proton uptake by cytochrome c oxidase on reduction and on ligand binding

ELSEVIER Biochimica et Biophysica Acta 1186 (1994) 19-26

BB Biochi~i et Biophysica A~ta

Proton uptake by cytochrome c oxidase on reduction and on ligand binding

Roy Mitchell, Peter R. Rich *

Glynn Research Institute, Bodmin, PL30 4A U, UK

(Received 28 September 1993; revised manuscript received 7 February 1994)

Abstract

On reduction, cytochrome oxidase was found to take up 2.4 + 0.1 protons in the pH range 7.2-8.5, of which 2 are associated with the binuclear centre, and the remaining fractional proton with haem a /Cu A. Ligation to oxidised cytochrome oxidase of the azide, formate, fluoride or cyanide anions is accompanied by uptake of one proton. In the case of the reduced enzyme, no protonation changes are observed on binding 0 2 (Hall6n S. and Nilsson T. (1992) Biochemistry 31, 11853-11859) or CO. Cyanide binding to reduced oxidase is, in contrast, still accompanied by uptake of a proton. These findings are discussed in terms of our previously-published proposal for the ligand chemistry of the binuclear site. The results overall suggest a principle of electroneutrality of redox and iigand state changes of the binuclear centre, with charge compensations provided only by protonation reactions.

Key words: Cytochrome c oxidase; Protonation; Ligand binding; Carbon monoxide; Anion ligation

1. Introduction

From measurements of protonation changes accompanying the steps of the four-electron catalytic cycle of 0 2 reduction to water [1], it appeared that reduction of haem a3/CUl3 at the oxygen-binding site of cytochrome oxidase must be accompanied by the uptake of two protons. A simple model was proposed in which protonation of hydroxide ligands of the binuclear site metals occurred on reduction. The present work was undertaken in order to measure directly the uptake of protons on reduction, and to study the effects of ligand binding.

From the pH dependence of inhibition by cyanide, azide and formate [2-4] it was suggested that these ligands are bound in their protonated forms. From infrared spectroscopy it now appears that cyanide is bound as M - C - N rather than as M-N-= C -H [5], so that in this case, and for the other ligands, these observations are consistent with displacement of a hydroxide ligand by the inhibitor. Where spectral observations of binding constants to the oxidised enzyme

* Corresponding author. Fax: +44 208 821575.

0005-2728/94/$07.00 © 1994 Elsevier Science B.V. All rights reserved SSDI 0005-2728(94)00021-V

have been carried out, these show a p H dependence similar to that of inhibition [3]. Direct measurements of proton uptake, however, using excess azide, showed much less than the expected one proton per haem a 3, particularly at high p H [6]. With other haem proteins, results differed according to the protein species [7]. With reduced enzyme, on the other hand, such evidence as is available is consistent with displacement of a water ligand. No pH change occurs on forming the 0 2 adduct [8]. The p H dependence o f the dissociation constants of hydrazine and hydroxylamine indicates binding of the deprotonated (uncharged) forms [9].

By direct pH measurements on binding of a range of ligands to both oxidised and reduced enzyme, we have a t tempted in this work to reveal any general pat tern of such ligand-induced protonation changes.

2. Materials and methods

Beef heart oxidase was prepared as previously de- scribed (prep. D of Ref. [10]) by a modified Kuboyama method, but made up finally with a bicine concentration of 5 mM instead of 50 mM. This method yields largely ' fast ' enzyme, i.e., showing little or no slow

20 R. Mitchell, P.R. Rich/Biochimica et Biophysica Acta 1186 (1994) 19-26

phase of cyanide binding. Optical measurements were carried out in a single-beam spectrophotometer con- structed in-house, using 1 cm light path cuvettes. For anaerobic experiments (Figs. 1-3), the cuvette was completely filled with 3.1 ml de-oxygenated medium so as to exclude any gas phase, and fitted with a ground glass stopper, bored through to allow insertion of sy- ringe needles when making additions. The 3.0 ml remaining in the cuvette was stirred by means of a glass-covered magnetic follower. A hood just large enough to enclose the top of the cuvette and stopper, but with a small entry port for additions, was kept continuously flushed with nitrogen. By this means, the aerobic experiments were also carried out under a nitrogen a tmosphere to avoid uptake of CO 2 and con- sequent acidification. Illumination for photoreduction was provided by a 300 W projector lamp. Photolysis of the CO compound was carried out using a frequency- doubled N d : Y A G laser (Spectron Laser Systems, Rugby, UK) with a pulse energy of 115 mJ at 532 nm, pulse half width 10 ns. p H measurements were carried out using either Phenol red or Cresol red, calibrating with standard acid and alkali solutions, and checking the absolute value with a p H electrode after each experiment. For further details, see figure legends.

D 2

I o . o 2 A I

5 rain ,

KOH KOH

Fig. ]. Proton uptake and release associated with redox cycling at pH 8.0. The medium (see Methods) contained 50 mM potassium sulfate, 1 mM Mes, 50/zM Cresol red, 10/~M FMN, 10/~M anthraquinone 2,6-disulfonate and (initially) 1/zM hexammineruthenium trichloride with 3 p.M cytochrome oxidase. After illumination to deplete residual oxygen and reduce the oxidase, as monitored at 604-502 nm (upper trace) the enzyme was alternately oxidised with 10 /zl 0 2- saturated H20 and photoreduced, monitoring pH at 576-620 nm (lower trace). Calibrations were carried out with 0.33 p.M n 2 s o 4 and 0.66/.~M KOH as 2/zl additions, causing pH steps of about 0.08 unit. Small arrows indicate short periods of illumination, 20 s initially, thereafter 10 or 5 s. The second and third redox cycles are shown, also the effect of a second addition (--* 2 /zM) of hexammineruthenium(III) to the reduced system.

3. Results

3.1. Proton uptake on reduction

The enzyme was photoreduced using FMN [11] with Mes as the ult imate reductant. At p H 8.0, where Mes has little buffering action, this system acts purely as a hydrogen donor, provided there is negligible net reduction of FMN, so that reduction of oxygen to water causes no pH change. With a suitable redox mediator such as hexammineruthenium, allowing rapid transfer of electrons between FMN and cytochrome oxidase, the enzyme can easily be subjected to repeated cycles of reduction and oxidation by alternately illuminating and reoxidising with oxygen. If the enzyme were an electron acceptor only, i.e., in the absence of proton uptake, reduction of cytochrome oxidase would be expected to be accompanied by release of four protons. Anthraquinone 2,6-disulfonate, with a midpoint potential of - 1 8 4 mV at p H 7 lying between hexammineruthenium (78 mV) and the more strongly-reducing FMN ( - 215 mV), was used as a hydrogen-donating redox buffer, causing no pH change on partial oxidation with oxygen, and preventing significant redox change of the FMN.

Fig. 1 shows the effect on pH, measured at 576-620 nm in the presence of Cresol red, of switching between fully-reduced and almost fully oxidised states of the enzyme. Reduction was continued a little beyond the

point where the enzyme appeared to be fully reduced, until no further pH change occurred, so that the hexammineruthenium, but little or no anthraquinone disulfonate, was also reduced. Oxygen was then added in slight excess over reducing capacity, so that a second addition prior to re-reduction (not shown) had little or no effect on either haem oxidation or pH. On adding the oxygen virtually all of the hexammineruthenium becomes oxidised, as was verified by the increased proton uptake when its concentration was increased, as does most of the oxidase. The extent of oxidation of the oxidase was estimated from a no-indicator control, t ak ing A~604_620 = 23.5 mM -1 cm -1 (based on Ae6o3 red - ox = 25 m M - l cm-x [12]). The 604-502 nm pair shown in Fig. 1 is pH-insensitive but, as well as measuring haem redox changes, is liable to interference from small redox changes of the FMN.

The number of protons coupled to redox changes of the oxidase were calculated from the extent of the oxygen-induced p H step. With a pure electron donor like hexammineruthenium(II) , one proton per electron will be consumed when it is oxidised by oxygen from the reaction:

4Ru(NH3)~ + + 4H + ,4Ru(NH3)36 + + 2 H 2 0

(1)

R. Mitchell, P.R. Rich/Biochirnica et Biophysica Acta 1186 (1994) 19-26 21

For fully-reduced oxidase, which contains four electrons and n protons, (4 - n) protons will be consumed on oxidation by 02 according to the reaction:

oxidase(4e-,nH +) + 0 2 + (4 -- n )H +

, oxidase + H 2 0 (2)

Hence, the difference between the number of protons taken up on oxidation by molecular oxygen of the hexammineruthenium(II) plus reduced oxidase and the number of electrons transferred to molecular oxygen gives the number of protons released from the oxidase oxidation.

At pH 7, any small amount of reduced anthraquinone sulfonate which might also become oxidised makes no difference since it is a hydrogen atom donor:

2AQDSH2 + 02 ~ 2H20 (3)

However, since the pKal of anthraquinone-2,6-disulfonate is 8.1 [13], proton uptake would occur on oxidation at pH values of 8 and above, which could hence lead to an overestimate of the number of redox protons associated with the oxidase if the anthraquinone disulfonate had become significantly reduced before the oxygen pulse. That this was not the case was confirmed by repeating such experiments with anthraquinone-l,5-disulfonate ( P K a l = 10.5, Era7 = - 170 mV [ 1 3 ] ) .

A small dilution correction was made for the O E- saturated water, and the nearly-negligible no-indicator control at 576-620 nm was subtracted. Several cycles of oxidoreduction could be carried out, with only a slight decrease in the extent of the redox change. Values obtained over a range of pH are shown in Table 1. There appears to be no significant effect of pH on the stoichiometry of proton release and, taking all values, a mean of 2.4 __ 0.1 is obtained for n in reaction (2) above.

In principle, the acidification seen on photoreduction could also be used to make the calculation, provided that the buffer acting as the ultimate reduc-

Table 1 Protons lost from cytochrome oxidase

pH H +/oxidase + S.E.

Fully-reduced 7.2-7.5 2.4+0.1 (6) 8.0 2.2+0.1 (8) 8.5 2.5+0.1 (4) All 2.4+0.1 (18)

Mixed-valence 8.0 0.4+0.1 (3) CO Compound CN Compound 7.1, 8.1 1.6+0.1 (4)

Summary of the calculated values for the number of protons lost from cytochrome oxidase on complete oxidation of the fully-reduced and mixed-valence states and from full reduction of the cyanide- ligated oxidised enzyme (No. of measurements in parentheses).

tant [11] was purely a hydrogen donor. This appeared to be the case with the present system at pH 8, where the pH change during consumption of residual oxygen before the oxidase became reduced was negligible, but not at pH 7.5 or below.

Under similar experimental conditions to Fig. 1, after photoreduction to a point where haem a became partly reduced, 50/xl gaseous CO was introduced into the stirred cuvette to form the mixed-valence compound, with haem a now mostly reoxidised. After CO binding was complete, further reduction showed the spectral characteristics of pure haem a reduction in the visible region (Cu A was not monitored, but was presumably also reduced). The concomitant acidification, measured at 570-626 nm (not shown) indicated that 0.4 protons were taken up per haem a reduced (Table 1). Thus, assuming that CO has no effect on haem a/CUA, of the 2.4 protons taken up by unli- ganded oxidase, approx. 0.4 are associated with haem a / C u A reduction, with the remaining 2 protons associated with the binuclear centre.

In order to elucidate further the relationship of the individual redox centres with proton uptake, we have also carried out some experiments using cyanide ligated enzyme where both coppers as well as haem a can be reduced independently of haem a 3. pH was measured at 578-625 nm and haem a reduction at 605-502 nm. The enzyme was incubated 15-20 min at room temperature with 10 mM cyanide, by which time binding to the fast enzyme was complete. The enzyme (3 /xM) was then reduced with 10 /~M NADH, in the presence of 0.1 ~M PMS, taking about 1 min:

E-CN(Ox.) + 1.5NADH , E-CN(3e-)

+ 1.5NAD÷+ 1-5H ÷ (4)

We find that this three-electron reduction of oxidised, cyanide-liganded oxidase at pH 7 or 8, is not accompanied by a significant pH change (not shown), implying that about 1.5 protons are taken up by the oxidase (1.6 + 0.1 S.E. from 4 experiments). There is, therefore, some evidence that of the two protons taken up on reduction of the binuclear centre, one may be associated with haem a 3 and one with CuB, or at least that each electron is accompanied by a proton.

3.2. Ligand binding to reduced oxidase

From flow-flash studies of Hall6n and Nilsson [8], it was evident that no net proton uptake or release occurred during the formation of the O 2 adduct of reduced cytochrome oxidase. Likewise, from the lack of protonation changes on forming the peroxy interme- diate by displacement of CO by oxygen from the mixed-valence compound [1,14] it may be inferred that release of CO is accompanied by no pH change. We


A B 5 9 8 nm

5 8 8 n m /

0.005 A

20 ms I I

5 9 8 nm

Fig. 2. Flash photolysis of the fully-reduced CO compound at pH 8. Cytochrome oxidase, to a final concentration of 3.0/xM, was added to a CO-saturated 50 mM solution of potassium sulfate, and reduced with 1 mM ascorbate with 3/~M hexammineruthenium(IIl) chloride as mediator. Photolysis of the CO compound was measured at 588 nm and pH at 598 nm (nearly isosbestic for CO ligation). Traces show averages of 10 pulses. (A) No-indicator control; (B) 50 ~zM Cresol red present, traces scaled up by a factor of 2.1.

Fig. 3 shows the effect on pH of cyanide binding to the reduced enzyme. The slight cyanide-induced acid drift was always found, but its cause is unknown. The small alkaline step on adding cyanide was due to the buffering capacity of the cyanide (pK~ = 9.3), since similar changes were seen with subsequent additions, where relatively little further binding occurred (Fig. 3B). The pH change due to binding was measured by extrapolating back to the point where binding was 50% complete, as determined from simultaneous measurement at 589-496 nm (omitted from Fig. 3A, but similar to that shown for Fig. 3B). The proton uptake associated with the second cyanide addition was subtracted. Cyanide binding was likewise measured, after subtract- ing the second small binding step at 589-496 nm, using an extinction coefficient estimated to be 14.3 mM-~. Six experiments gave an average value of 0.1 H ÷ released per cyanide bound. Since unbound cyanide at pH 7 is almost completely protonated, this indicates uptake of one proton, within experimental error, per cyanide anion bound.

have confirmed this in the case of the fully-reduced CO complex by flash photolysis in the presence of Cresol red at pH 8 (Fig. 2). The extent of the 588 nm signal in Fig. 2A corresponds to about 1.6 /xM CO compound (4.8 nmol). Because of the screening effect of the Cresol red, the photolysis yield was lower in the experiment shown in Fig. 2B, and the traces were accordingly scaled up by a factor of 2.1. Acid and alkali calibrations corresponding to 20 nmol H ÷ (not shown) caused absorbance changes of 0.040 at 598 nm, so that as plotted, release of one proton per CO photolysed would have caused an absorbance change of -0.0096 at 598 nm in B, i.e., approximately the same as the actual 588 nm trace. Clearly, any proton release or uptake is very small, certainly less than 0.1 per CO photolysed.

3.3. Ligand binding to oxidised enzyme

The cyanide binding experiment was repeated under oxidised conditions (not shown) in the absence of FMN, anthraquinone disulfonate and hexammineruthenium, using 5 mM additions of cyanide. In this case, cyanide binding, monitored at 584-556 nm, took several min- utes. No accompanying pH changes were seen, except for a small, instantaneous pH step on addition, independent of binding. The same result was obtained at pH 7 and 8. The conclusion, as above, is that close to one proton per cyanide anion is bound.

In the case of formate (pK a = 3.75), in order to achieve a sufficient binding rate, a concentration of 5 mM was required at pH 7, 20 mM at pH 8. Subsequent

A B Z 2min

H o4 . ,so , J .~____ .~__ ~ " ~ - ~ - ~

KCN "l I 0.02 A 559 - 622nm

2 mln | ~ . . . . ' ' k, . ~ + + +

,KOH KOH KCN KCN H2SO 4 KOH

Fig. 3. Cyanide binding to reduced oxidase. Conditions were as for Fig. 1, except Ches instead of Mes, Phenol red instead of Cresol red (pH 7.1), anthraquinone disulfonate omitted. After complete reduction, measured at 604-622 nm, 3/zl additions of anaerobic M KCN (approx. pH 7.2) were made as shown. Calibrations as Fig. 1. (A) pH measured at 559-622 nm. (B) No-indicator control at 559-622 nm, also showing cyanide

binding at 589-496 nm.

R. Mitchell, P.R. Rich/Biochimica et Biophysica Acta 1186 (1994) 19-26 23

additions could not be used as controls for the pH step due to formate buffering, since the degree of acid dissociation was markedly concentrat ion-dependent in this region. Rather than attempting to estimate the total pH change, the binding and pH time-courses were compared as shown in Fig. 4. The scaling factor was based o n p H calibrations (not shown), taking the extinction coefficient for formate binding at 412-430 nm as 25 mM -1 cm -1 (J. Moody unpublished data). This value is appropriate for the approx. 70% of the enzyme population that binds formate relatively rapidly. The wavelength pairs showed negligible cross-interference, as confirmed by a no-indicator control. Ap- prox. 1.0 H ÷ was taken up per cytochrome oxidase present at both pH 7 and 8.

Azide (pK a = 4.59) shows two spectrally distinct phases of binding, both rapid, but with different affini- ties, see Fig. 5 and [15] where Wever et al. found a 10-fold faster dissociation rate at 558 nm than when measured at 432 nm. Only the high-affinity binding was examined for protonation changes. Clearly, the high-affinity binding is not easily quantifiable from the spectrum in the Soret region. However, the peak at 562 nm appears to be very little affected by the low-affinity binding which, in addition to the well-known Soret red-shift, gives rise to a peak at 606 nm (pH 6) to 615 nm (pH 8) in the 20 mM - 2 mM difference spectrum (not shown). Measurements of the pH effect of azide binding were carried out with 0.2 mM additions under conditions similar to those of Fig. 4, except that ferricyanide was omitted, and Cresol red was used as indicator, pH was measured at 585-640 nm. The extent of binding at the high-affinity site was estimated from the peak to trough absorbance at about 562-540 nm (the peak position is slightly pH-dependent) in no-indi-

I

0.02 A

2 rain I

xeXeX= xixexix= xBxexex= x=x=xex= x=x=

• l l x lX

iX X

i x

X

)msxmXilXe X iX iX l xe

Fig. 4. pH change accompanying formate binding at pH 7.0. The cuvette contained 1.0 ml of 50 m M potassium sulfate, 0.1 mM potassium ferricyanide, 50 /zM Phenol red and 3 ~zM cytochrome oxidase. 5 m M potassium formate (5 M solution pH 7.95) was added as indicated by the arrow. Formate binding at 412-430 nm is shown (crosses), together with pH at 564-454 nm, scaled up by a factor of 2.86, so that / 1 . 4 5 6 4 _ 4 5 4 for 1 nmol H + consumed = z ~ ` 4 4 1 2 _ 4 3 0 for 1 nmol formate bound.

=,x5 A

2

0.2 x,~.=.,,~o.2~

2

I I I I I

500 6 0 0

X (nm) Fig. 5. Azide binding spectra: pH and concentration effects. Medium contained 50 mM potassium phosphate and 0.5 mM EDTA, with 3.0 ;zM cytochrome oxidase. A, pH 6.0; B, pH 7.0; C, pH 8.0. Numbers against traces indicate millimolar concentration of azide.

cator control experiments, assuming binding at this site to be complete at 40 mM azide. At 0.2 mM azide, typically 65-75% binding was found at pH 7.0, but only 30-40% at pH 8.0. In some experiments, a saturating amount of azide was added between the first and second 0.2 mM additions, so that no significant binding occurred with the second 0.2 mM addition. The same result was obtained if the large addition was omitted and the slight additional binding corrected for. Within experimental error, one proton is taken up per azide anion bound (Table 2).

Fluoride ligation to our 'fast' enzyme caused spectral changes similar to those shown for 'resting' enzyme by Muijsers et al. [16] in the visible region, but the Soret band shift was larger, giving a peak to trough extinction coefficient in the difference spectrum of about 21 mM -1 cm -1, approximately double that of the resting enzyme. Proton uptake during binding of 5 mM fluoride was measured using Phenol red at pH 7.0, monitoring pH at 576-618 nm and fluoride ligation at 642-666 nm taking ~5642_666 as 1.9 mM - l cm - l (see

24 R. Mitchell, P.R. Rich/Biochirnica et Biophysica Acta 1186 (1994) 19-26

Table 2 Protons bound to cytochrome oxidase

Ligand Oxidase state pH H ÷ taken up Ref. _+ S.E.

CO R 8 0 02 R 7-8.9 0 N2H 4 R 7.4-9 0 NH2OH R < 7.4-8.5 0 CN- R 7 0.9_+0.1 (6)

O 7, 8 1.0_+0.1 (9) HCOO- O 7, 8 1.0 N 3 O 7, 8 1.1_+0.2 (6) F- O 7 0.9_+0.1 (4)

[8] [9] [91

Summary of the calculated numbers of protons bound to cytochrome oxidase on binding of ligands. Note that the proton changes refer to binding of the form of the ligand shown. Several of these ligands (e.g. cyanide at pH values below 9) will be protonated in solution, so that overall proton uptake from the solution will be less than the calculated value.

Fig. 3 of Ref. 7), a value that we have confirmed. We find binding to more than 80% of the oxidase molecules following a 5 mM addition, consistent with a dissociation constant of around 1 mM rather than the 7-10 mM reported for resting oxidase [16]. Again, we find close to one proton taken up per fluoride anion bound (Table 2).

4. Discussion

Measurements by Oliveberg et al. of p H changes on oxidation of fully-reduced cytochrome oxidase showed approx. 3 protons released per oxidase [14], later revised to 2.6 at p H 7 -8 [8], with which our present measurement of 2.4 protons in the range pH 7.5-8.5 is in reasonable agreement . Of these protons, approx. 0.4 appears to be associated with haem a / C u A, and the remaining two with the binuclear centre. From our observations that displacement of CO from the mixed- valence CO compound by oxygen to form the peroxy (P) state causes no proton uptake, while P formation from oxidised enzyme and H 2 0 2 causes no proton release, we had already concluded that the reduced binuclear centre contains two additional net protons compared with the oxidised centre [1]. It is an interest- ing question whether reduction of cyanide-liganded oxidase, where both coppers as well as haem a can be reduced independently of haem a3, is accompanied by uptake of more than 0.4 protons. Published values of 1.0 [17] and 0.6 [18] protons per electron indicate that this is the case. However, some caution is necessary as the former figure appears to refer to electrons transferred from cytochrome c to the enzyme, while the latter refers to electrons transferred to haem a, and where less than three electrons are taken up, all three centres will not be equally occupied. Under conditions

where we have been careful to ensure complete reduction of both coppers and haem a, we find a total of 1.6 protons taken up.

In view of the generally-accepted pH dependence of the redox midpoints of both haems and CuB, the finding that protons are taken up on reduction was to be expected. The measured amounts, as reported both here and elsewhere [8], are consistent with a slope approaching - 6 0 m V / p H each for haem a 3 and CUB, and - 3 0 m V / p H for the sum of effects on haem a and Cu A. Only the value suggested for Cu B is significantly different from previous estimates. From CO binding studies, Lindsay et al. [19] concluded that the preliminary 2-electron reduction is affected by pH to the extent of some - 3 0 mV per unit. Assuming that haem a 3 has a slope close to - 6 0 mV [20], Cu s would be pH-independent . However, from the anticoopera- tive interaction of Cu B with haem a in cyanide-liganded oxidase, it has been inferred that the pH dependence is similar to that of haem a [21,22]. On the other hand, in anaerobic redox titrations, the appearance of the high- spin haem E P R signal as the potential is lowered, presumably reflecting Cu B reduction, occurs at a potential about 80 mV lower at pH 8.5 than at 7.0 [23]. There is therefore some support from pH dependence studies of a one proton per electron coupling for Cu B, in accordance with uptake measurements.

As regards ligand binding to the reduced enzyme, no pH changes occur with CO, whereas with cyanide anion, a proton is taken up. With oxidised enzyme, all ligands tested were taken up with a proton (Table 2). Although our current preparative method yields enzyme which binds cyanide nearly monophasically ( > 90% fast), it cannot be assumed that it is therefore homogeneous. Formate binding, for example, shows at least two phases [10], of which we here deal with only the larger and more rapid (Fig. 4). Neither can it be assumed that the classical view of binding of a single ligand group on haem a 3 is correct. In the case of cyanide and of azide, it appears increasingly likely, particularly at high ligand concentrations, that Cu B may be involved, either as a separate binding site, or with a bridging ligand [5,24,25]. Starting with the per- haps rather naive prediction, based on our previously- published model [1], that a hydroxide ion should be displaced from oxidised haem a 3, we find that pH changes on ligation are consistent with this. Either a hydroxide is displaced, or a proton is taken up with the ligand. With the reduced enzyme, no pH changes are predicted or occur on replacing the postulated water ligand with CO, 0 2 [8], N2H 4 [9] or N H 4 O H [9]. On the other hand, we find that with cyanide a proton is taken up with the anion, making a total of about 3.4 protons on reduction and cyanide anion binding. If cyanide is indeed bound in its anionic state, the situa- tion as regards the electrical charge of the centre,

tL Mitchell, P.R. Rich/Biochimica et Biophysica Acta 1186 (1994) 19-26 25

where the Fe E+ charge is neutralised by the porphyrin ring, appears to be unusual. To our knowledge, no other anion has been reported to bind to reduced oxidase (the carbon rather than the nitrogen probably provides the Fe bond [5]). It seems very plausible, since alkyl isocyanides can be ligands of reduced haem a 3 [26], that hydrocyanic acid binds in its tautomeric HNC- state, displacing H20. It has been suggested [5] that the unusually high cyanide binding affinity of reduced cytochrome oxidase compared with other ferrous haem proteins is a result of hydrogen bonding. This, of course, suggests a simple law that holds for all cases here tested, that anion binding to cytochrome oxidase is accompanied by uptake of a proton (or release of OH-) while binding of uncharged ligands causes no pH change. More generally still, as regards the binuclear centre, uptake or release of negative charges (anions or electrons) is always charge-compensated by protons. On the basis of recent FTIR studies [27], it has been proposed that as many as three cyanide binding sites, two on CUB, one on haem a 3 may be occupied under fully-reduced conditions. The question of the number of cyanide ligands per oxidase cannot be resolved by our pH change measurements, which, since the proton is already associated with the free cyanide, merely show that one proton per cyanide is taken up. In the case of azide, by measuring at only 0.2 mM, we aimed to avoid significant occupancy of the low-affinity site [25]. This may explain the difference between our results and those of Nicholls and Wrigglesworth [6], where at most less than half a proton was taken up per enzyme molecule on adding excess azide, although it seems more likely to be due to their use of 'resting' enzyme, where the high affinity site may not be available for azide binding. At our low concentration of azide, and also in the case of formate and of fluoride, the stoichiometry of proton uptake strongly suggests single ligand binding to the enzyme molecule.

The data are as expected from principles of electrical neutrality given that, except for added ligands, the binuclear centre is probably accessible to no ions present other than H ÷ (or OH-). We conclude that the present findings are compatible with our proposal [1] of a protonatable hydroxide ligand on haem a 3 and on Cu B . However, this represents the simplest case, and protonation changes of other groups remains possible. It might be argued against our proposal that such a hydroxy form has in fact been identified by resonance Raman spectroscopy, but only as a transient intermedi- ate species, with a lifetime measured in milliseconds [28-30]. It should be borne in mind, however, that these observations were made in the presence of CO and reductant. Under other conditions, where the oxidase was directly photoreduced, with no CO present, the species exhibiting a signal attributed to Fe-OH disappeared only as the enzyme slowly relaxed to the

resting, or slow state [31]. We do not find this disap- pearance surprising, since it seems likely that the de- creased rates of reduction and ligand binding of the binuclear centre in the slow form are caused by a modified ligand structure [10]. This might, for example, come about through loss of H:O to leave a bridging oxygen.

There appears to be an inconsistency between our observation that the P ~ F transition is accompanied by uptake of one proton as well as one electron and the pH dependence of the P / F ratio as reported by Wikstr6m [32]. This dependence indicates a net 2-proton uptake, and to some extent the calculation of the number of protons translocated at this step depends on this value [33]. A possible explanation is that the pH dependence measurement was made in mitochondria with a high transmembrane electrical potential, which is expected to raise the local 'pH' of the binuclear centre so that an additional site might be protonatable. This discrepancy requires further investigation.

Acknowledgements

We are grateful to the Wellcome Trust for a grant (No. 034505/Z/91/Z) supporting this work; also to Robert Harper for preparation of the enzyme, to Si- mon Brown and John Moody for helpful comments on the manuscript, and to the Glynn Research Founda- tion for general support and facilities.

References

[1] Mitchell, R., Mitchell, P. and Rich, P.R. (1992) Biochim. Bio- phys. Acta 1101, 188-191.

[2] Stannard, J.N. and Horecker, B.L. (1948) J. Biol. Chem. 172, 599-608.

[3] Wever, R., Muijsers, A.O., Van Gelder, B.F., Bakker, E.P. and Van Buuren, K.J.H. (1973) Biochim. Biophys. Acta 325, 1-7.

[4] NichoUs, P. (1976) Biochim. Biophys. Acta 430, 13-29. [5] Yoshikawa, S. and Caughey, W.S. (1990) J. Biol. Chem. 265,

7945-7958. [6] Nicholls, P. and Wrigglesworth, J.M. (1979) Oxidases and Re-

lated Redox Systems (King, T.E., Mason, H.S. and Morrison, M., eds.), pp. 1149-1160, Pergamon Press, Oxford.

[7] Millar, F., Wrigglesworth, J.M. and Nicholls, P. (1981) Eur. J. Biochem. 117, 13-17.

[8] Hall6n, S. and Nilsson, T. (1992) Biochemistry 31, 11853-11859. [9] Kubota, T. and Yoshikawa, S. (1993) Biochem. J. 292, 519-524.

[10] Moody, A.J., Cooper, C.E. and Rich, P.R. (1991) Biochim. Biophys. Acta 1059, 189-207.

[11] Traber, R., Kramer, H.E.A. and Hemmerich, P. (1982) Bio- chemistry 21, 1687-1693.

[12] Antalis, T.M. and Palmer, G. (1982) J. Biol. Chem. 257, 6194- 6206.

[13] Clark, W.M. (1960) Oxidation-Reduction Potentials of Organic Systems, Bailli~re, Tindall & Cox, London.

[14] Oliveberg, M., Hall6n, S. and Nilsson, T. (1991) Biochemistry 30, 436-440.


[15] Wever, R., Muijsers, A.O. and Van Gelder, B.F. (1973) Biochim. Biophys. Acta 325, 8-15.

[16] Muijsers, A.O., Van Buuren, K.J.H. and Van Gelder, B.F. (1974) Biochim. Biophys. Acta 333, 430-438.

[17] Miiller, M. and Azzi, A. (1986) Biochimie 68, 401-406. [18] Capitanio, N., De Nitto, E., Villani, G., Capitanio, G. and Papa,

S. (1990) Biochemistry 29, 2939-2945. [19] Lindsay, J.G., Owen, C.S. and Wilson, D.W. (1975) Arch.

Biochem. Biophys. 169, 492-505. [20] Blair, D.F., Ellis, W.R., Wang, H., Gray, H.B. and Chan, S.I.

(1986) J. Biol. Chem. 261, 11524-11537. [21] Wikstr6m, M. (1988) Ann. NY Acad. Sci. 550, 199-206. [22] Moody, A.J. and Rich, P.R. (1990) Biochim. Biophys. Acta 1015,

205-215. [23] Wilson, D.F., Erecinska, M. and Owen, C.S. (1976) Arch.

Biochem. Biophys. 175, 160-172.

[24] Yoshikawa, S. and Caughey, W.S. (1992) J. Biol. Chem. 267. 9757-9766.

[25] Li, W. and Palmer, G. (1993) Biochemistry 32, 1833-1843. [26] Yamamoto, T. and Orii, Y. (1973) J. Biochem. 73, 1049-1059. [27] Tsubaki, M. (1993)Biochemistry 32, 164-173 [see Correction:

Biochemistry 32, 9262]. [28] Han, S., Ching, Y. and Rousseau, D.L. (1990) Nature 348.

89-90. [29] Varotsis, C., Zhang, Y., Appelman, E.H. and Babcock, G.T.

(1993) Proc. Natl. Acad. Sci. USA 90, 237-241. [30] Babcock, G.T. and Varotsis, C. (1993) J. Bioenerg. Biomemb.

25, 71-80. [31] Hart, S., Ching, Y.-C. and Rousseau, D.L. (1989) J. Biol. Chem.

264, 6604-6607. [32] Wikstr6m, M. (1988) Chem. Scr. 28A, 71-74. [33] Wikstr6m, M. (1989) Nature 338, 776-778.

proton uptake by cytochrome c oxidase on reduction and on ligand binding

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