas JW, Puustinen A, Alben JO, Gennis RB & Wikstro ¨ m M (1993) Substitution of aspartate-135 in subunit I of the cytochrome bo ubiquinol oxidase of Escherichia coli eliminates proton-pumping activity. Biochemistry 32, 10923–10928. 6 Hosler JP, Ferguson-Miller S, Calhoun MW, Thomas JW, Hill J, Lemieux L, Ma J, Georgiou C, Fetter J, Shapleigh J, Tecklenburg MMJ, Babcock GT & Gennis RB (1993) Insight into the active-site structure and func- tion of cytochrome oxidase by site-directed mutants of bacterial cytochrome aa 3 and cytochrome bo. J Bioenerg Biomembr 25, 121–136. 7 Pfitzner U, Odenwald A, Ostermann T, Weingard L, Ludwig B & Richter O-MH (1998) Cytochrome c oxi- dase (heme aa 3 ) from Paracoccus denitrificans: analysis of mutations in putative proton channels of subunit I. J Bioenerg Biomembr 30, 89–97. 8 Tsukihara T, Aoyama H, Yamashita E, Tomashi 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 A ˚ . Science 272, 1136–1144. 9 Abramson J, Riistama S, Larsson G, Jasaitis A, Svens- son-Ek M, Laakkonen L, Puustinen A, Iwata S & Wikstro ¨ m M (2000) The structure of the ubiquinol oxi- dase from Escherichia coli and its ubiquinone binding site. Nat Struct Biol 7, 910–917. 10 Soulimane T, Buse G, Bourenkov GP, Bartunik HD, Huber R & Than ME (2000) Structure and mechanism of the aberrant ba 3 -cytochrome c oxidase from Thermus thermophilus. EMBO J 19, 1766–1776. 11 Svensson-Ek M, Abramson J, Larsson G, To ¨ rnroth S, Brzezinski P & Iwata S (2002) The X-ray crystal struc- tures of wild-type and EQ (I-286) mutant cytochrome c oxidase from Rhodobacter sphaeroides. J Mol Biol 321, 329–339. 12 Konstantinov AA, Siletsky S, Mitchell D & Kaulen A (1997) The roles of the two proton input channels in cytochrome c oxidase from Rhodobacter sphaeroides probed by the effects of site-directed mutations on time- resolved electrogenic intraprotein proton transfer. Proc Natl Acad Sci USA 94, 9085–9090. 13 Brzezinski P & A ¨ delroth P (1998) Pathways of proton transfer in cytochrome c oxidase. J Bioenerg Biomembr 30, 99–107. 14 Ruitenberg M, Kannt A, Bamberg E, Fendler K & Michel H (2002) Reduction of cytochrome c oxidase by a second electron leads to proton translocation. Nature 417, 99–102. 15 A ¨ delroth P, Gennis RB & Brzezinski P (1998) Role of the pathway through K (I-362) in proton transfer in cytochrome c oxidase from R. sphaeroides. Biochemistry 37, 2470–2476. 16 Ruitenberg M, Kannt A, Bamberg E, Ludwig B, Michel H & Fendler K (2000) Single-electron reduction of the oxidized state is coupled to proton uptake via the K pathway in Paracoccus denitrificans cytochrome c oxidase. Proc Natl Acad Sci USA 97, 4632–4636. 17 Forte E, Scandurra FM, Richter O-MH, D’Itri E, Sarti P, Brunori M, Ludwig B & Giuffre ` A (2004) Proton uptake upon anaerobic reduction of the Paracoccus denitrificans cytochrome c oxidase: a kinetic investiga- tion of the K354M and D124N mutants. Biochemistry 43, 2957–2963. 18 Bra ¨ nde ´ n M, Tomson F, Gennis RB & Brzezinski P (2002) The entry point of the K-proton-transfer pathway in cytochrome c oxidase. Biochemistry 41, 10794–10798. 19 Kannt A, Lancaster CRD & Michel H (1998) The role of electrostatic interactions for cytochrome c oxidase function. J Bioenerg Biomembr 30, 81–87. 20 Ma J, Tsatsos PH, Zaslavsky D, Barquera B, Thomas JW, Katsonouri A, Puustinen A, Wikstro ¨ m M, Brzezin- ski P, Alben JO & Gennis RB (1999) Glutamate-89 in subunit II of cytochrome bo 3 from Escherichia coli is required for the function of the heme-copper oxidase. Biochemistry 38, 15150–15156. 21 Tomson FL, Morgan JE, Gu G, Barquera B, Vygodina TV & Gennis RB (2003) Substitutions for glutamate 101 in subunit II of cytochrome c oxidase from Rhodo- bacter sphaeroides result in blocking the proton-conduct- ing K-channel. Biochemistry 42, 1711–1717. 22 Hellwig P, Behr J, Ostermeier C, Richter O-MH, Pfitz- ner U, Odenwald A, Ludwig B, Michel H & Ma ¨ ntele W O M. H. Richter et al. Proton access to Paracoccus cytochrome c oxidase FEBS Journal 272 (2005) 404–412 ª 2004 FEBS 411 (1998) Involvement of glutamic acid 278 in the redox reaction of the cytochrome c oxidase from Paracoccus denitrificans investigated by FTIR spectroscopy. Bio- chemistry 37, 7390–7399. 23 Behr J, Hellwig P, Ma ¨ ntele W & Michel H (1998) Redox dependent changes at the heme propionates in cytochrome c oxidase from Paracoccus denitrificans: direct evidence from FTIR difference spectroscopy in combination with propionate 13 C labelling. Biochemistry 37, 7400–7406. 24 Behr J, Michel H, Ma ¨ ntele W & Hellwig P (2000) Func- tional properties of the heme propionates in cytochrome c oxidase from Paracoccus denitrificans: evidence from FTIR difference spectroscopy and site-directed muta- gensis. Biochemistry 39, 1356–1363. 25 Hellwig P, Pfitzner U, Behr J, Rost B, Pesavento RP, Donk WV, Gennis RB, Michel H, Ludwig B & Ma ¨ ntele W (2002) Vibrational modes of tyrosines in cytochrome c oxidase from Paracoccus denitrificans: FTIR and elec- trochemical studies on Tyr-D4-labeled and on Tyr280- His and Tyr35Phe mutant enzymes. Biochemistry 41, 9116–9125. 26 Giuffre ` A, Barone MC, Brunori M, D’Itri E, Ludwig B, Malatesta F, Mu ¨ ller H-W & Sarti P (2002) Nitric oxide reacts with the single-electron reduced active site of cytochrome c oxidase. J Biol Chem 277, 22402–22406. 27 Brunori M, Giuffre ` A, D’Itri E & Sarti P (1997) Inter- nal electron transfer in Cu-heme oxidases: thermo- dynamic or kinetic control? J Biol Chem 272, 19870– 19874. 28 Kannt A, Soulimane T, Buse G, Becker A, Bamberg E & Michel H (1998) Electrical current generation and proton pumping catalyzed by the ba 3 -type cytochrome c oxidase from Thermus thermophilus. FEBS Lett 434, 17–22. 29 Sarti P, Jones MG, Antonini G, Malatesta F, Colosimo A, Wilson MT & Brunori M (1985) Kinetics of redox- linked proton pumping activity of native and subunit III-depleted cytochrome c oxidase: a stopped-flow inves- tigation. Proc Natl Acad Sci USA 82, 4876–4880. 30 Hellwig P, Rost B, Kaiser U, Ostermeier C, Michel H &Ma ¨ ntele W (1996) Carboxyl group protonation upon reduction of the Paracoccus denitrificans cytochrome c oxidase: direct evidence by FTIR spectroscopy. FEBS Lett 385, 53–57. 31 Hellwig P, Grzybek S, Behr J, Ludwig B, Michel H & Ma ¨ ntele W (1999) Electrochemical and ultraviolet ⁄ vis- ible ⁄ infrared spectroscopic analysis of heme a and a 3 redox reactions in the cytochrome c oxidase from Para- coccus denitrificans: separation of heme a and a 3 contri- butions and assignment of vibrational modes. Biochemistry 38, 1685–1694. 32 Venyaminov SY & Kalnin NN (1990) IR spectrophoto- metry of peptide compounds in water (H 2 O) solutions. I. Spectral parameters of amino acid residue absorption bands. Biopolymers 30, 1259–1271. 33 Pfitzner U, Hoffmeier K, Harrenga A, Kannt A, Michel H, Bamberg E, Richter O-MH & Ludwig B (2000) Tra- cing the d-pathway in reconstituted site-directed mutants of cytochrome c oxidase from Paracoccus deni- trificans. Biochemistry 39, 6756–6762. 34 Raitio M, Tuulikki J & Saraste M (1987) Isolation and analysis of the genes for cytochrome c oxidase in Para- coccus denitrificans. EMBO J 6, 2825–2833. 35 Steinru ¨ cke P, Gerhus E & Ludwig B (1991) Paracoccus denitrificans mutants deleted in the gene for subunit II of cytochrome c oxidase also lack subunit I. J Biol Chem 266, 7676–7681. 36 Kleymann G, Ostermeier C, Ludwig B, Skerra A & Michel H (1995) Engineered Fv fragments as a tool for the one-step purification of integral multisubunit mem- brane protein complexes. Biotechnology 13, 155–160. 37 Darley-Usmar VM, Capaldi RA, Takamiya S, Millett F, Wilson MT, Malatesta F & Sarti P (1987) Mitochon- dria – a Practical Approach (Darley-Usmar VM, Rick- wood D & Wilson MT, eds), pp. 143–152. IRL Press, Oxford. 38 Moss DA, Nabedryk E, Breton J & Ma ¨ ntele W (1990) Redox-linked conformational changes in proteins detected by a combination of infrared spectroscopy and protein electrochemistry: evaluation of the technique with cytochrome c. Eur J Biochem 187, 565–572. Proton access to Paracoccus cytochrome c oxidase O M. H. Richter et al. 412 FEBS Journal 272 (2005) 404–412 ª 2004 FEBS . Probing the access of protons to the K pathway in the Paracoccus denitrificans cytochrome c oxidase Oliver-M. H. Richter 1 , Katharina L. Du ¨ rr 1 , Aimo Kannt 2 , Bernd Ludwig 1 , Francesca. employing this tech- nique in combination with site-directed mutants and isotopic labeling. Here mutants of E78 II in the Paracoccus cyto- chrome c oxidase are characterized in terms of cata- lytic. distance of 9 A ˚ to the lysine of the K channel (K3 54) [3]. Subsequently this residue gained considerable attention due to a potential importance for access of protons into the K channel of heme-copper