In contrast, mutations of this residue E78II in the Paracoccus denitrificans cytochrome c oxidase do not affect its catalytic activity at all E78IIQ or reduce it to about 50% E78IIA; in t
Trang 1Paracoccus denitrificans cytochrome c oxidase
Oliver-M H Richter1, Katharina L Du¨rr1, Aimo Kannt2, Bernd Ludwig1, Francesca M Scandurra3, Alessandro Giuffre`3, Paolo Sarti3and Petra Hellwig4
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 (CuA) located in the periplasmic domain of
subunit II Subsequently heme a and another binuclear
center consisting of heme a3 and CuB, 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 a3 [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 (E78II) in the Paracoccus denitrificans cytochrome c oxidase do not affect its catalytic activity at all (E78IIQ) or reduce it to about 50% (E78IIA); in the latter case, the mutation causes no drastic decrease in heme a3reduction 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 E78II 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)1excludes the protonation
of E78IIin the course of the redox reaction in the studied pH range, although shifts of vibrational modes at 1570 and 1400 cm)1reflect 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 E78IIin the Paracoccus enzyme
Abbreviations
SHE¢, standard hydrogen electrode (at pH 7); SVD, singular value decomposition; VIS, visible.
Trang 2expected 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 bo3
quinol oxidase (E89IIQ⁄ 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 bo3 oxidase,
mutation of E89IIto Q has been proposed to block the
reduction of the heme-copper binuclear center, lending
support to the idea that E89II 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 a3reduction 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 E78II in the Paracoccus
cyto-chrome c oxidase are characterized in terms of
cata-lytic and proton pumping activity, kinetics of heme a3
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 E78II 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 E78IIQ matched that of wild-type, while the activity of the E78IIA mutant was lowered
to about 50% (Table 1)
Given the lower activity of the E78IIA 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 a3 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
a
Stopped-flow apparatus;bpotentiometric method.
Trang 3The time course of heme a3 reduction in the E78IIA
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
E78IIA mutation only slightly affects the rate of
heme a3 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
a3 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 O2 consumption ( 80 mol O2Æmol6
enzy-me)1Æs)1) As previously shown for the Paracoccus enzyme [26], this is expected given the unfavorable redox equilibrium between heme a and heme a3, if it is taken into account that our measurements were carried out in the absence of NO acting as a trapping ligand for reduced heme a3[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 E78IIQ and the E78IIA mutant oxidases show unimpaired proton-pumping (Table 1) The H+⁄ e– ratio for E78IIQ was found to be around 1.0 when determined by the reductant pulse potentio-metric method [28] The E78IIA 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 E78IIA 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 E78IIA and E78IIQ 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 a3(inset) (Bottom panel) Time courses of heme a3reduction
as obtained by SVD analysis Fitted rate constants with relative
amplitudes in brackets: wild-type enzyme, k1¼ 18.7 s)1 (70%),
k2¼ 1.1 s)1 (30%); E78 II A mutant, k1¼ 14.5 s)1 (35%), k2¼
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].
Trang 4charge 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⁄ a3, CuB or CuA) 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 E78IIQ (A) and E78IIA (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 subspec-traction of the differ-ence spectra for the mutant enzymes from wild-type (C) The glutamine of E78IIQ 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(NH2) 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 E78IIQ (C, line) and E78IIA (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.
Trang 5[32] The increase of a difference signal at 1676 and at
1608 cm)1, however, is also observed for the E78IIA
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)1and
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–)smodes 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 E78II 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 E78II side chain at pH 7,
electro-chemically induced FTIR difference spectra were
obtained for the E78IIQ 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 E78IIQ
mutant enzyme equilibrated at the same pH values (B)
as well as the direct comparison of wild-type (dotted
line) with the E78IIQ 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)1indicates that residues close to the surface, or in the vicinity of sev-eral water molecules are involved here Comparing the spectra of the E78IIQ 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 E78IIto 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 pKA
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 E78IIQ (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 E78IIQ mutant at pH 4.8 (C).
Trang 6value 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 denitrificanscytochrome 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 bo3 ubiquinol oxidase [20]
and E101 in the R sphaeroides aa3cytochrome 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 bo3) and to
8 and 14% (R sphaeroides aa3), 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 E78IIA to about 50% of the wild-type level, the
E78IIQ mutation shows unchanged catalytic
compet-ence More importantly, proton-pumping is essentially
unaffected in both mutant enzymes It therefore seems
that E78II lacks a direct role in the overall enzymatic
reaction of the Paracoccus enzyme in contrast even to
the closely related R sphaeroides aa3 oxidase [21]
Interestingly, mutation of E78II to A causes only a
slight effect on the kinetics of heme a3reduction, 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 a3 reduction is considered as diagnostic
for mutated residues in the K channel, we conclude
that in the P denitrificans aa3 cytochrome c oxidase
E78II does not represent the dominant entry point for protons into the K-channel We used redox-induced FTIR difference spectroscopy to monitor whether E78II 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 E78IIA, 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 aa3 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 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 ctaC to 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 Fv antibody fragment was per-formed as described [33,36] For electrochemistry the pro-tein samples were further concentrated to 0.5 mm aa3 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
Trang 7dodecyl-b-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 20C 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 N2-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, 4C) 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, 4C) and the resulting liposome
suspension stored at 4C
Proton translocation of reconstituted oxidase
A suspension containing 0.4 lm E78IIA or wild-type oxidase
proteoliposomes, 60 lm 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 25C and 555.6 nm, which was deter-mined to be the isosbestic point for cytochrome c under the experimental conditions E78IIQ 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)1range, 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; Em:)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
Trang 8This 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
coxidase (cytochrome aa3) from Paracoccus
denitrifi-cans Proc Natl Acad Sci USA 77, 196–200
2 Haltia T, Puustinen A & Finel M (1988) The
Paracoc-cus denitrificanscytochrome aa3has 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 colieliminates 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 aa3and 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 aa3) 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 ba3-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 denitrificanscytochrome 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 bo3from 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 sphaeroidesresult 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
Trang 9(1998) Involvement of glutamic acid 278 in the redox
reaction of the cytochrome c oxidase from Paracoccus
denitrificansinvestigated 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 propionate13C 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
coxidase 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
coxidase 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 ba3-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 a3 redox reactions in the cytochrome c oxidase from Para-coccus denitrificans: separation of heme a and a3 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 (H2O) 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 denitrificansmutants 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