Báo cáo Y học: Role of histidine 42 in ascorbate peroxidase Kinetic analysis of the H42A and H42E variants

doi:10.1046/j.1432-1033.2002.02998.x

Eur. J. Biochem. 269, 3182–3192 (2002) (cid:3) FEBS 2002

Role of histidine 42 in ascorbate peroxidase Kinetic analysis of the H42A and H42E variants

Latesh Lad1, Martin Mewies1, Jaswir Basran2, Nigel S. Scrutton2 and Emma L. Raven1 1Department of Chemistry and 2Department of Biochemistry, University of Leicester, UK

Compound I (k¢1). Values for k¢1 and Kd were, respectively, 4.3 ± 0.2 s)1 and 30 ± 2.0 mM (H42A) and 28 ± 1.0 s)1 and 0.09 ± 0.01 mM (H42E). Photodiode array experi- ments for H42A revealed wavelength maxima for this intermediate at 401 nm, 522 nm and 643 nm, consistent with the formation of a transient [H42A–H2O2] species. Rate constants for Compound I formation for H42A were independent of pH, but for rAPX and H42E were pH- dependent [pKa ¼ 4.9 ± 0.1 (rAPX) and pKa ¼ 6.7 ± 0.2 (H42E)]. The results provide: (a) evidence that His42 is critical for Compound I formation in APX; (b) confirmation that titration of His42 controls Compound I formation and an assignment of the pKa for this group; (c) mechanistic and spectroscopic evidence for an intermediate before Com- pound I formation; (d) evidence that a glutamic acid residue at position 42 can act as the acid–base catalyst in ascorbate peroxidase.

Keywords: ascorbate peroxidase; Compound I; histidine 42.

To examine the role of the distal His42 residue in the cata- lytic mechanism of pea cytosolic ascorbate peroxidase, two site-directed variants were prepared in which His42 was replaced with alanine (H42A) or glutamic acid (H42E). Electronic spectra of the ferric derivatives of H42A and H42E (pH 7.0, l ¼ 0.10 M, 25.0 (cid:1)C) revealed wavelength maxima [kmax (nm): 397, 509, (cid:2) 540sh, 644 (H42A); 404, 516, (cid:2) 538sh, 639 (H42E)] consistent with a predominantly five- co-ordinate high-spin iron. The specific activity of H42E for oxidation of L-ascorbate (8.2 ± 0.3 UÆmg)1) was (cid:2) 30-fold lower than that of the recombinant wild-type enzyme (rAPX); the H42A variant was essentially inactive but activity could be partially recovered by addition of exogen- ous imidazoles. The spectra of the Compound I intermedi- ates of H42A [kmax (nm) ¼ 403, 534, 575sh, 645] and H42E [kmax (nm) ¼ 404, 530, 573sh, 654] were similar to those of rAPX. Pre-steady-state data for formation of Compound I for H42A and H42E were consistent with a mechanism involving accumulation of a transient enzyme intermediate (Kd) followed by conversion of this intermediate into

mechanistic, structural and spectroscopic scrutiny that it has become the benchmark against which all other peroxidases are measured.

The plant peroxidase superfamily has been classified [1] into three major categories: class I contains the enzymes of prokaryotic origin, class II contains the fungal enzymes (e.g. lignin peroxidase) and class III manganese peroxidase, contains the classical secretory peroxidases [e.g. horseradish peroxidase (HRP)]. The most notable member of the class I peroxidase subgroup is cytochrome c peroxidase (CcP), which was first identified in 1940 [2]. In spite of the fact that CcP has some rather unusual features, most notably the existence of a stable tryptophan radical during catalysis [3–6] and the utilization of a large macromolecular substrate (cytochrome c), it has been the subject of such intense

that

More recently, it has been possible to isolate and purify in good yields a second member of the class I peroxidase subgroup, ascorbate peroxidase (APX) [7,8]. Ascorbate- dependent peroxidase activity was first reported in 1979 [9,10] and the enzyme catalyses the reduction of potentially damaging H2O2 in plants and algae using ascorbate as a source of reducing equivalents [11,12]. APX was known from sequence comparisons [13] to contain the same active- site Trp residue (Trp179) as is used by CcP (Trp191) during catalysis. With high-resolution structural information avail- able for the recombinant pea cytosolic enzyme (rAPX) [14] (Fig. 1), APX has provided a new opportunity to reassess the functional properties of CcP and to determine whether it is indeed representative of class I peroxidases. As detailed functional information has emerged, however, it seems that APX has several rather curious features of its own, and, in some ways, more questions have been raised than answered. (In fact, even the current classification of APX as a class I enzyme has been recently questioned [15].) For example, Trp179 in APX is not a necessary requirement for oxidation of ascorbate [16] and there is general agreement from kinetic [17–19] and EPR data [20] the initial product (Compound I) of the reaction of APX with H2O2 is a porphyrin p-cation intermediate and not a protein-based trytophan radical. Equally intriguing is the existence of a

Correspondence to E. L. Raven, Department of Chemistry, University of Leicester, University Road, Leicester LE1 7RH, UK. Fax: + 44 (0)116 2523789, Tel.: + 44 (0)116 2522099, E-mail: emma.raven@le.ac.uk Abbreviations: APX, ascorbate peroxidase; pAPX, wild-type pea cytosolic APX; rAPX, recombinant wild-type pea cytosolic APX; H42A, a variant of rAPX in which His42 has been replaced with alanine; H42E, a variant of rAPX in which His42 has been replaced with glutamic acid; CcP, cytochrome c peroxidase; HRP, horseradish peroxidase; sh, shoulder. Enzymes: ascorbate peroxidase (EC 1.11.1.11); horseradish peroxidase (EC 1.11.1.7); cytochrome c peroxidase (EC 1.11.1.5). (Received 27 December 2001, revised 8 April 2002, accepted 9 May 2002)

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particular interest is the role of the distal histidine residue (His42 in APX), which has been proposed [31,32] to act as an acid–base catalyst, by accepting a proton from H2O2 and releasing it subsequently as water. Site-directed mutagenesis studies on CcP and HRP have provided evidence to support these predictions (reviewed in [33–37]). In the work reported here, we replaced the distal histidine residue of APX (Fig. 1) with alanine and glutamic acid (H42A and H42E variants, respectively). The aims of the work were severalfold. First, to establish a definitive role for His42 in Compound I formation by replacing it with a residue that is not capable of hydrogen bonding (H42A) and to examine whether other residues at this position are able to act as alternative acid– base catalysts (H42E). Secondly, to use these variants to provide information on the origin of the pH-dependent kinetic rate profile for Compound I formation [17]. Finally, as we anticipated that the replacement of His42 would probably generate variant enzymes that may well have altered kinetic properties, we sought to utilize these alter- ations in intimate mechanism to probe in more detail the formation of Compound I in APX. As such, we present the first spectroscopic evidence for the nature of the interme- diate formed during the reaction of APX with H2O2.

potassium-binding site (not present in CcP), located (cid:2) 8 A˚ from the a-carbon of Trp179; the functional role of this site (if, indeed, there is one) is not yet fully understood.

Fig. 1. Active site of ascorbate peroxidase [14]. Hydrogen bonds (dotted lines) are indicated.

M A T E R I A L S A N D M E T H O D S

Materials

Although steady-state kinetic analyses have been a fairly prominent feature of most of the early literature on APX (reviewed in [12]), pre-steady-state kinetic data have been very much more limited, largely as a result of insufficient quantities of enzyme, and only preliminary mechanistic information is available [16–19,21,22]. The enzyme operates through a classical peroxidase mechanism in which the ferric enzyme is oxidized by two electrons to a so-called Compound I intermediate with concomitant release of one molecule of water, followed by two successive single- electron reductions of the intermediate by ascorbate (HS) to regenerate ferric enzyme.

APX þ H2O2 (cid:4)!

k1 Compound I þ H2O ð1Þ

Compound I þ HS (cid:4)!

k2 Compound II þ S(cid:8)

ð2Þ

L-Ascorbic acid (Aldrich Chemical Co.), guaiacol, imida- zole, 1,2-dimethylimidazole (Sigma Chemical Co.) and the chemicals used for buffers (Fisher) were of the highest analytical grade (more than 99% purity) and used without further purification. H2O2 solutions were freshly prepared by dilution of a 30% (v/v) solution (BDH): exact concen- trations were determined using the published absorption )1Æcm)1 [38]). Aqueous solutions coefficient (e240 ¼ 39.4 M were prepared using water purified through an Elgastat Option 2 water purifier, which itself was fed with deionized water. All pH measurements were made using a Russell pH- electrode attached to a digital pH-meter (Radiometer Copenhagen, model PHM 93).

Compound II þ HS (cid:4)!

ð3Þ

k3 APX þ S(cid:8) þ H2O

Mutagenesis and protein purification

Although Eqn (1) is commonly written as a single step, experimental [23–26] and theoretical [27–30] evidence sug- gests that this is an over-simplification. A more complex mechanism, as first suggested by Poulos and Kraut [31] – involving binding of neutral peroxide to the enzyme, concomitant proton transfer from the bound peroxide to the distal histidine residue, followed by O–O bond cleavage and release of H2O – has been suggested (Scheme 1). Of

Site-directed mutagenesis was performed according to the QuikchangeTM protocol (Stratagene Ltd, Cambridge, UK). Two complementary oligonucleotides encoding the desired mutation were synthesized and purified (PerkinElmer). For the primers were: 5¢-CGTTTGGCATGGGCT H42A, TCTGCTGGTAC-3¢ (forward primer) and 3¢-GCAAAC CGTACCCGAAGACGACCATG-5¢ (reverse primer). For H42E the primers were: 5¢-CGTTTGGCATGGGAATC TGCTGGTAC-3¢ (forward primer) and 3¢-GCAAACC GTACCCTTAGACGACCATG-5¢ (reverse primer). To confirm the identity of the transformants, overnight cultures containing 100 lgÆmL)1 ampicillin were incubated at 37 (cid:1)C with vigorous shaking (250 r.p.m.). The plasmid DNA was isolated using the Hybaid mini-plasmid system and sequenced to confirm the desired mutation. Automated fluorescent sequencing, using New England Biolabs pUC and malE primers, was performed by the Protein and Nucleic Acid Chemistry Laboratory, University of Leices- ter, on an Applied Biosystems 373-Stretch machine, and

Scheme 1. Proposed steps in the formation of Compound I. The mechanism depicts the neutral peroxide-bound and anionic peroxide- bound intermediates. The distal histidine residue that acts as the acid– base catalyst is indicated (B).

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sequence data were analysed using the program SeqED (Applied Biosystems). Individual mutations were confirmed by sequencing across the whole rAPX-coding gene.

pH-dependent assays, a mixed sulfonic acid buffer system (l ¼ 95–110 mM depending on the exact pH) that buffered over the entire pH range was used; reactions were initiated by the addition of H2O2 (to 0.10 mM). In these cases, [L-ascorbic acid] ¼ 0.70 mM (rAPX) and 0.50 mM (H42E), [guaiacol] ¼ 30 mM (rAPX) and 11 mM (H42E), and [enzyme] ¼ 25 nM. Specific activities ([enzyme] ¼ 25 nM, sodium phosphate, pH 7.0, l ¼ 0.10 M, 25.0 (cid:1)C) were calculated from initial slopes of activity measurements; 1 unit of activity is defined as the amount of enzyme that oxidizes 1 lmol substrate per minute (lmolÆmin)1Æmg)1).

concentrations

Transient-state kinetics

)1Æcm)1 for rAPX [40], e397 ¼ 83 mM

)1Æcm)1 for H42E.

Bacterial fermentation of cells and purification of rAPX were carried out according to published procedures [7]. Enzyme purity was assessed by examination of the ASoret/ A280 value; in all cases an ASoret/A280 value > 1.9 for rAPX, H42A and H42E was considered pure. Enzyme purity was additionally assessed using SDS/PAGE, and the prepara- tions were judged to be homogeneous by the observation of a single band on a Coomassie Blue-stained reducing SDS/ polyacrylamide gel. Enzyme (pH 7.0, l ¼ 0.10 M, 25.0 (cid:1)C) were determined using the pyridine haemochromagen method [39]: absorption coefficients were )1Æcm)1 e403 ¼ 88 mM for H42A and e404 ¼ 95 mM

UV/visible spectroscopy

Spectra were recorded using a variable-slit Perkin-Elmer Lambda 14 UV/visible spectrometer, linked to an Exacta 486D computer, and an Epsom-LQ-1060 printer. Tempera- ture was controlled (± 0.1 (cid:1)C) using a thermally jacketed cell holder connected to a circulating water bath (Julabo U3) and a water cooler (MK Refrigeration Limited), which was operated in tandem.

Mass spectrometry

Samples were analysed using a Micromass Quattro BQ (Tandem Quadrupole) electrospray mass spectrometer. Horse heart myoglobin (Sigma) was prepared as described for rAPX below, and used to calibrate the spectrometer in the range 600–1400 m/z. Protein samples were introduced into the instrument at a flow rate of 5 lLÆmin)1. Trace salt was removed using a Centricon-10 concentrator (Amicon) and successive centrifugation and dilution with highly purified water (Elgastat). Samples ((cid:2) 2 mgÆmL)1, 20 lL) were then diluted 10-fold with a solution of 50 : 50 (v/v) acetonitrile/water containing 0.1% acetic acid.

Transient-state kinetics were performed using a SX.18 MV microvolume stopped-flow spectrophotometer (Applied Photophysics) fitted with a Neslab RTE200 circulating water bath (± 0.1 (cid:1)C). Reported values of kobs are an average of at least three measurements. All curve fitting was performed using the Grafit software package. All data were analysed using nonlinear least-squares regression analysis on an Archimedes 410–1 microcomputer using Spectra- kinetics software (Applied Photophysics). Pseudo-first- order rate constants for the formation of Compound I (k1,obs) were monitored at 403 nm (rAPX), 404 nm (H42E) and 397 nm (H42A), in single mixing mode by mixing enzyme (0.5–1.0 lM) with various concentrations of H2O2. Absorbance changes were independent of [H2O2]; observed changes in absorbance were 97–99% of the calculated values. The pH-jump method was used to examine the pH-dependence of Compound I formation, to avoid enzyme instability problems below pH 5 and above pH 8.5. Enzyme samples were prepared in water, adjusted to pH 7 with trace amounts of phosphate buffer (5 mM, pH 8.0); H2O2 solutions were made up in buffers of twice the final concentration. The buffers used were sodium phosphate in the pH range 5.5–8.5 (l ¼ 0.20 M), citrate-phosphate in the pH range 4.0–6.0 (l ¼ 0.20 M) and carbonate buffer in the range 8.0–9.0 (l ¼ 0.20 M). The pH of the solution was measured after mixing to ensure consistency. pH-dependent data were fitted to the Henderson–Hasselbach equation for a single-proton process (Eqn 4):

Steady-state measurements

k ¼

ð4Þ

A þ B (cid:9) 10pH(cid:4)pKa 1 þ 10pH(cid:4)pKa

substrates were as

)1Æcm)1 [41]; guaiacol, e470 ¼ 22.6 mM

Stock solutions of L-ascorbic acid, guaiacol, H2O2 and enzyme were prepared in sodium phosphate (l ¼ 0.10 M, pH 7.0, 25.0 (cid:1)C). Enzyme assays were performed in a 1-mL quartz cuvette: various concentrations of substrate and 25 nM enzyme were preincubated for 3 min in buffer and the reaction was initiated by the addition of H2O2 ((cid:2) 2.5 lL, (cid:2) 30 mM) to a final concentration of 0.1 mM. The wavelengths and absorption coefficients used for follows: L-ascorbic acid, various )1Æcm)1 e290 ¼ 2.8 mM [42]. Activities were determined by dividing the change in absorbance by the absorption coefficient of the substrate. Values for kcat were calculated by dividing the maximum )1Æs)1) by the micromolar concentration rate of activity (lM of enzyme; values for Km were determined by a fit of the data to the Michaelis–Menten equation using a nonlinear regression analysis program (Grafit32 version 3.09b; Erithacus Software Ltd). All reported values are the mean of three independent assays. Errors on kcat and Km are estimated to ± 5% and ± 10%, respectively. For

where A and B are the rate constants for Compound I formation at the extremes of acidic and basic pH, respect- ively, and k is the rate constant (either second-order, k1, for rAPX or limiting first-order, k¢1, for H42A/H42E) for Compound I formation. Formation of Compound I in the presence of exogenous imidazole was carried out using single-wavelength mode (397 nm), where one syringe con- tained H42A (1 lM) and the other H2O2 (0.5–35 mM) in the presence of either imidazole or 1,2-dimethylimidazole (20 mM) (relatively low concentrations of exogenous imi- dazole and a high buffer concentration were used to minimize the effect of fluctuating imidazole levels on the ionic strength and pH and to prevent binding of the imidazole to the haem). Time-dependent spectra of the various reactions were obtained by multiple-wavelength stopped-flow spectroscopy using a photodiode array detec- tor and X-SCAN software (Applied Photophysics). Spectral

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Steady-state kinetics

deconvolution was performed by global analysis and integration methods using PROKIN software numerical (Applied Photophysics).

R E S U L T S

Mass spectrometry

The specific activity of rAPX for oxidation of L-ascorbic acid (256 ± 6 UÆmg)1) is comparable to published data (411 UÆmg)1) for pAPX [8]. The specific activity of H42E (8.2 ± 0.3 UÆmg)1) was (cid:2) 30-fold lower than that of rAPX. Under conditions identical with those used for rAPX and H42E, H42A exhibited no activity with L-ascorbic acid, although residual activity was detected (9.2 ± 0.3 · 10)2 UÆmg)1) when a higher enzyme concen- tration was used ([H42A] ¼ 200 nM).

The integrity of the variant proteins was examined to ensure that post-translational modification of the protein had not occurred. Analysis of H42A and H42E (data not shown) gave average masses for the apoproteins of 27126.9 ± 0.8 Da and 27185.1 ± 0.5 Da, respectively, in good agree- ment with the calculated masses of 27126.74 Da (H42A) and 27184.88 Da (H42E).

Electronic spectra

Electronic spectra of the ferric derivatives of H42A and H42E (pH 7.0, l ¼ 0.10 M, 25.0 (cid:1)C) are shown in Fig. 2. Wavelength maxima (Table 1) for H42A and H42E were found to be slightly different from those for rAPX, but are consistent with a predominantly five-coordinate high-spin iron. In the presence of excess cyanide, spectra for H42A and H42E were consistent with the formation of a low-spin haem species, with absorption maxima [H42A–CN kmax )1Æcm)1)) ¼ 420 (102), 540, 574sh; H42E–CN (nm) (e (mM )1Æcm)1)) ¼ 420 (109), 540, 573sh] similar kmax (nm) (e (mM to those for the corresponding derivative of rAPX [rAPX– )1Æcm)1)) ¼ 419 (104), 539, 572sh]. The CN kmax (nm) (e (mM spectra of both ferric H42A and H42E were, on the other hand, unaffected by the addition of either azide or fluoride, suggesting that these (weak field) ligands do not bind to the haem under these conditions.

Steady-state data (kcat, Km and the arithmetically calcu- lated selectivity coefficient, kcat/Km) for oxidation of L-ascorbic acid and guaiacol by rAPX and H42E are shown in Table 2. (The oxidation of L-ascorbic acid by rAPX does not obey standard Michaelis kinetics [8,22,43] and, in this case, data were fitted to the Hill equation. Oxidation of guaiacol by rAPX obeys Michaelis kinetics and data were fitted to the Michaelis–Menten equation. The origin of the different concentration-dependencies for these two sub- strates is not known. Oxidation of both L-ascorbic acid and guaiacol by H42E was observed to obey Michaelis–Menten kinetics.) For both substrates, kcat values for H42E are (cid:2) 50- fold lower than for rAPX, with Km values largely unaffected (about threefold lower for H42E). The H42A variant was inactive with both L-ascorbic acid (above) and guaiacol. The dependence of the rate of substrate oxidation (lMÆs)1) vs. pH yielded a pH optimum for rAPX at (cid:2) 7 for the oxidation of both L-ascorbic acid (pH optimum 7.0) and guaiacol (pH optimum 6.9) (data not shown) (the pH optimum for L-ascorbic acid is consistent with that reported previously for pAPX [8]). For H42E, the pH optimum is shifted by (cid:2) 1 pH unit for both L-ascorbic acid (pH optimum 8.1) and guaiacol (pH optimum 8.0).

)1Æcm)1) for the ferric and Compound I derivatives of rAPX, H42A

Fig. 2. UV/visible spectra of ferric rAPX (solid line), H42A (dotted line) and H42E (dashed line). The region 450–700 nm has been multi- plied by a factor of five. Sample conditions: sodium phosphate, pH 7.0, l ¼ 0.10 M, 25.0 (cid:1)C.

Table 1. Wavelength maxima (nm) and in parentheses absorption coefficients (mM and H42E.

Derivative rAPX H42A H42E

FeIII Compound I 403(88), 506, 540sh, 636 404(59), 529, 583sh, 650 397(83), 509, (cid:2)540, 644 403(62), 534, 575sh, 645 404(95), 516, 538sh, 639 404(66), 530, 573sh, 654

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L-Ascorbate

Table 2. Values for kcat, Km and kcat/Km for the oxidation of L-ascorbic acid and guaiacol by rAPX and H42E (sodium phosphate, pH 7.0, l ¼ 0.10 M).

Guaiacol

)1Æs)1)

)1Æs)1)

Enzyme kcat/Km (mM kcat/Km (mM kcat (s)1) Km (mM) kcat (s)1) Km (mM)

Pre-steady-state kinetics

Spectra of the transient Compound I intermediates of H42A and H42E, formed by reaction of the ferric deriva- tives with 10 equivalents of H2O2, were obtained by photodiode array experiments. These preliminary experi- ments showed that Compound I formation for both variants is much slower than for rAPX, reactions being complete in less than 1000 s for H42A and 100 s for H42E, compared with less than 300 ms for rAPX under identical conditions. Wavelength maxima and absorption coefficients for the Compound I intermediate of H42A and H42E were found to be similar to those of rAPX (Table 1) and to those previously published for wild-type pAPX (kmax ¼ 404 nm [17]). For H42A and H42E, Compound I is surprisingly stable, for up to 30 s, but does not spontaneously convert into Compound II as is observed for rAPX. Instead, Compound I for both H42A and H42E slowly returns to a spectrum with a slightly red-shifted Soret band, with wavelength maxima at 405, 514, 550sh and 640 nm for H42A and 406, 516, 552sh and 638 nm for H42E.

248 ± 28 3.9 ± 0.2 rAPX H42E 0.41 ± 0.04 0.17 ± 0.09 605 22.9 66 ± 3 1.5 ± 0.1 12.3 ± 0.9 2.9 ± 0.35 5.4 0.5

M

The dependence of the observed rate constant, k1,obs (individual traces were monophasic in all cases), on the concentration of H2O2 for H42A and H42E (Fig. 3) exhibits hyperbolic behaviour. For rAPX (data not shown and [21,22]) and pAPX [17], a linear dependence on [H2O2] is observed; in this work, a second-order rate constant of )1Æs)1 was derived for reaction of rAPX (6.1 ± 0.1) · 107 with H2O2. Saturation behaviour of the kind exhibited by H42A and H42E is consistent with a mechanism involving a pre-equilibrium step which precedes Compound I forma- tion (Eqns 5 and 6):

Ka

X

ð5Þ

E þ H2O2 (cid:4)(cid:4)*)(cid:4)(cid:4)

ð6Þ

k0 1 Compound I þ H2O X (cid:4)!

M

(E ¼ H42A or H42E). This mechanism predicts a linear (first-order) dependence at low concentrations of peroxide and a zero-order dependence at high concentrations. An expression for k1,obs can be derived (Eqn 7):

ð7Þ

k1;obs ¼

k0 1 1 þ Kd=½H2O2(cid:12)

where Kd is the dissociation constant of the bound complex in Eqn (5) (Kd ¼ 1/Ka) and k¢1 is the limiting first-order rate constant at high peroxide concentrations. A fit of these data for H42A and H42E to Eqn (7) (Fig. 3) yields values for k¢1 and Kd of 4.3 ± 0.2 s)1 and 30 ± 2.0 mM, respectively (H42A) and 28 ± 1.0 s)1 and 0.09 ± 0.01 mM, respect- ively (H42E). These data pass through the origin, indicating

that the second step of the reaction is irreversible. At low concentrations of peroxide, where a linear dependence is observed, it is possible to extract an approximate value for the second-order rate constant for reaction with H2O2 [Eqn (5) where Kd is related to the microscopic second-order (ka) and first-order (kb) rate constants for this step (Kd ¼ kb/ )1Æs)1 and 1.1 ± 0.2 · ka)]: values for ka of 84 ± 6 M )1Æs)1 were obtained for H42A and H42E, respectively. 105 The mechanistic scheme implicated by the above data suggested the accumulation of a reaction intermediate, the conversion of which to product was rate-limiting at high peroxide concentrations. To examine the nature of this intermediate, photodiode array experiments were carried out for H42A (Fig. 4). Intermediate spectra were obtained from a spectrally deconvoluted model: A fi B fi C, where A corresponds to ferric H42A, B corresponds to the intermediate (tentatively assigned as the [H42A–H2O2] complex, vide infra) and C corresponds to Compound I. Wavelength maxima for the proposed intermediate were at 401 nm, 522 nm and 643 nm. The model yielded rate constants for each step: kA (A fi B) and kB (B fi C) of

Fig. 3. Dependence of k1,obs on [H2O2] concentration for the reaction of H42A (A) and H42E (B) with H2O2 (sodium phosphate, pH 7.0 l ¼ 0.10 M, 5.0 (cid:1)C, [H42A] ¼ 0.5 lM, [H42A] ¼ 0.5 lM). Data were fitted using a nonlinear least-squares fitting procedure to Eqn (7).

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mental data. For H42E, no intermediate complex could be detected under these conditions.

Fig. 5. pH-dependence of Compound I formation for (A) rAPX, (B) H42A and (C) H42E. The solid line for rAPX and H42E represents a fit of the data to the Henderson–Hasselbach equation for a single proton process (Eqn 4); data for H42A were fitted to y ¼ c. Conditions: l ¼ 0.10 M, 5.0 (cid:1)C. array spectra (sodium phosphate,

The pH-dependences of the observed rate constants for Compound I formation for rAPX, H42A and H42E enzymes are shown in Fig. 5. Nonlinear dependencies with zero intercepts on [H2O2] were observed at all pH values for H42A and H42E. The use of acetate and nitrate buffers has been avoided to prevent anionic effects previously observed in pH-dependent studies on HRP and CcP [44–46]. Compound I spectra for each enzyme were identical at all pHs (data not shown). Determination of second-order (k1) and limiting first-order (k¢1) rate constants for rAPX and H42E, respectively, revealed pH-dependent behaviour for Compound I formation in both cases (Fig. 5A,C). These data were fitted to a single-proton process (Eqn 4), and pKa values for rAPX and H42E of 4.9 ± 0.1 and 6.7 ± 0.2, respectively, were obtained. In contrast, H42A exhibits no detectable pH-dependence for the rate constant for Com- pound I formation (k¢1) from pH 5.5–8.5 (Fig. 5B).

Recovery of H42A activity

147 s)1 and 1.19 s)1, respectively ([H2O2] ¼ 35 mM), which is in approximate agreement with the limiting rate constant of 4.3 s)1 determined above. The maximum concentration of the intermediate under these conditions was 91% of the initial enzyme concentration. Under these experimental conditions and in contrast with rAPX, there was no clear isosbestic point for the conversion of ferric H42A into Compound I, indicating that there are more than two absorbing species in solution and therefore consistent with the formation of an intermediate. At 342 nm (Fig. 4, inset), the isosbestic point between ferric enzyme and the interme- diate, the absorbance–time trace shows a lag phase before Compound I formation, providing further evidence to support the proposed intermediate; the filled circles in this inset are simulated data points derived from the model and show good agreement between the simulated and experi-

Steady-state and pre-steady-state analyses of H42A indi- cated that the enzyme was essentially inactive. Rate constants for Compound I formation for H42A in the presence of either imidazole or 1,2-dimethylimidazole (20 mM) were also determined (sodium phosphate,

pH 7.0, Fig. 4. Photodiode l ¼ 0.10 M, 5.0 (cid:1)C) for the reaction between H42A (1 lM) and H2O2 (35 mM). The reaction was monitored over a time base of 2 s, and 100 spectra were recorded with a time interval of 20 ms between each scan. Experimental data were fitted to an A fi B fi C model, using PROKIN software. Deconvoluted spectra from this analysis are shown in the Soret (A) and visible (B) regions. The solid line shows the spectrum of ferric H42A; the dotted and dashed lines show the inter- mediate spectra derived from the model, and represent the spectra of the proposed [H42A–H2O2] complex and Compound I, respectively. (B) Inset: time course at 342 nm. (d) Simulated data points derived from the model calculated using the PROKIN software.

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1,2-dimethylimidazole were (cid:2) 10-fold higher than for imidazole itself and were observed to increase linearly with increasing 1,2-dimethylimidazole concentration (1–40 mM); no evidence for saturation was observed in this case and addition of 1,2-dimethylimidazole to H42A did not generate an observable low-spin derivative.

D I S C U S S I O N

To investigate the catalytic role of the conserved His42 residue in APX catalysis, two site-directed variants were prepared in which the distal histidine was replaced by alanine (H42A) and glutamic acid (H42E) residues. The results provide: (a) unambiguous evidence that His42 is critical for efficient Compound I formation in APX; (b) confirmation that titration of this residue controls the rate constant for Compound I formation and an assignment of the pKa for this group; (c) mechanistic evidence for an intermediate before Compound I formation; (d) spectro- scopic evidence on the nature of this intermediate. The detailed implications of these data are discussed below.

Examination of the electronic spectra for the two variants indicates that large-scale structural alterations have not been induced as a direct consequence of the mutations. Hence, wavelength maxima for ferric H42A and H42E are shifted slightly compared with ferric rAPX (Table 1), but are consistent with a largely five-coordinate, high-spin haem geometry. As ferric rAPX has itself been found [47] to be a mixture of five and six-coordinate high-spin haem together with some six-coordinate low-spin haem (the relative proportions of which are themselves likely to be affected by sample and storage conditions), it is probable that the slight differences in the exact wavelength maxima reflect differences in the spin state/coordination number distribu- tion for each variant compared with rAPX. In particular, we note the broad Soret band, distinct shoulder ((cid:2) 375 nm) and red-shifted CT1 band (644 nm) for H42A, all of which are consistent with increased five-coordinate character for this variant compared with both rAPX and H42E. How- ever, it is not possible to make fully quantitative assessments from these data, and more detailed spectroscopic analyses were not attempted [although resonance Raman data (unpublished work) for rAPX and H42A at neutral pH indicate that the five and six-coordinate high-spin haem represent major and minor components, respectively].

[These values are slightly different s)1

(k¢1 ¼ 4.3 ± 0.2

[H42A] ¼ 1 lM; Fig. 6). l ¼ 0.20 M, 5.0 (cid:1)C, pH 7.0, [Higher concentrations of imidazole were not experiment- ally accessible for two main reasons: (a) high concentra- tions of either imidazole led to large changes in both ionic strength and solution pH; (b) binding of imidazole to the leading to iron is observed at higher concentrations, enzyme inhibition.] The spectrum of Compound I formed under these conditions (kmax ¼ 403, 534, 575sh, and 645 nm) was identical with that obtained in the absence of exogenous imidazoles. Formation of Compound I again showed saturation kinetics (Fig. 6), and a nonlinear least-squares fit of the data to Eqn (7) in the pres- imidazole and 1,2-dimethylimidazole yields ence of k¢1 ¼ 40 ± 3 s)1 and 168 ± 30 s)1, respectively, and Kd ¼ 25 ± 5 mM and 23 ± 8 mM, respectively. Compar- ison with the corresponding data obtained under similar experimental conditions but in the absence of exogenous ligand [k¢1 ¼ 4.1 ± 0.1 s)1, Kd ¼ 30 ± 2.0 mM (sodium phosphate, pH 7.0 l ¼ 0.20 M], indicates that imidazole and 1,2-dimethylimidazole lead to (cid:2) 10-fold and (cid:2) 40-fold increases in k¢1, respectively, with the values for Kd largely from unaffected. those determined above and Kd ¼ 30 ± 2.0 mM), because the two determinations were carried out at a slightly different ionic strengths.]

[H2O2] ¼ 0.10 mM,

imidazole (data not

In parallel steady-state experiments ([H42A] ¼ 100 nM, [guaiacol] ¼ 30 mM, sodium phos- phate, pH 7.0, l ¼ 0.20 M, 25.0 (cid:1)C), the oxidative activity of H42A with guaiacol was observed to increase with increasing concentration of imidazole and to saturate at high concentration of shown). (L-Ascorbic acid activity was not examined because both L-ascorbic acid and imidazole absorb strongly between 265 and 290 nm.) Separate UV/visible experiments (data not shown) provided independent evidence for binding of imidazole to the iron (Kd ¼ 51 ± 6 mM for H42A; Kd ¼ 0.3 ± 0.1 mM for rAPX) to generate a low-spin H42A–imidazole derivative (kmax ¼ 412, 533 and 565sh nm) that inhibited activity. Guaiacol activities in the presence of

The steady-state oxidation of L-ascorbate by pAPX [8] and rAPX [43] cannot be satisfactorily fitted to a standard Michaelis–Menten treatment, and sigmoidal kinetics are observed; steady-state oxidation of guaiacol by rAPX obeys Michaelis–Menten kinetics. Although it was initially pro- posed that the origin of the sigmoidal dependence may arise from the dimeric structure of the enzyme, site-directed mutagenesis work [43] has indicated that this is unlikely to be the case. It is possible that nonspecific effects, perhaps involving radical chemistry associated with oxidation of ascorbate, or the existence of more than one substrate binding site, may be influential. A sigmoidal dependence on substrate concentration is not observed for H42E-catalysed oxidation of L-ascorbate, an observation that we are not, at present, able to fully rationalize. Replacement of the distal His42 by alanine renders the H42A enzyme essentially inactive, which is presumably directly linked to the very poor reactivity of this variant with H2O2.

Fig. 6. Dependence of k1,obs, the pseudo-first-order rate constant for Compound I formation in H42A, on H2O2 concentration in the absence and presence of exogenous imidazoles (20 mM). Conditions: sodium phosphate, l ¼ 0.20 M, 5.0 (cid:1)C, pH 7.0, [H42A] ¼ 1 lM. (n) H42A; (s) H42A + imidazole; (d) H42A + 1,2-dimethylimidazole.

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support a key catalytic role for His42 and indicate that the glutamic acid residue is able to replace the distal histidine residue such that reasonably efficient Compound I forma- tion is effected. The corresponding H42A and H42E variants in HRP have been shown to lower the rate constant for Compound I formation by six and four orders of magnitude, respectively [49–51,57], although no evidence for an intermediate was found for these variants. Large effects on the rate constant of Compound I formation have also been reported for other His42 variants of HRP [58–60] and for CcP [61,62].

M

There is now general agreement that the Compound I species formed immediately after reaction of both rAPX and pAPX with H2O2 contains a porphyrin p-cation radical and not a protein-based radical as observed in CcP [17–20]. Spectra of Compound I obtained in this work for rAPX agree with those previously published for pAPX (kmax ¼ 404 nm [17]) and those for HRP in which a porphyrin p-cation radical is known to be generated [37]. Spectra for the Compound I intermediates of H42A and H42E were similar to those of rAPX and are also consistent with a porphyrin p-cation formulation. The decay of Compound I of rAPX in the absence of substrate through the normal Compound II/ferric route is not straightforward and generates spectra that are similar to, but not identical with, authentic samples of either Compound II or ferric rAPX [48]. It has been proposed [48] that (noncatalytic) protein radical chemistry involving a Trp amino acid occurs under these conditions, leading to a permanent alteration of the haem structure that is reflected in the absorption maxima of the final (ferric-like) decay product. For H42A and H42E in the absence of substrate, Compound I does not spontaneously convert into a Compound II species as observed for rAPX, and the Compound I intermediates for both enzymes slowly generate species with a slightly red-shifted Soret band [kmax (H42A) ¼ 405 nm; kmax (H42E) ¼ 406 nm] compared with the ferric state. These maxima are analogous to those observed for the decay of rAPX (kmax ¼ 406 nm), suggesting that similar radical chemistry may occur in the variants, although this has not been examined in detail in this work. The failure to observe Compound II for the two variants under these conditions is intriguing (particularly as H42E is clearly active against both L-ascorbate and guaiacol) and has been noted previously for the H42E [49,50] and H42A and H42V [51] variants of HRP. These data suggest that the distal histidine residue stabilizes Compound II through a hydrogen-bond- ing structure involving the Ne of His42 and the ferryl oxygen. However, although hydrogen-bonding interactions of this kind have been structurally confirmed for the ferrous- oxy derivative of CcP [52], no such interaction was detected in the crystal structure of Compound I [53,54]. Instead, stabilizing hydrogen-bonding interactions from the distal arginine residue have been identified for Compound I [53,54]. Indeed, Compound II of the R38A variant of HRP was also not detected [25] and the Compounds I of the R48L, R48K [55] and R48E [56] variants of CcP are similarly unstable. Hence, it is possible that these hydrogen- bonding interactions involving Arg38 in rAPX have been simultaneously disrupted as a secondary consequence of the His42 mutations and, together with His42, may be influen- tial in defining the stability of Compound II.

The pre-steady-state data (Fig. 3) are consistent with a mechanism involving formation of an enzyme–substrate intermediate, the conversion of which into product is rate- limiting at high concentrations of peroxide. The simplest mechanism that is consistent with these data is described by Eqns (5) and (6). Although mechanisms involving conform- ational gating of the reaction at high peroxide concentra- tions [55] or before Compound I formation [63] are possible, the observation of a spectroscopically distinct intermediate for H42A provides good evidence for the proposed mechanism (but leaves this interpretation slightly more open for H42E). This enzyme intermediate has wavelength maxima at 401 nm, 522 nm and 643 nm and we assign the intermediate as arising from a transient [H42A–H2O2] species (see below). Our ability to detect this species for the first time arises from the lowering of the observed rate constant compared with rAPX as a conse- quence of the mutation, such that high concentrations of H2O2 (above the Kd) are experimentally accessible in the stopped-flow experiment. It seems unlikely that rAPX would utilize a different mechanism and we assume that detection of the intermediate is not possible in this case because the concentrations of H2O2 required to satisfy the inequality [H2O2] (cid:13) Kd would generate observed rate constants outside of the experimental stopped-flow limit. As such, only the linear part of the nonlinear k1,obs vs. [H2O2] dependence is experimentally accessible for rAPX, and O–O bond cleavage is not rate-limiting under any experimental conditions. [In fact, for all APXs [17–19] and variants of rAPX [16,21,22] examined so far, a linear dependence on [H2O2] is observed and second-order rate )1Æs)1).] Indeed, this constants are derived (k1 (cid:2) 107 intermediate has eluded detection for this very reason in other peroxidase systems and its exact structure is still not clear. For example, Baek & Van Wart [23,24] identified an intermediate, designated Compound 0 and proposed to be a hyperporphyrin (FeIII-OOH) complex, in the reaction of HRP with various peroxides under cryogenic conditions (kmax (cid:2) 330, 410 nm for HRP-H2O2). Transient interme- diates have also been detected for HRP in polyethylene glycol [26] and for the R38L/R38G and H42L variants of HRP [25,64]. Comparison of the spectra obtained in this work with theoretical calculations [29] on the hyperporph- yrin (FeIII-OOH) vs. neutral peroxide (FeIII-HOOH) struc- ture for this intermediate, together with the similarity of the spectrum of the intermediate to that of the ferric derivative, indicates that the new intermediate for H42A is probably a neutral ferric peroxide complex. This suggests that the substrate is bound initially as a neutral peroxide, consistent with the known pKa of H2O2 (pKa ¼ 11.6), which dictates that the peroxide molecule is predominantly in the proto- nated form under the conditions used in this work

The replacement of the distal histidine residue by alanine or glutamic acid clearly has a profound influence on the ability of the rAPX enzyme to react with H2O2 and is clearly demonstrated by the nonlinear dependence of the observed rate constant on peroxide concentration. A comparison of rate constants for rAPX and variant proteins is only possible by comparing the second-order rate constant derived from the linear part of Fig. 3 with the second-order rate constant for formation of Com- pound I in rAPX. These values indicate that a (cid:2) 106-fold and (cid:2) 102-fold decrease has occurred for H42A and H42E, respectively. These data provide convincing evidence to

3190 L. Lad et al. (Eur. J. Biochem. 269)

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(deprotonation of the bound hydroperoxide species is assumed to occur before O–O bond cleavage).

assistance with MS analyses. Drs Ian Ashworth and Brian Cox (Syngenta formerly Zeneca) are gratefully acknowledged for supporting this work. We are also grateful to Mr Kuldip Singh for technical assistance.

R E F E R E N C E S

1. Welinder, K.G. (1992) Superfamily of plant, fungal and bacterial peroxidases. Curr. Opin. Struct. Biol. 2, 388–393. 2. Altschul, A.M., Abrams, R. & Hogness, T.R. (1940) Cytochrome c peroxidase. J. Biol. Chem. 136, 777–794.

3. Scholes, C.P., Liu, Y., Fishel, L.A., Farnum, M.F., Mauro, J.M. & Kraut, J. (1989) Recent ENDOR and pulsed electron para- magnetic resonance studies of cytochrome c peroxidase Com- pound I and its site-directed mutants. Isr. J. Chem. 29, 85–92. 4. Erman, J.E., Vitello, L.B., Mauro, J.M. & Kraut, J. (1989) Detection of an oxy-ferryl porphyrin p cation radical in the reaction between hydrogen peroxide and a mutant yeast cyto- chrome c peroxidase. Evidence for tryptophan-191 involvement in the radical site of compound I. Biochemistry 28, 7992–7995. 5. Sivaraja, M., Goodin, D.B., Smith, M. & Hoffman, B.M. (1989) Identification by ENDOR of Trp191 as the free-radical site in cytochrome c peroxidase Compound ES. Science 245, 738–740. 6. Mauro, J.M., Fishel, L.A., Hazzard, J.T., Meyer, T.E., Tollin, G., Cusanovich, M.A. & Kraut, J. (1988) Tryptophan-191-phenyl- alanine, a proximal-side mutation in yeast cytochrome c peroxi- dase that strongly affects the kinetics of ferrocytochrome c oxidation. Biochemistry 27, 6243–6256.

7. Patterson, W.R. & Poulos, T.L. (1994) Characterization and crystallization of recombinant pea cytosolic ascorbate peroxidase. J. Biol. Chem. 269, 17020–17024.

8. Mittler, R. & Zilinskas, B.A. (1991) Purification and character- isation of pea cytosolic ascorbate peroxidase. Plant Physiol. 97, 962–968. 9. Kelly, G.J. & Latzko, E. (1979) Soluble ascorbate peroxidase. Naturewissenschaften 66, 617–618.

The pH-dependent data for Compound I formation, Fig. 5, are particularly informative. The absence of a pH- dependence for H42A unambiguously assigns this residue as being responsible for the pH-dependent reaction between rAPX and H2O2. The pKa of His42 can also be derived directly (pKa ¼ 4.9) from the rAPX data and indicates that this group must be deprotonated for efficient reaction with H2O2 (Scheme 1). By analogy with the rAPX data, the new pKa observed for H42E (pKa ¼ 6.7) can be assigned as arising from titration of the glutamic acid residue at position 42: although this is slightly higher than the pKa of the free amino acid (pKa ¼ 4.1), it is well known that protein pKa values are a sensitive function of the protein environment and that protein structure is often able to modulate thermodynamic proton-transfer events over a wide range (for example as evidenced from the range of pKa values exhibited by the distal residue in peroxidases [65]). The pKa for His42 corresponds closely to that reported previously (pKa ¼ 5.0 [17]) for pAPX, although for the pAPX enzyme no assignment of this titratable residue was possible. For peroxidases that exhibit pH-dependent kinetics of Com- pound I formation, the pKa values are in a similar range to that found in this work (for example, HRP (pKa ¼ 2.5–5.3 [66,67]), myeloperoxidase (pKa ¼ 4.0 [68,69]) and Coprinus cinereus peroxidase (pKa ¼ 5.0 [70]) and have been assigned as arising from titration of the distal residue. For peroxid- ases that exhibit pH-independent kinetics, for example, lignin [71] and manganese peroxidases [72], titration of the distal histidine presumably still influences Compound I formation, but the pKa is not experimentally accessible. For CcP, examination of the role of the distal histidine residue in Compound I formation has been complicated by specific buffer effects that alter the kinetic profile [46,62]. Where pH- dependent rate constants have been reported, the pKa values (pKa ¼ 5.4 [62], 4.0 [62]) are in a similar range to those reported here for rAPX.

10. Groden, D. & Beck, E. (1979) H2O2 destruction by ascorbate- dependent systems from chloroplasts. Biochim. Biophys. Acta 546, 426–435.

11. Dalton, D.A. (1991) Ascorbate peroxidase. In Peroxidases in Chemistry and Biology (Everse, J., Everse, K.E. & Grisham, M.B., eds), pp. 139–154. CRC Press, Boca Raton.

12. Raven, E.L. (2000) Subcellular Biochemistry: Enzyme Catalysed Electron and Radical Transfer (Holzenberg, A. & Scrutton, N.S., eds), pp. 318–350. Kluwer Academic Publishers, New York. 13. Mittler, R. & Zilinskas, B.A. (1991) Molecular cloning and nucleotide sequence analysis of a cDNA encoding pea cytosolic ascorbate peroxidase. FEBS Lett. 289, 257–259.

14. Patterson, W.R. & Poulos, T.L. (1995) Crystal structure of recombinant pea cytosolic ascorbate peroxidase. Biochemistry 34, 4331–4341.

15. Heering, H.A., Jansen, M.A.K., Thorneley, R.N.F. & Smulevich, G. (2001) Cationic asorbate peroxidase isozyme II from tea: structural insights into the heme pocket of a unique hybrid per- oxidase. Biochemistry 40, 10360–10370.

The catalytic deficiency of H42A can be partially com- pensated for by the use of exogenous imidazole [57]; we observed 1,2-dimethylimidazole to be more effective than imidazole in recovering activity against guaiacol. This is probably related to the relative affinities of the two imidazole derivatives for binding to the haem iron: binding of imidazole but not 1,2-dimethylimidazole is observed, leading to a linear dependence of the activity over the entire concentration range for 1,2-dimethylimidazole, but not for imidazole. The effect of exogenous imidazoles is also evident for the rate constant for Compound I formation for H42A, which was enhanced by (cid:2) 10-fold and (cid:2) 40-fold for imidazole and 1,2- dimethylimidazole, respectively. For both imidazoles, how- ever, the rate constant for Compound I formation and the peroxidase activity are still much lower than for rAPX itself, clearly highlighting the very specific role that the distal acid– base catalyst plays in peroxidase catalysis.

16. Pappa, H., Patterson, W.R. & Poulos, T.L. (1996) The homo- logous tryptophan critical for cytochrome c peroxidase function is not essential for ascorbate peroxidase activity. J. Biol. Inorg. Chem. 1, 61–66.

A C K N O W L E D G E M E N T S

17. Marquez, L.A., Quitoriano, M., Zilinskas, B.A. & Dunford, H.B. (1996) Kinetic and spectral properties of pea cytosolic ascorbate peroxidase. FEBS Lett. 389, 153–156.

18. Kvaratskhelia, M., Winkel, C. & Thorneley, R.N.F. (1997) Purification and characterisation of a novel class III peroxidase isozyme from tea leaves. Plant Physiol. 114, 1237–1245.

19. Kvaratskhelia, M., Winkel, C., Naldrett, M.T. & Thorneley, R.N.F. (1999) A novel high activity cationic ascorbate peroxidase This work was supported by grants from the BBSRC (Special Studentship to L. L. and project grant 91/B11469), the Royal Society (grants 18851 and 21138) and Zeneca (CASE award to L. L.). Mr J. Lamb and Professor Peter Farmer (Centre for Mechanisms of Human Toxicity, Leicester University) are gratefully acknowledged for

Catalytic mechanism of ascorbate peroxidase (Eur. J. Biochem. 269) 3191

(cid:3) FEBS 2002

42. Santimone, M. (1975) Titration study of guaiacol oxidation by HRP. Can. J. Biochem. 53, 649–657. from tea (Camellia sinensis): a class III peroxidase with unusual substrate specificity. J. Plant Physiol. 154, 273–282.

20. Patterson, W.R., Poulos, T.L. & Goodin, D.B. (1995) Identifica- in ascorbate peroxidase 43. Mandelman, D., Schwarz, F.P., Li, H. & Poulos, T.L. (1998) The role of quaternary interactions in the stability and activity of ascorbate peroxidase. Protein Sci. 7, 2089–2098. tion of a porphyrin p-cation radical Compound I. Biochemistry 34, 4342–4345.

21. Bursey, E.H. & Poulos, T.L. (2000) Two substrate binding sites in ascorbate peroxidase: the role of arginine 172. Biochemistry 39, 7374–7379. 44. Dowe, R.J. & Erman, J.E. (1982) The reaction of hydrogen- peroxide with the dimethyl ester heme derivative of cytochrome-c peroxidase. J. Biol. Chem. 257, 2403–2405.

22. Mandelman, D., Jamal, J. & Poulos, T.L. (1998) Identification of two-electron transfer sites in ascorbate peroxidase using chemical modification, enzyme kinetics, and crystallography. Biochemistry 37, 17610–17617.

45. Araiso, T. & Dunford, H.B. (1980) Horseradish peroxidase. XLI. Complex formation with nirtrate and its effect on compound I formation. Biochem. Biophys. Res. Commun. 94, 1177–1182. 46. Vitello, L.B., Huang, M. & Erman, J.E. (1990) pH-dependent spectral and kinetic properties of cytochrome c peroxidase – comparison of freshly isolated and stored enzyme. Biochemistry 29, 4283–4288. 23. Baek, H.K. & Van Wart, H.E. (1989) Elementary steps in the formation of horseradish peroxidase Compound I: direct observation of Compound 0, a new intermediate with a hyperporphyrin spectrum. Biochemistry 28, 5714–5719.

47. Nissum, M., Neri, F., Mandelman, D., Poulos, T.L. & Smulevich, G. (1998) Spectroscopic characterisation of recombinant pea cytosolic ascorbate peroxidase: similarities and differences with cytochrome c peroxidase. Biochemistry 37, 8080–8087. 24. Baek, H.K. & Van Wart, H.E. (1992) Elementary steps in the reaction of horseradish peroxidase with several peroxides: kinetics and thermodynamics of formation of Compound 0 and Com- pound I. J. Am. Chem. Soc. 114, 718–725.

25. Rodriguez-Lopez, J.N., Smith, A.T. & Thorneley, R.N.F. (1996) Role of Arg38 in horseradish peroxidase. J. Biol. Chem. 271, 4023– 4030. 48. Hiner, A.N.P., Martı´ nez, J.I., Arnao, M.B., Acosta, M., Turner, D.D., Raven, E.L. & Rodrı´ guez-Lo´ pez, J.N. (2001) Detection of a radical intermediate in the reaction of ascorbate peroxidase with hydrogen peroxide. Eur. J. Biochem. 268, 3091–3098.

26. Ozaki, S.-I., Inada, Y. & Watanabe, Y. (1998) Characterisation of polyethylene glycolated horseradish peroxidase in organic solvents: generation and stabilization of transient catalytic intermediates at low temperature. J. Am. Chem. Soc. 120, 8020– 8025.

49. Tanaka, M., Ishimori, K. & Morishima, I. (1996) The distal glu- tamic acid as an acid-base catalyst in the distal site of horseradish peroxidase. Biochem. Biophys. Res. Commun. 227, 393–399. 50. Tanaka, M., Ishimori, K., Kitagawa, T. & Morishima, I. (1997) Catalytic activities and structural properties of horseradish per- oxidase distal His42Glu or Gln mutant. Biochemistry 36, 9889– 9898.

27. Wirstam, M., Blomberg, M.R.A. & Siegbahn, P.E.M. (1999) Reaction mechanism of Compound I formation in Haem perox- idases: a density functional study. J. Am. Chem. Soc. 121, 10178– 10185. 51. Newmyer, S.L. & Ortiz de Montellano, P.R. (1995) Horseradish peroxidase His42Ala, His42Val and Phe41Ala mutants. J. Biol. Chem. 270, 19430–19438.

28. Loew, G. & Dupuis, M. (1996) Structure of a model transient peroxide intermediate of peroxidases by ab initio methods. J. Am. Chem. Soc. 118, 10584–10587. 52. Miller, M.A., Shaw, A. & Kraut, J. (1994) 2.2 A˚ structure of oxy- peroxidase as a model for the transient enzyme:peroxide complex. Nat. Struct. Biol. 1, 524–530.

53. Fulop, V., Phizackerley, R.P., Soltis, S.M., Clifton, I.J., Wakat- suki, S., Erman, J., Hajdu, J. & Edwards, S.L. (1994) Laue dif- fraction study on the structure of cytochrome c peroxidase compound I. Structure 2, 201–208.

29. Harris, D.L. & Loew, G.H. (1996) Identification of putative per- oxide intermediates of peroxidases by electronic structure and spectra calculations. J. Am. Chem. Soc. 118, 10588–10594. 30. Filizola, M. & Loew, G. (2000) Role of protein environment in horseradish peroxidase compound I formation: molecular dynamics simulations of horseradish peroxidase-HOOH complex. J. Am. Chem. Soc. 122, 18–25. 31. Poulos, T.L. & Kraut, J. (1980) The stereochemistry of peroxidase 54. Edwards, S.L., Xuong, N., Hamlin, R.C. & Kraut, J. (1987) Crystal structure of cytochrome c peroxidase Compound I. Bio- chemistry 26, 1503–1511. catalysis. J. Biol. Chem. 255, 8199–8205.

32. Finzel, B.C., Poulos, T.L. & Kraut, J. (1984) Crystal structure of cytochrome c peroxidase refined at 1.7 A˚ resolution. J. Biol. Chem. 259, 13027–13036. 33. Veitch, N.C. & Smith, A.T. (2001) Horseradish peroxidase. Adv. Inorg. Chem. 51, 107–162. 34. Smith, A.T. & Veitch, N.C. (1998) Substrate binding and catalysis 55. Vitello, L.B., Erman, J.E., Miller, M.A., Wang, J. & Kraut, J. (1993) Effect of arginine on the reaction between cytochrome c peroxidase and hydrogen peroxide. Biochemistry 32, 9807–9818. 56. Bujons, J., Dikiy, A., Ferrer, J.C., Banci, L. & Mauk, A.G. (1997) Charge reversal of a critical active-site residue of cytochrome c peroxidase: characterization of the Arg48Glu variant. FEBS Lett. 243, 72–84. in heme peroxidases. Curr. Opin. Chem. Biol. 2, 269–278. 35. Erman, J.E. (1998) Cytochrome c peroxidase: a model heme 57. Newmyer, S.L. & Ortiz de Montellano, P.R. (1996) Recue of the catalytic activity of an H42A mutant of horseradish peroxidase by exogenous imidazoles. J. Biol. Chem. 271, 14891–14896. protein. J. Biochem. Mol. Biol. 31, 307–327.

36. English, A.M. (1994) Encyclopedia of Inorganic Chemistry: Iron: Haem Proteins, Peroxidases and Catalases (King, R.B., ed.), pp. 1682–1697. Wiley, Chichester. 58. Savenkova, M.I., Newmyer, S.L. & Ortiz de Montellano, P.R. (1996) Rescue of His42Ala horseradish peroxidase by a Phe41His mutation. J. Biol. Chem. 271, 24598–24603.

59. Savenkova, M.I., Kuo, J.M. & Ortiz de Montellano, P.R. (1998) Improvement of peroxygenase activity by relocation of a catalytic histidine within the active site of horseradish peroxidase. Bio- chemistry 37, 10828–10836. 37. Dunford, H.B. (1999) Heme Peroxidases. John Wiley, Chichester. 38. Nelson, D.P. & Kiesow, L.A. (1972) Anal. Biochem. 49, 474–478. 39. Antonini, M. & Brunori E.B. (1971) Hemoglobin and Myoglobin and their Reactions with Ligands. North Holland Publishers, Amsterdam.

60. Rodriguez-Lopez, J.N., Smith, A.T. & Thorneley, R.N.F. (1996) Recombinant horseradish peroxidase isozyme C: the effect of distal haem cavity mutations (His42Leu and Arg38Leu) on com- pound I formation and substrate binding. J. Biol. Inorg. Chem. 1, 136–142. 40. Hill, A.P., Modi, S., Sutcliffe, M.J., Turner, D.D., Gilfoyle, D.J., Smith, A.T., Tam, B.M. & Lloyd, E. (1997) Chemical, spectro- scopic and structural investigation of the substrate-binding site in ascorbate peroxidase. Eur. J. Biochem. 248, 347–354. 41. Asada, K. (1984) Chloroplasts: formation of active oxygen and its 61. Erman, J.E., Vitello, L.B., Miller, M.A. & Kraut, J. (1992) Active- site mutations in cytochrome c peroxidase: a critical role for his- scavenging. Methods Enzymol. 105, 422–427.

3192 L. Lad et al. (Eur. J. Biochem. 269)

(cid:3) FEBS 2002

(Everse, J., Everse, K.E. & Grisham, M.B., eds), pp. 1–24. CRC Press, Boca Raton. tidine-52 in the rate of formation of Compound I. J. Am. Chem. Soc. 114, 6592–6593.

62. Erman, J.E., Vitello, L.B., Miller, M.A., Shaw, A., Brown, K.A. & Kraut, J. (1993) Histidine 52 is a critical residue for rapid for- mation of cytochrome c peroxidase Compound I. Biochemistry 32, 9798–9806. 68. Furtmuller, P.G., Burner, U., Jantschko, W., Regelsberger, G. & Obinger, C. (2000) Two-electron reduction and one-electron oxi- dation of organic hydroperoxides by human myleoperoxidase. FEBS Lett. 484, 139–143.

69. Marquez, L.A., Huang, J.T. & Dunford, H.B. (1994) Spectral and kinetic studies on the formation of myeloperoxidase compound I and compound II – roles of hydrogen peroxide and superoxide. Biochemistry 33, 1447–1454. 63. Rasmussen, C.B., Hiner, A.N.P., Smith, A.T. & Welinder, K.G. (1998) Effect of calcium, other ions, and pH on the reactions of barley peroxidase with hydrogen peroxide and fluoride. J. Biol. Chem. 273, 2232–2240.

ferulic 64. Rodriguez-Lopez, J.N., Lowe, D.J., Hernandez-Ruiz, J., Hiner, (2001) A.N.P., Garcia-Canovas, F. & Thorneley, R.N.F. Mechanism of reaction of hydrogen peroxide with horseradish peroxidase: identification of intermediates in the catalytic cycle. J. Am. Chem. Soc. 123, 11838–11847. 70. Abelskov, A.K., Smith, A.T., Rasmussen, C.B., Dunford, H.B. (1997) pH dependence and structural & Welinder, K.G. interpretation of the reactions of Coprinus cinereus peroxidase with hydrogen peroxide, acid and 2,2¢-azinobis (3-ethylbenzthiazoline-6-sulphonic acid). Biochemistry 36, 9453– 9463.

65. Dunford, H.B. (2001) How do enzymes work? Effect of electron circuits on transition state acid dissociation constants. J. Biol. Inorg. Chem. 6, 819–822. 71. Andrawis, A., Johnson, K.A. & Tien, M. (1988) Studies on compound-I formation of the lignin peroxidase from Phanero- chaete chrysosporium. J. Biol. Chem. 263, 1195–1198.

72. Wariishi, H., Dunford, H.B., MacDonald, I.D. & Gold, M.H. (1989) Manganese peroxidase from the lignin-degrading basidio- mycete Phanerochaete chrysosporium. J. Biol. Chem. 264, 3335– 3340. 66. Rodriguez-Lopez, J.N., George, S.J. & Thorneley, R.N.F. (1998) The structure of carbonyl horseradish peroxidase: spec- troscopic and kinetic characterisation of the carbon monoxide complexes of His42Leu and Arg38Leu mutants. J. Biol. Inorg. Chem. 3, 44–52.

67. Dunford, H.B. (1991) Horseradish peroxidase: structure and kinetic properties. In Peroxidases in Chemistry and Biology