Electrochemical, FT-IR and UV/VIS spectroscopic properties of the caa 3 oxidase from T. thermophilus Petra Hellwig 1 , Tewfik Soulimane 2 * and Werner Ma¨ ntele 1 1 Institut fu ¨ r Biophysik der Johann-Wolfgang-Goethe-Universita ¨ t, Frankfurt/M., Germany; 2 Institut fu ¨ r Biochemie der Rheinisch- Westfa ¨ lischen-Technischen Hochschule, Aachen, Germany The caa 3 -oxidase from Thermus thermophilus has been studied with a combined electrochemical, UV/VIS and Fourier-transform infrared (FT-IR) spectroscopic approach. In this oxidase the electron donor, cytochrome c, is covalently bound to subunit II of the cytochrome c oxidase. Oxidative electrochemical redox titrations in the visible spectral range yielded a midpoint potential of )0.01 ± 0.01 V (vs. Ag/AgCl/3 M KCl, 0.218 V vs. SHE¢) for the heme c. This potential differs for about 50 mV from the midpoint potential of isolated cytochrome c, indicating the possible shifts of the cytochrome c potential when bound to cytochrome c oxidase. For the signals where the hemes a and a 3 contribute, three potentials, ¼ )0.075 V ± 0.01 V, Em 2 ¼ 0.04 V ± 0.01 V and Em 3 ¼ 0.17 V ± 0.02 V (0.133, 0.248 and 0.378 V vs. SHE¢, respectively) could be obtained. Potential titrations after addition of the inhibitor cyanide yielded a midpoint potential of )0.22 V ± 0.01 V for heme a 3 -CN – and of Em 2 ¼ 0.00 V ± 0.02 V and Em 3 ¼ 0.17 V ± 0.02 V for heme a ()0.012 V, 0.208 V and 0.378 V vs. SHE¢, respectively). The three phases of the potential-dependent development of the difference signals can be attributed to the cooperativity between the hemes a, a 3 and the Cu B center, showing typical behavior for cyto- chrome c oxidases. A stronger cooperativity of Cu B is dis- cussed to reflect the modulation of the enzyme to the different key residues involved in proton pumping. We thus studied the FT-IR spectroscopic properties of this enzyme to identify alternative protonatable sites. The vibrational modes of a protonated aspartic or glutamic acid at 1714 cm )1 concomitant with the reduced form of the protein can be identified, a mode which is not present for other cytochrome c oxidases. Furthermore modes at positions characteristic for tyrosine vibrations have been identified. Electrochemically induced FT-IR difference spectra after inhibition of the sample with cyanide allows assigning the formyl signals upon characteristic shifts of the m(C¼O) modes, which reflect the high degree of similarity of heme a 3 to other typical heme copper oxidases. A comparison with previously studied cytochrome c oxidases is presented and on this basis the contributions of the reorganization of the polypeptide backbone, of individual amino acids and of the hemes c, a and a 3 upon electron transfer to/from the redox active centers discussed. Keywords: caa 3 oxidase; cytochrome c oxidase; UV/VIS- spectroscopy; FT-IR-spectroscopy; Thermus thermophilus. Cytochrome c oxidase is the terminal enzyme of the respiratory chain in mitochondria and many prokaryotes. As an integral membrane protein it catalyzes the reduction of dioxygen to water using electrons from cytochrome c. Four redox-active sites are involved in the electron transfer. Electrons from cytochrome c are first transferred to a homobinuclear copper A site (Cu A ) and then subsequently to heme a, and to heme a 3 , which is located close to copper B(Cu B ), forming a heterobinuclear metal center where oxygen is reduced to water. Protons needed for water formationaretakenupfromthecytosolicsideinbacterial membranes or from the matrix side in mitochondria. The proton consumption and the coupled translocation of nH + /e – across the membrane contribute to the proton gradient needed to synthesize ATP. Two pathways have been proposed to serve for consumed and pumped protons on the basis of site-directed mutagen- esis [1,2] and later using the crystal structures [3–5]. These pathways are highly conserved among most studied cyto- chrome oxidases [2,6]. However, cytochrome oxidases have been reported that lack amino acids disputed to be essential in proton translocation. In the case of caa 3 -oxidases from T. thermophilus, for example, as well as from Rhodothermus marinus, the amino acid Glu278 (numbering for Paracoccus denitrificans), which is proposed to pass protons in the D-pathway to the binuclear center, is missing, but proton- pumping activity is observed [3,7–9]. A highly conserved Tyr–Ser couple was suggested to replace Glu278 [9]. In the current understanding, two pathways are necessary for the catalytic activity, but different residues may be involved. In an important step for the understanding of the essentials for cytochrome c oxidase activity and coupled proton pump- ing, the crystal structure of the aberrant ba 3 -oxidase from T. thermophilus was determined [10] and alternative path- ways discussed. Correspondance to P. Hellwig, Institut fu ¨ rBiophysikderJohann- Wolfgang-Goethe-Universita ¨ t, Theodor-Stern-Kai 7 Haus 74, 60590 Frankfurt/M., Germany. Fax: + 49 69 6301 5838, Tel.: + 49 69 6301 4227, E-mail: hellwig@biophysik.uni-frankfurt.de Abbreviations: FT-IR, Fourier-transform infrared; SHE¢, standard hydrogen electrode; TMPD, N,N,N¢,N¢-tetramethyl-p-phenylenedi- amine dihydrochloride *Present address: Paul Scherrer Institut, Structural Biology Group, 5232-CH, Villigen PSI, Switzerland. (Received 13 March 2002, revised 6 August 2002, accepted 14 August 2002) Eur. J. Biochem. 269, 4830–4838 (2002) Ó FEBS 2002 doi:10.1046/j.1432-1033.2002.03182.x Under restricted O 2 supply, the thermophilic Gram negative bacterium T. thermophilus expresses two different cytochrome c oxidases. The heme types incorporated belong to the caa 3 -andba 3 -type cytochrome c oxidases, respectively. The caa 3 -oxidase contains analogous central subunits and catalytic entity to the mitochondrial aa 3 - oxidases, however, including a covalently bound type-c heme [11]. This is currently only found in a few bacteria [9,12]. Recent results showed that the enzyme is made of two fusion proteins. The smaller protein consists of a typical oxidase subunit II sequence, which provides the homonu- clear Cu A binding site and is fused to a cytochrome c domain [11,13]. The larger protein is a fusion product of subunit I, that has the hemes a, a 3 and the Cu B sites, and subunit III [8,12,13]. The heme c center in the caa 3 -oxidase is proposed to serve as the first electron acceptor from a bc 1 complex [14]. We note, however, that no bc 1 complex has yet been described for T. thermophilus. No activity was detected for a reaction with soluble horse heart cyto- chrome c, c 552 from T. thermophilus and yeast iso1 cyto- chrome c, which serve as natural reductands for cytochrome c oxidases, but a reduction can be noted for nonphysiological reducing agents such as N,N,N¢,N¢-tetra- methyl-p-phenylenediamine dihydrochloride (TMPD) [15]. The caa 3 -oxidase may be regarded as an integrated version of the noncovalent redox complex between cytochrome c and cytochrome c oxidase. Previous reports on the caa 3 -oxidase from T. thermophi- lus concluded that this enzyme is a typical member of the heme copper oxidase family [12], with the exception of a different titrimetric behavior of the redox centers in the electron transfer [16] and the lack of some key residues as mentioned above. In this work we study the electrochemical, UV-VIS and FT-IR spectroscopic properties of the caa 3 - oxidase from T. thermophilus in the presence and absence of cyanide, and compare the observed properties to previous reports on members of the heme copper oxidase family such as cytochrome c oxidase from bovine heart and P. denitri- ficans, and the aberrant ba 3 -oxidase from T. thermophilus. MATERIALS AND METHODS Sample preparation The caa 3 -type cytochrome c oxidase from T. thermophilus was prepared as described previously in Gerscher et al. [17]. For electrochemistry the sample was solubilized in n-decyl-b- D -maltopyranoside, 100 m M phosphate buffer (pH 7) containing 100 m M KCl and concentrated to approximately 0.5 m M using Microcon ultrafiltration cells (Millipore). Exchange of H 2 O against D 2 O was performed by repeatedly concentrating the enzyme and rediluting it in a D 2 O phosphate-buffer. H/D exchange was better than 80% as judged from the shift of the amide-II mode at 1550 cm )1 in the FT-IR absorbance spectra (data not shown). For inhibition with cyanide, samples were diluted with 500 lL of 100 m M phosphate buffer containing 20 m M KCN (pH 7), incubated overnight and reconcentrated to 0.5 m M . Electrochemistry The ultra-thin layer spectroelectrochemical cell for the UV/ VIS and IR was used as described previously [18]. 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 pathlength set to 6–8 lm. The gold grid working electrode was chemically modified by a 2-m M cysteamine solution as reported previously [19]. In order to accelerate the redox reaction, 15 different mediators were added as reported by Hellwig et al. [19], with the exception of K 4 [Fe(CN) 6 ], to a total concentration of 40 l M each. At this concentration, and with the pathlength below 10 lm, no spectral contributions from the mediators in the VIS and IR range could be detected in control experiments with samples lacking the protein, except for the PO modes of the phosphate buffer between 1200 cm )1 and 1000 cm )1 .Asa supporting electrolyte, 100 m M KCl was added. Approxi- mately 5–6 lL of the protein solution were sufficient to fill the spectroelectrochemical cell. Potentials quoted with the data refer to the Ag/AgCl/3 M KCl reference electrode, adding + 208 mV for SHE¢ (pH 7) potentials. Midpoint potentials are described for both electrode types. Spectroscopy FT-IR and UV/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 4000–1000 cm )1 range and a dispersive spectrometer for the 400–900 nm range as reported previously [18]. First, the protein was equilibrated with an initial potential at the electrode, and single beam spectra in the VIS and IR range were recorded. A potential step to the final potential was then applied, and single beam spectra of this state were again recorded after equilibration. Difference spectra as presented here were then calculated from the two single beam spectra with the initial single beam spectrum taken as a reference. No smoothing or deconvolution procedures were applied. The equilibration process for each potential applied was followed by monitoring the electrode current and by successively recording spectra in the visible range until no further changes were observed. The equilibration generally took less than 8 min under the conditions reported (protein concentration, electrode modification, mediators) for the full potential step from )0.5V to 0.5V and to selected potentials. Typically, 128 interferograms at 4 cm )1 resolution were coadded for each single beam IR spectrum and Fourier-transformed using triangular apodization. Differences in sample concentration and pathlength were taken into account by normalizing the FT-IR difference spectra on the difference signal of the sample in the UV/VIS at 602 nm. Redox titrations The redox-dependent absorbance changes of the caa 3 - oxidase from T. thermophilus were studied performing electrochemical redox titrations in the UV/VIS spectral range. The redox titrations were performed by stepwise setting the potential and recording the spectrum after sufficient equilibration. Typically data were recorded at steps of 30–50 mV. All measurements were performed at 5 °C. The midpoint potentials E m and the number n of transferred electrons were obtained by adjusting a calcula- ted Nernst curve to the measured absorbance change at a Ó FEBS 2002 Characterization of T. thermophilus caa 3 oxidase (Eur. J. Biochem. 269) 4831 single wavelength by an interactive fit. All parameters have to be adjusted manually until the theoretical Nernst curve and the measured data match well (fit by eye). RESULTS AND DISCUSSION UV/VIS difference spectra Figure 1A shows the oxidized-minus-reduced UV/VIS difference spectra of the caa 3 -oxidase from T. thermophilus obtained for a potential step from )0.5 V to 0.5 V (vs. Ag/ AgCl/3 M KCl). In the oxidized-minus-reduced spectra the positive signals correlate with the oxidized and the negative signals with the reduced form of the enzyme. For the reduced form, the Soret band can be observed at 415 and 442 nm, and for the oxidized form at 403 and 422 nm. The b–band can be seen at 517 nm and the a–band at 547 nm and 603 nm. The difference signals that can be observed between 400 and 700 nm include the contributions of the hemes c, a and a 3 . The difference signals observed at 403, 415, 517 and 547 nm are characteristic for heme c. In electrochemically induced difference spectra of horse heart cytochrome c the Soret band was reported to absorb at 418 nm, the b-band at 520 nm and the a–band at 550 nm [18]. The deviations of approximately 3 nm between the signals of horse heart cytochrome c and the heme c in the caa 3 -oxidase from T. thermophilus can be attributed to the different environ- ment of the heme centers. The difference signals observed at 442 nm and 603 nm can be assigned to the contributions of the hemes a and a 3 , with the position of the a-band showing a downshift in relation to bovine heart oxidase. In Fig. 1B the oxidized-minus-reduced UV/VIS differ- ence spectra of the caa 3 -oxidase poisoned with cyanide obtained for a potential step from )0.5 V to 0.5 V (solid line) and from )0.5–0.05 V (dotted line) can be seen. The a-band shifts to 599 nm upon binding of cyanide to heme a 3 . This shift is reflected clearly in the critical potential step from )0.5 to )0.05 V, where mainly heme a 3 and heme c contribute. Addition of cyanide was used in the electro- chemically induced FT-IR difference spectra to separate contributions of heme a 3 . UV/VIS redox titrations In Fig. 2A the potential dependent development of the a–band from heme c at 548 nm in an oxidative titration can be seen (filled circles). The theoretical Nernst fit described in Materials and methods yields a midpoint potential of E m ¼ )0.01 ± 0.01 V (vs. Ag/AgCl, and 0.218 V vs. SHE¢) for heme c. This value was also reported by Yoshida and Fee [16]. The midpoint potential of soluble horse heart cytochrome c is 0.048 V (0.256 V vs. SHE¢, as obtained with the same method as described here and in [18]); other cytochrome c types show a close midpoint potential. The midpoint potential of the cytochrome c in the cyto- chrome c–cytochrome c oxidase complex is unknown and may be the origin for this downshift. Alternatively, the electron transfer directly from the bc 1 complex, as suggested for a possible mechanism, could require a lower potential. Figure 2B (filled circles) shows the potential-dependent development of the difference signal at 443 nm from )0.4 V to + 0.6 V. Three phases can be clearly discriminated. The theoretical Nernst fit yields midpoint potentials Em 1 ¼ )0.075 ± 0.01 V, Em 2 ¼ 0.04 ± 0.01 V and Em 3 ¼ 0.17 ± 0.02 V for an n value of 0.9–1 (these values correspond to 0.133, 0.248 and 0.378 V vs. SHE¢, respect- ively). As reported previously, cytochrome c oxidase from bovine heart shows a complex titration curve reflecting the cooperative interactions between the hemes a and a 3 ,and the Cu B center [20–22]. For cytochrome c oxidase from bovine heart E m values near 200, 260 and 340 mV have been reported [23–25], and analyzed in detail by several groups [20–22]. For the caa 3 -oxidase we observe a noteworthy pronounced phase at 40 mV (248 mV vs. SHE¢) but essentially a similar titrimetric behavior. In a previously reported potential titration of the caa 3 -oxidase from T. thermophilus, Yoshida and Fee [16] describe a compar- able potential and n-value for the first step, but report a second step with an n-value of two at approximately 160 mV for Cu B and heme a 3 .Onthisbasis,Cu B and heme a 3 have been suggested to act as a two-electron acceptor [12] in contrast to bovine heart oxidase, where subsequent one-electron transfer is reported. The three phasic curve, with a step of n ¼ 1 for each step as found here, shows a significantly more comparable titrimetric behavior in comparison to other typical oxidases, but in contrast to the work by Yoshida and Fee [16]. The small difference to other oxidases found here, reflected in the stronger second step at 40 mV, may be attributed to the Fig. 1. Oxidized-minus-reduced UV/VIS difference spectra of the caa 3 - oxidase from T. thermophilus. Results obtained for a potential step from )0.5 V to 0.5 V (vs. Ag/AgCl/3 M KCl) in the absence (A) or the presence (B, solid line) of cyanide, and for a potential step from )0.5 to 0.05 V in the presence of cyanide (B, dotted line). 4832 P. Hellwig et al.(Eur. J. Biochem. 269) Ó FEBS 2002 covalently attached heme c or to a generally changed cooperativity of the other redox centers. In order to discriminate the contributions of the cofac- tors, inhibitors uncoupling or changing the cooperativity can be used. Addition of cyanide strongly shifts the heme a 3 potential and thus uncouples or changes cooperativity in the binuclear center [20,22,28]. The shifts of the titration curve upon addition of cyanide can be seen in Fig. 2B (open circles). The theoretical Nernst fit yielded midpoint poten- tials Em 1 ¼ )0.22 ± 0.01 V, Em 2 ¼ 0.00 ± 0.01 V and Em 3 ¼ 0.17 ± 0.02 V (the values correspond to )0.012, 0.208 and 0.378 V vs. SHE¢, respectively). The potential at )220 mV can be attributed to the heme a 3 –CN – signal, this shift reflecting the characteristic behavior of cytochrome c oxidases. heme a is expected to contribute with two steps, reflecting the remaining interactions with Cu B .Asseenin Fig. 2B (open circles), a further interaction is observed, presenting additional evidence for a different cooperativity of the redox centers. Whereas in the Soret Band heme a and heme a 3 contribute almost equally, the heme a contribution domin- ates the a-band. Figure 2C shows a comparison of the potential-dependent development of the modes at 599 nm (triangles) and 442 nm (open circles) in the presence of cyanide. As seen for the curve that represents the titration curve at 602 nm (triangles) a smaller ratio is present for the potential at )220 mV than for the titration curve measured at 442 nm in the same conditions, supporting the assign- ment to heme a 3 . Heme c, however, shows a relatively small difference in midpoint potential of 15 mV in the presence of cyanide (Fig. 2A, empty circles) and thus does not indicate a noteworthy cooperativity between heme a 3 and the heme c centers. Heme c can be ruled out as origin for the distinct second phase at 40 mV in the titration curve for the hemes a and a 3 . It may be suggested that the different cooperativity, as well as the lower heme a 3 potential, is necessary to compensate the differences caused by the presence of different key residues in the D-pathway, since the potentials are assumed to be crucial for the coupling of electron and proton transfer. Interestingly, for the caa 3 -oxidase from R. marinus which also lacks the above-mentioned Glu278 side chain, downshifted potentials for the hemes a and a 3 have been described, although the cooperativity is not discussed [9]. In the case of the aberrant ba 3 -cytochrome c oxidase from T. thermophilus, a completely different titri- metric behavior was observed [26], also indicating that the midpoint potentials and cooperativity are adapted to the varying proton path residues. To emphasize this suggestion further, future comparative studies on the varying oxidases could be performed. FT-IR difference spectra Figure 3 shows the oxidized-minus-reduced FT-IR differ- ence spectra of the caa 3 -oxidase from T. thermophilus for a potential step from )0.5 V to 0.5 V equilibrated in H 2 O(A) and D 2 O buffer (B). Numerous distinct sharp bands appear throughout the spectrum, with half-widths typically below 5–10 cm )1 . The noise level in these difference spectra can be estimated at approximately 25–50 · 10 )6 absorbance units at frequencies above 1750 cm )1 , where no signals appear. Only in regions of strong absorbance of the sample, such as Fig. 2. Potential dependent development of the hemes in the caa 3 -oxid- ase from T. thermophilus . Heme c was monitored at 548 nm in the absence (filled circles) and presence (open circles) of cyanide (A) and a midpoint potential of )0.01 V ± 0.01 V (vs. Ag/AgCl/3 M KCl or 0.218 V vs. SHE¢) was obtained by a theoretical Nernst fit (solid line). The hemes a and a 3 were monitored at 442 nm in the absence (filled circles) and presence (open circles) of cyanide. Midpoint poten- tials of Em 1 ¼ )0.075V±0.01V, Em 2 ¼ 0.04 V ± 0.01 V and Em 3 ¼ 0.17 V ± 0.02 V were determined (these values correspond to 0.133 V, 0.248 V and 0.378 V vs. SHE¢, respectively). After addition of the inhibitor cyanide (open circles) a midpoint potential of )0.22 V ± 0.01 V for heme a 3 -CN – and of Em 2 ¼ 0.00 V ± 0.02 V and Em 3 ¼ 0.17 V ± 0.02 V for heme a canbeseen(thevalues correspond to )0.012 V, 0.208 V and 0.378 V vs. SHE¢, respectively). (B) Comparison of the potential dependent development of the modes at 599 nm (triangles) and 442 nm (open circles) in the presence of cyanide. The theoretical Nernst fit is shown as a solid line (C). Ó FEBS 2002 Characterization of T. thermophilus caa 3 oxidase (Eur. J. Biochem. 269) 4833 around 1650 cm )1 (water OH-bending mode and amide-I C¼O mode), was the noise level slightly higher, though never exceeding 10 )4 absorbance units. The entirety of difference signals represent the total molecular changes concomitant with the redox reactions. In the electrochemically induced FT-IR difference spectra, contributions from the porphyrin ring, the heme propio- nates and the vinyl substituent can be expected, originating from heme c, with contributions from the formyl groups and from the geranyl side chain expected from heme a and a 3 . In addition to the signals of the hemes, the reorgan- ization of the polypeptide backbone and amino acid side chains occurring upon electron transfer of the five redox active centers heme c, a, a 3 ,Cu A and Cu B , and coupled processes such as proton transfer can be expected to manifest themselves in the spectra. In the following paragraph the difference spectra will be described and discussed. Tentative assignments will be presented on the basis of the comparison to IR and Raman spectra of heme model compounds, other oxidases, spectra of isolated amino acids as model compounds and information on contributions from the secondary structure from infrared absorbance spectra and the deconvolution of the amide-I region. A particular problem of the assignment in the difference spectra is the superposition of signals from different constituents of the oxidase, which can lead to the possibility of multi component bands and may present ambiguities in the assignment. A spectral region particularly susceptible for overlapping bands is the amide-I range. Although in this range (approx. 1690–1610 cm )1 ) typical contributions from secondary structure elements are expected, and signals may point to the alterations of local protein conformation in the course of the redox reaction, we keep in mind that the heme formyl mode also contributes here, as well as specific modes from amino acid side chains. For a clearer discrimination of these overlapped bands, we used deuteration of the sample and FT-IR spectra studies in the presence of the inhibitor cyanide support our tentative assignments. Tentative assignments of difference signals to polypeptide backbone modes Amide-I signals are predominantly caused by the C¼O stretching vibration of the polypeptide backbone. For different secondary structure elements, characteristic absorptions can be distinguished. In the electrochemically induced FT-IR difference spectra, contributions from the reorganization of the polypeptide backbone upon electron transfer to and from the cofactors can be expected, and a partial attribution of the signals observed in the amide-I region (1690–1610 cm )1 ) to amide-I modes is conceivable. The different secondary structure elements show a different sensitivity to H/D exchange. In Fig. 3A strong positive signals can be observed at 1694 cm )1 , 1684 cm )1 , 1674 cm )1 and 1646 cm )1 , and prominent negative differ- ence modes are present at 1660 cm )1 , 1626 cm )1 and 1614 cm )1 . After H/D exchange (Fig. 3B) the increase of the signal at 1696 cm )1 and 1626 cm )1 can be observed. A clear shift from 1634 cm )1 to 1658 cm )1 and to 1650 cm )1 is visible. The modes involved in the signals at 1660 cm )1 contribute in the range characteristic for the absorbance from a–helical secondary structure elements. However, absorbance changes induced by the reorganization of a-helical secondary structure elements are expected to show very small shifts after H/D exchange at most (2–10 cm )1 ) and an assignment is unlikely. Unordered secondary structure elements show a higher sensitivity to H/D exchange and also contribute in this spectral range. An involvement of reorganizations of b–sheet secondary struc- ture elements is possible for the difference in the signals at 1696–1674 cm )1 , and at 1646–1620 cm )1 . However, con- clusive assignments are difficult in this spectral range where difference bands strongly overlap. In the amide-II region (1575 cm )1 )1480 cm )1 ), strong negative signals at 1546 cm )1 and 1516 cm )1 as well as positive signals at 1562 cm )1 and 1498 cm )1 can be observed. An assignment of the signals in the amide-II region in the electrochemically induced FT-IR difference spectrum of the caa 3 -oxidase from T. thermophilus to amide-II modes, however, appears less probable since little or no shift for H/D exchange is observed. Assignment of heme vibrational modes Formyl substituent. The C¼O bond of the formyl group at the porphyrin ring of hemes a and a 3 can be expected to contribute between 1680 cm )1 and 1606 cm )1 , depending on hydrogen bonding with neighboring amino acids. The formyl substituent of heme a is predicted to form a Fig. 3. Oxidized-minus-reduced FT-IR difference spectra of the caa 3 - oxidase from T. thermophilus. Results obtained for a potential step from )0.5 V to 0.5 V (vs. Ag/AgCl/3 M KCl) equilibrated in H 2 O(A) and D 2 O buffer (B). 4834 P. Hellwig et al.(Eur. J. Biochem. 269) Ó FEBS 2002 hydrogen bond with an arginine side chain, while the same substituent for heme a 3 appears to be free from H-bonding to nearby amino acid residues. Different frequencies for the m(C¼O) stretching mode of the formyl group can thus be expected. In resonance Raman spectra of caa 3 -oxidase from T. thermophilus a signal at 1611 cm )1 could be assigned to the m(C¼O) CHO from reduced heme a and at 1649 cm )1 to the oxidized form [17]. Comparable signals can be observed here at 1650 cm )1 for the oxidized form and at 1608 cm )1 for the reduced form. The resonance Raman spectroscopic characterization of the m(C¼O) CHO vibrational modes for heme a 3 showed the presence of a mode for the reduced form at 1664 cm )1 and for the oxidized form at 1673 cm )1 [17]. In the electrochemically induced FT-IR difference spectra in Fig. 3A corresponding bands can be seen at 1678 cm )1 (oxidized form) and 1660 cm )1 (reduced form). These modes have been previously attributed to the formyl side chain from cytochrome c oxidase from bovine heart [27–29] reportedtobesensitivetoCN – binding in a characteristic way [27]. In Fig. 4A the spectra in the presence of cyanide clearly reflect a shift of the mode at 1678–1668 cm )1 and of the mode at 1660–1652 cm )1 , supporting the assignment to the heme a 3 formyl mode. In a direct comparison of these vibrational modes to those observed for the cytochrome c oxidase from P. denitrificans, an analog environment of the protein site of the heme a 3 formyl group in the absence and presence of cyanide can be concluded. Porphyrin ring vibrations. Porphyrin ring vibrations of the heme centers, for example the CaCm vibration (m 37 )or the CbCb vibration (m 38 ) can be expected between 1620 cm )1 and 1500 cm )1 and are involved in the electrochemically induced FT-IR difference spectra shown in Figs 3 and 4. On the basis of recent resonance Raman work on the caa 3 -oxidase from T. thermophilus [17] and a direct comparison to resonance Raman and FT-IR investigations on other oxidases [17,23,30] tentative as- signments of porphyrin ring vibrations have been made and summarized in Table 1. As described previously by Gerscher et al. [17], the spectroscopic properties of the hemes a and a 3 sites are comparable to other typical aa 3 oxidases. Additionally contributions of the heme c center can be expected. It is clear that, in addition to the modes assigned here, further C–C or C–N vibrations of the porphyrin ring (m 4 ,m 39 ) will contribute to the electrochemically induced FT-IR difference spectra. However, we refrain from dis- cussing and assigning these modes on the basis of the data presented here, in spite of the fact that bands in the difference spectra are observed in the region where the modes were attributed. The vibrational modes of bound cyanide Electrochemically induced FT-IR difference spectra of cyanide bound to heme a 3 were characterized to specify possible variations of the binuclear center in direct compari- son to other oxidases, as for example the cytochrome c oxidase from P. denitrificans. In the spectral range from 2200–2000 cm )1 contributions from cyanide ligand bound to heme a 3 can be expected. In the inset in Fig. 4 a strong positive mode can be seen at 2148 cm )1 and a negative signal at 2040 cm )1 for a potential step from )0.5 to 0.5 V (unbroken line) and for )0.5 to 0.05 V (dotted line). A small mode at 2092 cm )1 can be seen in the reduced state, indicating the presence of free cyanide upon reduction. The band at 2148 cm )1 may be assigned to the C–N stretching of the Fe 3+ –C¼N–Cu B 2+ entity (also a Fe 3+ –C¼N–Cu B 2+ – C¼N structure was discussed) based on the spectral shifts observed for isotopically labeled cyanide complexes [31–33]. Correspondingly, the band at around 2040 cm )1 of the reduced form could be attributed tentatively to the C–N stretching of a nonbridging cyanide ligand of heme a 3 .In the spectra observed for the critical potential step from )0.5 to 0.05 V, where the reorganization upon oxidation of the inhibited heme a 3 center is induced, the cyanide modes are completely developed. To allow the above-mentioned bridged structure to be present, Cu B must be oxidized at this potential in the presence of cyanide, since the contri- bution of the unbridged Fea 3 3+ –CN – structure was repor- ted to be observable at 2132 cm )1 . The position of the cynide vibrational modes are essentially identical to the ones observed for bovine heart oxidase [31] and from P. denitrificans (Hellwig et al. unpublished results) reflecting a close environment and ligand binding properties of the binuclear heme a 3 –Cu B center. Identification of protonable sites Aspartic and Glutamic acid side chains. The m(C¼O) mode of protonated aspartic and glutamic side chains absorb typically above 1710 cm )1 , the exact absorption depending on the hydrogen bonding. A negative mode is present at 1714 cm )1 , which shifts to 1716 cm )1 upon H/D exchange. This indicates the presence of a protonated Fig. 4. Oxidized-minus-reduced FT-IR difference spectra of the caa 3 - oxidase from T. thermophilus. Results obtained for a potential step from )0.5 V to 0.5 V (vs. Ag/AgCl/3 M KCl) in the absence (dotted line) and presence of cyanide (solid line). The inset shows an enlarged view of the spectral region characteristic for the CN modes from 2200 to 2000 cm )1 . Ó FEBS 2002 Characterization of T. thermophilus caa 3 oxidase (Eur. J. Biochem. 269) 4835 aspartic or glutamic acid side chain in the reduced state of the enzyme, which is coordinated with a strong hydrogen bond. A small very broad positive mode is found at 1744 cm )1 , indicating the possible contribution of a surface group in the oxidized form of the protein. This mode shifts to 1740 cm )1 upon H/D exchange. The signal may originate from heme c reduction, but also reflect a distinct protonable site in subunit I, involved in a different proton pathway. For the cytochrome c oxidase from P. denitrificans difference modes at 1746 cm )1 and 1734 cm )1 have been observed and attributed to Glu278 [19,34], and to the equivalent residues in the cytochrome bo 3 quinol oxidase from E. coli [35,36], a residue which lacks the caa 3 -oxidase as mentioned above. Correspondingly no analogous con- tribution can be seen here. Tyrosines. Pereira et al. [9] recently suggested a Tyr–Ser motif, conserved in several of the cytochrome c oxidases which lack the above-mentioned Glu278 residue, to be involved in proton pumping. For tyrosine side chains, the m 19 (CC) ring mode for the protonated form of tyrosines is proposed to absorb with an strong signal at 1518 cm )1 and for the deprotonated form at 1496–1486 cm )1 [37,38]. Clearly a difference mode in the spectral region character- istic for the protonated form can be seen concomitant with the reduced state at 1515 cm )1 and the mode typical for the deprotonated form at 1498 cm )1 , indicating the protonation of a tyrosine residue with the reduction of the protein. Also the m 7¢a (CO) and d(COH) of tyrosine side chains expected at approximately 1265 cm )1 and 1245 cm )1 respective to the protonation state, can be seen. We note that these assignments are highly tentative until this data can be supported by combining the technique with site-directed mutants or labeled compounds. Heme propionates. The heme propionates at the hemes a and a 3 are discussed to be involved in proton translocation Table 1. Summary of tentative assignments of the vibrational modes involved in the electrochemically induced FT-IR difference spectra of the caa 3 - oxidase from T. thermophilus. caa 3 FT-IR caa 3 RR [17] Redox state Tentative assignments Comparable modes for aa 3 P. denitrificans [30] 1744 ox m(C¼O) Glu278 for P. denitrificans 1746 – red m(C¼O) Glu278 for P. denitrificans 1734 1714 – red m(C¼O) Asp/Glu – 1708 ox m(C¼O) Asp/Glu 1708 1694 ox 1692 red amide-I (b-sheet) 1694 1684 ox amide-I (b-sheet) 1688 1682 red amide-I (b-sheet, loops) 1684 1678 1674 ox m(C¼O) CHO heme a 3 m(C¼O) heme propionates m(C¼O) Asn/Gln m(CN 3 H 5 ) as Arg 1676 1660 1665 red m(C¼O) CHO heme a 3 amide-I (a-helical) m(CN 3 H 5 ) as Arg 1662 1650 1650 ox amide-I (a-helical) m(C¼O) CHO heme a 1656/1644 1646/1636 ox amide-I (b-sheet) 1656/1644 1626 red d(NH 2 ) Asn/Gln m(CN 3 H 5 ) s Arg amide-I (b-sheet) 1632 1608 red m 37 heme c 1602 1604 ox m 37 heme a (m 8a / 8b (CC) Tyr-OH) 1608 1610 red m(C¼O) CHO heme a 1606 1597 ox m 37 heme c 1580 1585 ox m 37 heme a 3 1588 1562 1567/1558 ox m 38x heme a/a 3 m(COO – ) as heme propionate m(COO – ) as Asp/Glu m(CC) ring Tyr-O – 1564 1546 1545 red m 38y heme a 1548 1530 1532 red m 38y heme a 3 m(COO – ) as heme a 3 propionate 1528 1515 – red m 19 (CC) Tyr-OH – 1498 – ox m 19 (CC) Tyr-O – 1265 – ox m 7¢a (CO) Tyr-O – 1245 – red m 7¢a (CO) and d(COH) Tyr-OH 4836 P. Hellwig et al.(Eur. J. Biochem. 269) Ó FEBS 2002 during catalytic cycle [39]. The contributions of protonated and ionized carboxylic groups of the heme propionates for the cytochrome c oxidase from P. denitrificans were assigned by specific 13 C-labelling of the carboxylic groups of the four heme propionates and site-directed mutagenesis in the vicinity of its site [40,41]. A signal at 1676 cm )1 was attributed to contributions of protonated carboxylic groups. Difference bands at 1570 cm )1 and 1538 cm )1 were assigned to the m(COO – ) as vibration and at 1380 cm )1 to the m(COO – ) s vibration of deprotonated heme propionates [40,41]. Signals at comparable positions can be seen in the spectra shown for the caa 3 -oxidase. In addition the contributions of the heme propionates of the heme c can be expected in a comparable spectral region. CONCLUSIONS The superfamily of heme copper oxidases includes a number of enzymes, which show deviations to the centrally discussed oxidases. The study of these aberrant systems is important to understand the principles of these enzymes. The cytochrome c oxidase from T. thermophilus studied in this work lacks a key residue in the so-called D-pathway, although it does show proton pumping activity. A Tyr–Ser motif was previously suggested to replace the absent acidic group in several oxidases [9]. In this work we could observe modes at characteristic positions for the protona- tion of a tyrosine side chain concomitant with the reduction of the enzyme. A further alternative protonable sitecouldbeseenat1714cm )1 . This mode is observable in the spectral range characteristic for protonated aspartic or glutamic acid side chains and reflects its protonation with the reduction of the protein. We note that these assign- ments are tentative and can be supported by the combi- nation with site-directed mutagenesis or labeling experiments. Interestingly potential titrations of the enzyme show a slightly different redox-dependent behavior. It may be suggested that the stronger cooperativity displays the modulation of the enzyme to the different residues involved. This is in line with the observation reported previously for the caa 3 -oxidase from R. marinus and ba 3 -oxidase from T. thermophilus [9,26]. An influence of the attached heme c center is less likely on the basis of titrations in the presence of cyanide. The electrochemically induced FT-IR difference spectra also include the contributions of the heme centers c, a and a 3 . Together with the spectra in the presence of cyanide and in direct comparison to previous resonance Raman data it can be concluded that the hemes a and a 3 have a similar structural environment comparised with bovine heart and P. denitrificans oxidases [17]. In summary, the caa 3 -cytochrome c oxidase shows the characteristic complex redox behavior and shows several structural properties of a typical cytochrome c oxidase. The presence of the two proton pathways is discussed as one essential of the cytochrome c oxidase. The so-called D-pathway seems to involve different residues here, most likely a tyrosine and an aspartic or glutamic acid. It may also be suggested that the complex redox behavior is crucial for the cytochrome c oxidase mechanism, with some variations, as observed here. REFERENCES 1. Thomas, J.W., Puustinen, A., Alben, J.O., Gennis, R.B. & Wikstro ¨ m, M. (1993) Substitution of asparagine for aspartate- 135 in subunit I of the cytochrome bo ubiquinol oxidase of Escherichia coli eliminates-pumping activity. Biochemistry 32, 10923–10928. 2.Garcia-Horsman,J.A.,Barquera,B.,Rumbley,J.,Ma,J.& Gennis, R.B. (1994) The superfamily of heme-copper respiratory oxidases. J. Bacteriol. 176, 5587–5600. 3. Tsukihara, T., Aoyama, H., Yamashita, E., Tomizaki, T., Yamaguchi, H., Shinzawa-Itoh, K., Nakashima, R., Yaono, R. & Yoshikawa, S. (1995) Structures of metal sites of oxidized bovine heart cytochrome c oxidase at 2.8 A ˚ . Science 269, 1069– 1074. 4. Iwata, S., Ostermeier, C., Ludwig, B. & Michel, H. (1995) Struc- ture at 2.8 A ˚ resolution of cytochrome c oxidase from Paracoccus denitrificans. Nature 376, 660–669. 5. Ostermeier, C., Harrenga, A., Ermler, U. & Michel, H. (1997) Structure at 2.7 A ˚ resolution of the Paracoccus denitrificans two- subunit cytochrome c oxidase complexed with an antibody FV fragment. Proc. Natl Acad. Sci. 94, 10547–10553. 6. Pereira, M.M., Santana, M. & Teixeira, M. (2001) A novel scenario for the evolution of haem-copper oxygen reductases. Biochim. Biophys. Acta. 1505, 185–208. 7. Yoshida, T. & Fee, J.A. (1984) Studies on cytochrome c oxidase activity of the cytochrome c 1 aa 3 complex from Thermus thermo- philus. J. Biol. Chem. 259, 1031–1036. 8.Mather,M.W.,Springer,P.,Hensel,S.,Buse,G.&Fee,J.A. (1993) Cytochrome oxidase genes from Thermus thermophilus. Nucleotide sequence of the fused gene and analysis of the deduced primary structures for subunits I and III of cytochrome caa 3 . J. Biol. Chem. 268, 5395–5408. 9. Pereira, M.M., Verkhovskaya, M.L., Teixeira, M. & Verkhovsky, M.I. (2000) The caa 3 terminal oxidase of Rhodothermus marinus lacking the key glutamate of the D -channel is a proton pump. Biochemistry 39, 6336–6340. 10. Soulimane, T., Buse, G., Bourenkov, G.P., Bartunik, H.D., Huber, R. & Than, M.E. (2000) Structure and mechanism of the aberrant ba 3 cytochrome c oxidase from Thermus thermophilus. EMBO J. 19, 1766–1776. 11. Buse, G., Hensel, S. & Fee, J.A. (1989) Cytochrome oxidase genes from Thermus thermophilus. Nucleotide sequence of the fused gene and analysis of the deduced primarystructures for subunits I and III of cytochrome caa 3 . Eur J. Biochem. 181, 261–268. 12. Fee, J.A., Yoshida, T., Surerus, K.K. & Mather, M.W. (1993) cytochrome caa 3 from the thermophilic bacterium Thermus ther- mophilus: a member of the heme-copper oxidase superfamily. J. Bioenergetics Biomembranes 25, 103–114. 13. Mather, M.W., Springer, P. & Fee, J.A. (1991) Cytochrome oxi- dase genes from Thermus thermophilus. Nucleotide sequence and analysis of the deduced primary structure of subunit IIc of cyto- chrome caa 3 . J. Biol. Chem. 266, 5025–5035. 14. Trumpower, B.L. (1991) The three-subunit cytochrome bc 1 com- plex of Paracoccus denitrificans. Its physiological function, struc- ture, and mechanism of electron transfer and energy transduction. J. Bioenerg. Biomembr. 23, 241–255. 15. Soulimane,T.,vonWalter,M.,Hof,P.,Than,M.E.,Huber,R.& Buse, G. (1997) Cytochrome-c552 from Thermus thermophilus:a functional and crystallographic investigation. Biochem. Biophys. Res. Comm. 237, 572–576. 16. Yoshida, T. & Fee, J.A. (1985) Potentiometric study of cyto- chrome c 1 aa 3 from Thermus thermophilus. J. Inorg. Biochim. 23, 279–288. 17. Gerscher, S., Hildebrandt, P., Soulimane, T. & Buse, G. (1998) Resonance Raman spectroscopic study of the caa 3 oxidase from Thermus thermophilus. Biospectroscopy 4, 365–377. Ó FEBS 2002 Characterization of T. thermophilus caa 3 oxidase (Eur. J. Biochem. 269) 4837 18. Moss, D.A., Nabedryk, E., Breton, J. & Ma ¨ ntele, W. (1990) Redox-linked conformational changes in proteins detected by a combination of infrared spectroscopy and protein electro- chemistry. Evaluation of the technique with cytochrome c. Eur. J. Biochem. 187, 565–572. 19. Hellwig, P., Behr, J., Ostermeier, C., Richter, O.M., Pfitzner, U., Odenwald, A., Ludwig, B., Michel, H. & Ma ¨ ntele, W. (1998) Involvement of glutamic acid 278 in the redox reaction of the cytochrome c oxidase from Paracoccus denitrificans investigated by FTIR spectroscopy. Biochemistry 37, 7390–7399. 20. Wikstro ¨ m, M.K., Krab, K. & Saraste, M. (1981) Cytochrome oxidase, a synthesis. Academic Press, New York. 21. Babcock, G.T., Vickery, L.E. & Palmer, G. (1978) The electronic state of heme in cytochrome oxidase II. Oxidation–reduction potential interactions and heme iron spin state behavior observed in reductive titrations. J. Biol. Chem. 7, 2400–2411. 22. Hendler, R.W. & Westerhoff, H.V. (1992) Redox interactions in cytochrome c oxidase: from the ÔneoclassicalÕ toward ÔmodernÕ models. Biophy. J. 63, 1586–1604. 23. Reddy, K.V., Hendler, R.W. & Bunow, B. (1986) Complete analysis of the cytochrome components of beef heart mitochon- dria in terms of spectra and redox properties. Cytochromes aa 3 . Biophys. J. 49, 705–715. 24. Hendler, R.W., Reddy, K.V., Shrager, R.I. & Caughey, W.S. (1986) Analysis of the spectra and redox properties of pure cyto- chromes aa 3 . Biophys. J. 49, 717–729. 25. Sidhu, G.S. & Hendler, R.W. (1990) Characterization of two low Em forms of cytochrome a 3 and their carbon monoxide complexes in mammalian cytochrome c oxidase. Biophys. J. 57, 1125–1140. 26. Hellwig, P., Soulimane, T., Buse, G. & Ma ¨ ntele, W. (1999) Elec- trochemical, FTIR, and UV/VIS spectroscopic properties of the ba 3 oxidase from Thermus thermophilus. Biochemistry 38, 9648– 9658. 27. Babcock, G.T. (1988) Raman scattering by cytochrome oxidase and by heme a model compounds. In Biological Applications of Resonance Raman Spectroscopy (T.Spiro,ed.),pp.294.Wileyand Sons, New York. 28. Harmon, P.A., Hendler, R.W. & Levin, I.W. (1994) Resonance Raman and optical spectroscopic monitoring of heme a redox states in cytochrome c oxidase during potentiometric titrations. Biochemistry 33, 699–707. 29. Ching, Y.C., Argade, P.V. & Rousseau, D.L. (1985) Resonance raman spectra of CN-bound cytochrome oxidase: spectral isola- tion of cytochromes a2+, a3(2+), and a3(2+)(CN-). Biochem- istry. 27, 4938–4946. 30. Hellwig, P., Grzybek, S., Behr, J., Ludwig, B., Michel, H. & Ma ¨ ntele, W. (1999) Electrochemical and ultraviolet/visible/infra- red spectroscopic analysis of heme a and a 3 redox reactions in the cytochrome c from Paracoccus denitrificans:separationofhemea and a 3 contributions and assignment of vibrational modes. Biochemistry 38, 1685–1694. 31. Yoshikawa, S. & Caughey, W.S. (1990) Infrared evidence of cyanide binding to iron and copper sites in bovine heart cyto- chrome c oxidase. Implications regarding oxygen reduction. J. Biol. Chem. 265, 7945–7958. 32. Yoshikawa, S., Mochizuki, M., Zhao, X.J. & Caughey, W.S. (1995) Effects of overall oxidation state on infrared spectra of heme a 3 cyanideinbovineheartcytochromec oxidase. Evidence of novel mechanistic roles for CuB. J. Biol. Chem. 270, 4270–4279. 33. Tsubaki, M., Matsushita, K., Adachi, O., Hirota, S., Kitagawa, T. & Hori, H. (1997) Resonance Raman, infrared, and EPR investigation on the binuclear site structure of the heme-copper ubiquinol oxidases from Acetobacter aceti: effect of the heme peripheral formyl group substitution. Biochemistry 36, 13034– 13042. 34. 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 evi- dence by FTIR spectroscopy. FEBS-Lett. 385, 53–57. 35. Lu ¨ bben, M. & Gerwert, K. (1996) Redox FTIR difference spec- troscopy using caged electrons reveals contributions of carboxyl groups to the catalytic mechanism of haem-copper oxidases. FEBS-Lett. 397, 303–307. 36. Yamazaki, Y., Kandori, H. & Mogi, T. (1999) Effects of subunit I mutations on redox-linked conformational changes of the Escherichia coli bo-type ubiquinol oxidase revealed by Fourier- transform infrared spectroscopy. J. Biochem. (Tokyo) 126, 194–199. 37. Venyaminov, S.Y. & Kalnin, N.N. (1990) Quantitative IR spec- trophotometry of peptide compounds in water (H 2 O) solutions. I. Spectral parameters of amino acid residue absorption bands. Biopolymers 30, 1243–1257. 38. Hienerwadel, R., Boussac, A., Breton, J., Diner, B. & Berthomieu, C. (1997) Fourier transform infrared difference spectroscopy of photosystem II tyrosine D using site-directed mutagenesis and specific isotope labeling. Biochemistry 36, 14712–14723. 39. Michel, H., Behr. J., Harrenga, A. & Kannt, A. (1998) cytochrome c oxidase: structure and spectroscopy. Ann. Rev. Biophys. Biomol. Struct. 27, 329–356. 40. Behr, J., Hellwig, P., Ma ¨ ntele, W. & Michel, H. (1998) Redox dependent changes at the heme propionates in cytochrome c oxi- dase from Paracoccus denitrificans: direct evidence from FTIR difference spectroscopy in combination with heme propionate 13 C labeling. Biochemistry 37, 7400–7406. 41. Behr, J., Michel, H., Ma ¨ ntele, W. & Hellwig, P. (2000) Functional properties of the heme propionates in cytochrome c oxidase from Paracoccus denitrificans. Evidence from FTIR difference spectro- scopy and site-directed mutagenesis. Biochemistry 39, 1356–1363. 4838 P. Hellwig et al.(Eur. J. Biochem. 269) Ó FEBS 2002 . important step for the understanding of the essentials for cytochrome c oxidase activity and coupled proton pump- ing, the crystal structure of the aberrant. chains and reflects its protonation with the reduction of the protein. We note that these assign- ments are tentative and can be supported by the combi- nation