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

Role of histidine 42 in ascorbate peroxidase
Kinetic analysis of the H42A and H42E variants
Latesh Lad
1
, Martin Mewies
1
, Jaswir Basran
2
, Nigel S. Scrutton
2
and Emma L. Raven
1
1
Department of Chemistry and
2
Department of Biochemistry, University of Leicester, UK
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°C) revealed wavelength
maxima [k
max
(nm):397,509,% 540
sh
, 644 (H42A); 404, 516,
% 538
sh

, 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% 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 [k
max
(nm) ¼ 403, 534, 575
sh
, 645] and H42E
[k
max
(nm) ¼ 404, 530, 573
sh
, 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
(K
d
) followed by conversion of this intermediate into
Compound I (k¢
1
). Values for k¢

1
and K
d
were, respectively,
4.3 ± 0.2 s
)1
and 30 ± 2.0 m
M
(H42A) and 28 ± 1.0 s
)1
and 0.09 ± 0.01 m
M
(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–H
2
O
2
] species. Rate
constants for Compound I formation for H42A were
independent of pH, but for rAPX and H42E were pH-
dependent [pK
a
¼ 4.9 ± 0.1 (rAPX) and pK
a
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

anassignmentofthepK
a
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.
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.
manganese peroxidase, lignin peroxidase) and class III
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
mechanistic, structural and spectroscopic scrutiny that it has
become the benchmark against which all other peroxidases
are measured.
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 H
2

O
2
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] that the initial product
(Compound I) of the reaction of APX with H
2
O
2
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:

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)
Eur. J. Biochem. 269, 3182–3192 (2002) Ó FEBS 2002 doi:10.1046/j.1432-1033.2002.02998.x
potassium-binding site (not present in CcP), located % 8A
˚
from the a-carbon of Trp179; the functional role of this site
(if, indeed, there is one) is not yet fully understood.
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 H

2
O
2
À!
k
1
Compound I þ H
2
O ð1Þ
Compound I þ HS À!
k
2
Compound II þ S

ð2Þ
Compound II þ HS À!
k
3
APX þ S

þ H
2
O ð3Þ
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 H
2
O – has been suggested (Scheme 1). Of
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 H
2
O
2
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 H

2
O
2
.
MATERIALS AND METHODS
Materials
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. H
2
O
2
solutions were freshly prepared
by dilution of a 30% (v/v) solution (BDH): exact concen-
trations were determined using the published absorption
coefficient (e
240
¼ 39.4
M
)1
Æcm
)1
[38]). Aqueous solutions
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).
Mutagenesis and protein purification
Site-directed mutagenesis was performed according to the
Quikchange
TM
protocol (Stratagene Ltd, Cambridge, UK).
Two complementary oligonucleotides encoding the desired
mutation were synthesized and purified (PerkinElmer). For
H42A, the primers were: 5¢-CGTTTGGCATGGGCT
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 °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).
Fig. 1. Active site of ascorbate peroxidase [14]. Hydrogen bonds

(dotted lines) are indicated.
Ó FEBS 2002 Catalytic mechanism of ascorbate peroxidase (Eur. J. Biochem. 269) 3183
sequence data were analysed using the program SeqED
(Applied Biosystems). Individual mutations were confirmed
by sequencing across the whole rAPX-coding gene.
Bacterial fermentation of cells and purification of rAPX
were carried out according to published procedures [7].
Enzyme purity was assessed by examination of the A
Soret
/
A
280
value; in all cases an A
Soret
/A
280
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 concentrations (pH 7.0,
l ¼ 0.10
M
, 25.0 °C) were determined using the pyridine
haemochromagen method [39]: absorption coefficients were
e
403
¼ 88 m
M

)1
Æcm
)1
for rAPX [40], e
397
¼ 83 m
M
)1
Æcm
)1
for H42A and e
404
¼ 95 m
M
)1
Æcm
)1
for H42E.
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 °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 (% 2mgÆmL
)1
,20lL)
were then diluted 10-fold with a solution of 50 : 50 (v/v)
acetonitrile/water containing 0.1% acetic acid.
Steady-state measurements
Stock solutions of
L
-ascorbic acid, guaiacol, H
2
O
2
and
enzyme were prepared in sodium phosphate (l ¼ 0.10
M
,
pH 7.0, 25.0 °C). Enzyme assays were performed in a 1-mL
quartz cuvette: various concentrations of substrate and
25 n
M
enzyme were preincubated for 3 min in buffer and
the reaction was initiated by the addition of H
2

O
2
(% 2.5 lL, % 30 m
M
) to a final concentration of 0.1 m
M
.
The wavelengths and absorption coefficients used for
various substrates were as follows:
L
-ascorbic acid,
e
290
¼ 2.8 m
M
)1
Æcm
)1
[41]; guaiacol, e
470
¼ 22.6 m
M
)1
Æcm
)1
[42]. Activities were determined by dividing the change in
absorbance by the absorption coefficient of the substrate.
Values for k
cat
were calculated by dividing the maximum

rate of activity (l
M
)1
Æs
)1
) by the micromolar concentration
of enzyme; values for K
m
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 k
cat
and K
m
are estimated to ± 5% and ± 10%, respectively. For
pH-dependent assays, a mixed sulfonic acid buffer system
(l ¼ 95–110 m
M
depending on the exact pH) that buffered
over the entire pH range was used; reactions were initiated
by the addition of H
2
O
2
(to 0.10 m
M
). In these cases,
[

L
-ascorbic acid] ¼ 0.70 m
M
(rAPX) and 0.50 m
M
(H42E),
[guaiacol] ¼ 30 m
M
(rAPX) and 11 m
M
(H42E), and
[enzyme] ¼ 25 n
M
. Specific activities ([enzyme] ¼ 25 n
M
,
sodium phosphate, pH 7.0, l ¼ 0.10
M
,25.0°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
)1
Æmg
)1
).
Transient-state kinetics
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 °C). Reported values of k
obs
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
(k
1,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 l
M
) with various concentrations of H
2
O
2
.
Absorbance changes were independent of [H
2
O
2
]; 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 m
M
,pH8.0);H
2
O
2
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):
k ¼
A þ B Â 10
pHÀpK
a
1 þ 10
pHÀpK
a
ð4Þ
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, k
1
,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 l
M
)andtheotherH
2
O
2
(0.5–35 m
M
)inthe
presence of either imidazole or 1,2-dimethylimidazole
(20 m
M
) (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
3184 L. Lad et al.(Eur. J. Biochem. 269) Ó FEBS 2002
deconvolution was performed by global analysis and
numerical integration methods using
PROKIN
software
(Applied Photophysics).
RESULTS
Mass spectrometry
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 °C)areshowninFig.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 k
max

(nm) (e (m
M
)1
Æcm
)1
)) ¼ 420 (102), 540, 574
sh
; H42E–CN
k
max
(nm) (e (m
M
)1
Æcm
)1
)) ¼ 420 (109), 540, 573
sh
] similar
to those for the corresponding derivative of rAPX [rAPX–
CN k
max
(nm) (e (m
M
)1
Æcm
)1
)) ¼ 419 (104), 539, 572
sh
]. The
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 kinetics
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% 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 n
M
).
Steady-state data (k
cat

, K
m
and the arithmetically calcu-
lated selectivity coefficient, k
cat
/K
m
) 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, k
cat
values for H42E are % 50-
fold lower than for rAPX, with K
m
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 (l
M
Æs
)1
)vs.
pH yielded a pH optimum for rAPX at % 7forthe
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 % 1 pH unit for both
L
-ascorbic acid (pH
optimum 8.1) and guaiacol (pH optimum 8.0).
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 °C.
Table 1. Wavelength maxima (nm) and in parentheses absorption coefficients (m
M

)1
Æcm
)1
) for the ferric and Compound I derivatives of rAPX, H42A
and H42E.
Derivative rAPX H42A H42E
Fe
III
403(88), 506, 540
sh
, 636 397(83), 509, %540, 644 404(95), 516, 538
sh
, 639
Compound I 404(59), 529, 583
sh
, 650 403(62), 534, 575
sh
, 645 404(66), 530, 573
sh
, 654
Ó FEBS 2002 Catalytic mechanism of ascorbate peroxidase (Eur. J. Biochem. 269) 3185
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 H
2
O
2
, 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 (k
max
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, 550
sh
and 640 nm for
H42A and 406, 516, 552
sh
and 638 nm for H42E.
The dependence of the observed rate constant, k
1,obs
(individual traces were monophasic in all cases), on the
concentration of H
2
O
2
for H42A and H42E (Fig. 3)
exhibits hyperbolic behaviour. For rAPX (data not shown

and [21,22]) and pAPX [17], a linear dependence on [H
2
O
2
]
is observed; in this work, a second-order rate constant of
(6.1 ± 0.1) · 10
7
M
)1
Æs
)1
was derived for reaction of rAPX
with H
2
O
2
. 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):
E þ H
2
O
2
ÀÀ*
)ÀÀ
K
a
X ð5Þ

X À!
k
0
1
Compound I þ H
2
O ð6Þ
(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 k
1,obs
can be derived (Eqn 7):
k
1;obs
¼
k
0
1
1 þ K
d
=½H
2
O
2

ð7Þ
where K
d
is the dissociation constant of the bound complex

in Eqn (5) (K
d
¼ 1/K
a
)andk¢
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 K
d
of 4.3 ± 0.2 s
)1
and 30 ± 2.0 m
M
, respectively
(H42A) and 28 ± 1.0 s
)1
and 0.09 ± 0.01 m
M
, 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 H
2
O
2

[Eqn
(5) where K
d
is related to the microscopic second-order (k
a
)
and first-order (k
b
) rate constants for this step (K
d
¼ k
b
/
k
a
)]: values for k
a
of 84 ± 6
M
)1
Æs
)1
and 1.1 ± 0.2 ·
10
5
M
)1
Æs
)1
were obtained for H42A and H42E, respectively.

Themechanisticschemeimplicatedbytheabovedata
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–H
2
O
2
]
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: k
A
(A fi B) and k
B
(B fi C) of
Fig. 3. Dependence of k
1,obs
on [H
2
O
2
] concentration for the reaction of
H42A (A) and H42E (B) with H
2

O
2
(sodium phosphate, pH 7.0
l ¼ 0.10
M
,5.0°C, [H42A] ¼ 0.5 l
M
,[H42A]¼ 0.5 l
M
). Data were
fitted using a nonlinear least-squares fitting procedure to Eqn (7).
Table 2. Values for k
cat
, K
m
and k
cat
/K
m
for the oxidation of
L
-ascorbic acid and guaiacol by rAPX and H42E (sodium phosphate, pH 7.0,
l ¼ 0.10
M
).
L
-Ascorbate Guaiacol
Enzyme k
cat
(s

)1
) K
m
(m
M
)
k
cat
/K
m
(m
M
)1
Æs
)1
) k
cat
(s
)1
) K
m
(m
M
)
k
cat
/K
m
(m
M

)1
Æs
)1
)
rAPX 248 ± 28 0.41 ± 0.04 605 66 ± 3 12.3 ± 0.9 5.4
H42E 3.9 ± 0.2 0.17 ± 0.09 22.9 1.5 ± 0.1 2.9 ± 0.35 0.5
3186 L. Lad et al.(Eur. J. Biochem. 269) Ó FEBS 2002
147 s
)1
and 1.19 s
)1
, respectively ([H
2
O
2
] ¼ 35 m
M
), 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-
mental data. For H42E, no intermediate complex could be
detected under these conditions.
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 [H
2
O
2
] 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 (k
1
)
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
datawerefittedtoasingle-protonprocess(Eqn4),andpK
a
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
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 m
M
) were also determined (sodium phosphate,
Fig. 4. Photodiode array spectra (sodium phosphate, pH 7.0,
l ¼ 0.10
M
,5.0°C) for the reaction between H42A (1 l
M
) and H
2
O
2
(35 m
M
). 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 Cmodel,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–H
2
O
2
] 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.
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°C.
Ó FEBS 2002 Catalytic mechanism of ascorbate peroxidase (Eur. J. Biochem. 269) 3187
l ¼ 0.20
M
,5.0°C, pH 7.0, [H42A] ¼ 1 l
M
;Fig.6).
[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
iron is observed at higher concentrations, leading to
enzyme inhibition.] The spectrum of Compound I formed

under these conditions (k
max
¼ 403, 534, 575
sh
,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-
ence of imidazole and 1,2-dimethylimidazole yields
k¢
1
¼ 40 ± 3 s
)1
and 168 ± 30 s
)1
, respectively, and
K
d
¼ 25 ± 5 m
M
and23±8m
M
, 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

, K
d
¼ 30 ± 2.0 m
M
(sodium
phosphate, pH 7.0 l ¼ 0.20
M
], indicates that imidazole
and 1,2-dimethylimidazole lead to % 10-fold and % 40-fold
increases in k¢
1
, respectively, with the values for K
d
largely
unaffected. [These values are slightly different from
those determined above (k¢
1
¼ 4.3 ± 0.2 s
)1
and
K
d
¼ 30 ± 2.0 m
M
), because the two determinations were
carried out at a slightly different ionic strengths.]
In parallel steady-state experiments ([H42A] ¼ 100 n
M
,
[guaiacol] ¼ 30 m

M
,[H
2
O
2
] ¼ 0.10 m
M
, sodium phos-
phate, pH 7.0, l ¼ 0.20
M
,25.0°C), the oxidative activity
of H42A with guaiacol was observed to increase with
increasing concentration of imidazole and to saturate at
high concentration of imidazole (data not 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 (K
d
¼ 51 ± 6 m
M
for H42A;
K
d
¼ 0.3 ± 0.1 m
M

for rAPX) to generate a low-spin
H42A–imidazole derivative (k
max
¼ 412, 533 and 565
sh
nm)
that inhibited activity. Guaiacol activities in the presence of
1,2-dimethylimidazole were % 10-fold higher than for
imidazole itself and were observed to increase linearly with
increasing 1,2-dimethylimidazole concentration (1–40 m
M
);
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.
DISCUSSION
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 pK
a
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 (% 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].
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 H
2
O
2
.
Fig. 6. Dependence of k
1,obs
, the pseudo-first-order rate constant for
Compound I formation in H42A, on H
2
O
2
concentration in the absence
and presence of exogenous imidazoles (20 m
M

). Conditions: sodium
phosphate, l ¼ 0.20
M
,5.0°C, pH 7.0, [H42A] ¼ 1 l
M
.(n)H42A;
(s) H42A + imidazole; (d) H42A + 1,2-dimethylimidazole.
3188 L. Lad et al.(Eur. J. Biochem. 269) Ó FEBS 2002
There is now general agreement that the Compound I
species formed immediately after reaction of both rAPX
andpAPXwithH
2
O
2
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
(k
max
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 [k
max
(H42A) ¼ 405 nm; k
max
(H42E) ¼ 406 nm] compared with the ferric state. These
maxima are analogous to those observed for the decay of
rAPX (k
max
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 N
e
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 CcPare
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 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 H
2
O
2
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 % 10
6
-fold

and % 10
2
-fold decrease has occurred for H42A and H42E,
respectively. These data provide convincing evidence to
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].
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–H

2
O
2
] 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
H
2
O
2
(above the K
d
) 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 H
2
O
2
required to satisfy the
inequality [H
2
O
2
] ) K
d
would generate observed rate
constants outside of the experimental stopped-flow limit.

As such, only the linear part of the nonlinear k
1,obs
vs.
[H
2
O
2
] 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 [H
2
O
2
] is observed and second-order rate
constants are derived (k
1
% 10
7
M
)1
Æs
)1
).]Indeed,this
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 (Fe

III
-OOH) complex, in the reaction of
HRP with various peroxides under cryogenic conditions
(k
max
% 330, 410 nm for HRP-H
2
O
2
). 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 (Fe
III
-OOH) vs. neutral peroxide (Fe
III
-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 pK
a
of H
2
O
2

(pK
a
¼ 11.6), which dictates
that the peroxide molecule is predominantly in the proto-
nated form under the conditions used in this work
Ó FEBS 2002 Catalytic mechanism of ascorbate peroxidase (Eur. J. Biochem. 269) 3189
(deprotonation of the bound hydroperoxide species is
assumed to occur before O–O bond cleavage).
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 H
2
O
2
.ThepK
a
of His42 can also be derived
directly (pK
a
¼ 4.9) from the rAPX data and indicates that
this group must be deprotonated for efficient reaction with
H
2
O
2
(Scheme 1). By analogy with the rAPX data, the new
pK
a

observed for H42E (pK
a
¼ 6.7)canbeassignedas
arising from titration of the glutamic acid residue at position
42: although this is slightly higher than the pK
a
of the free
aminoacid(pK
a
¼ 4.1), it is well known that protein pK
a
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 pK
a
values
exhibited by the distal residue in peroxidases [65]). The pK
a
for His42 corresponds closely to that reported previously
(pK
a
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 pK
a
values are in a similar range to
that found in this work (for example, HRP (pK
a

¼ 2.5–5.3
[66,67]), myeloperoxidase (pK
a
¼ 4.0 [68,69]) and Coprinus
cinereus peroxidase (pK
a
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 pK
a
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 pK
a
values
(pK
a
¼ 5.4 [62], 4.0 [62]) are in a similar range to those
reported here for rAPX.
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 % 10-fold and % 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.
ACKNOWLEDGEMENTS
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
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.
REFERENCES
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 Trp
191
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.
10. Groden, D. & Beck, E. (1979) H
2
O
2
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.
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.
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
3190 L. Lad et al.(Eur. J. Biochem. 269) Ó FEBS 2002
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-
tion of a porphyrin p-cation radical in ascorbate peroxidase
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.
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.
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.
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.
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.
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.
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.
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
catalysis. J.Biol.Chem.255, 8199–8205.
32. Finzel,B.C.,Poulos,T.L.&Kraut,J.(1984)Crystalstructureof
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
in heme peroxidases. Curr. Opin. Chem. Biol. 2, 269–278.
35. Erman, J.E. (1998) Cytochrome c peroxidase: a model heme
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.
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.
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
scavenging. Methods Enzymol. 105, 422–427.
42. Santimone, M. (1975) Titration study of guaiacol oxidation by
HRP. Can. J. Biochem. 53, 649–657.
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.
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.
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.
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.
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.
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.
51. Newmyer, S.L. & Ortiz de Montellano, P.R. (1995) Horseradish
peroxidase His42Ala, His42Val and Phe41Ala mutants. J. Biol.
Chem. 270, 19430–19438.
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.
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.
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.
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.
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.
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.
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-
Ó FEBS 2002 Catalytic mechanism of ascorbate peroxidase (Eur. J. Biochem. 269) 3191
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.

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.
64. Rodriguez-Lopez, J.N., Lowe, D.J., Hernandez-Ruiz, J., Hiner,
A.N.P., Garcia-Canovas, F. & Thorneley, R.N.F. (2001)
Mechanism of reaction of hydrogen peroxide with horseradish
peroxidase: identification of intermediates in the catalytic cycle.
J.Am.Chem.Soc.123, 11838–11847.
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.
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
(Everse,J.,Everse,K.E.&Grisham,M.B.,eds),pp.1–24.CRC
Press, Boca Raton.
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.
70. Abelskov,A.K.,Smith,A.T.,Rasmussen,C.B.,Dunford,H.B.

& Welinder, K.G. (1997) pH dependence and structural
interpretation of the reactions of Coprinus cinereus peroxidase
with hydrogen peroxide, ferulic acid and 2,2¢-azinobis
(3-ethylbenzthiazoline-6-sulphonic acid). Biochemistry 36, 9453–
9463.
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.
3192 L. Lad et al.(Eur. J. Biochem. 269) Ó FEBS 2002