Within the scope of the present survey, we will limit the presentation of peroxidase structures and activities to some classical examples which have been extensively studied during recent decades:
a plant peroxidase (horseradish peroxidase), a yeast peroxidase (cytochromecperoxidase), three fungal peroxidases (chloroperoxidase, ligninase, and manganese peroxidase), and two human peroxidases (myeloperoxidase and thyroid peroxidase).
8.11.2.1 Horseradish Peroxidase (HRP)
A large number of publications on peroxidases have been devoted to horseradish peroxidase (HRP), and its properties in different oxidation states have been well studied.4–6Despite several efforts, a high-resolution X-ray structure of HRP was not obtained until 1997.7 Horseradish peroxidase contains at least seven major isoenzymes, but the majority of the physico-chemical data have been obtained with the mixture of isoenzymes B and C (C being the major component in highly purified samples). The X-ray structure was solved for HRP-C, which was expressed inE.
coli with a gene containing 309 amino-acid residues, only 306 of which are visible in the crystal structure. The prosthetic group of HRP consists of a ferric protoporphyrin IX with a histidine as proximal ligand (His-170) having an iron–nitrogen distance of 2.3 A˚ (Scheme 1). The proximal histidine (His-170) is separated from the Ca2þsite only by a threonine (Thr-171). In the active site are key conserved catalytic residues (arginine-38 with its guanidium acting as proton donor, and histidine-42 acting as base) for the activation of hydrogen peroxide; they are found in positions very similar to those in other heme-peroxidases with a pentacoordinated iron(III) center (see Figure 1for a view of the active site).
N HN
His-170 FeIII O H O
H H2N C
NH2 NH Arg-38 His-42
Scheme 1
The closest water molecule in the distal pocket is 3.2 A˚ away from the metal position. The structure shows the proximity of asparagine-70 to the distal histidine-42 and suggests its role in maintaining the basicity of His-42. That fact has been confirmed with a Val70Asn mutant that has a lower catalytic activity compared to the wild enzyme.8,9
A crystal structure of the peroxidase–substrate complex has been determined at 2.0 A˚ resolution and demonstrated the existence of an aromatic bonding pocket.10This hydrophobic distal pocket, containing a benzhydroxamic acid, is created by several phenylalanine residues (Phe-68, Phe-142, Phe-143, and Phe-179) and is close to His-42 and Arg-38. The shape of the distal cavity has been
investigated by proton NMR using the wild HRP and a mutant His42Ala. In this latter, modified enzyme, only small modifications of the distal pocket have been observed.11
The oxidation of HRP by hydrogen peroxide generates a reactive intermediate (called Com- pound I (Cpd I)) containing two redox equivalents (above the resting state of the native enzyme);
one-electron reduction of this first high-valent intermediate generates compound II (Cpd II), Compounds I and II have a Soret band at 400 (broad) and 420 nm, respectively. Cpd I consists of an iron(IV)–oxo species with a radical cation on the porphyrin ring (Scheme 2). The formation of Cpd I has been monitored by stop-flow UV-visible spectroscopy and is a chemically controlled reaction, with a pre-equilibrium of the enzyme with a neutral molecule of H2O2within the active site12 which generates a ferric hydroperoxo intermediate (called compound 0) with a hyperpor- phyrin spectrum (for calculations on this intermediate, see refs. 13 and 14). A recent study performed on wild HRP and three mutants (His42Leu, Arg38Leu, and Arg38Gly) by rapid- scan stopped-flow and EPR has shown the formation of an iron–H2O2intermediate, which serves as precursor of the FeIII–OOH intermediate (Cpd 0).15 At neutral pH values, His-42 is not protonated and acts as a base in the first deprotonation of hydrogen peroxide to generate FeIII–OOH.
Compounds I and II have been characterized by X-ray absorption and Raman spectroscopy.
Both methods demonstrated the presence of an iron(IV) oxo entity in both Cpds I and Cpd II, with a short FeẳO bond of 1.6 A˚ consistent with a ferryl structure.16 Although it has been suggested that the FeO bond distance of Cpd II was longer: 1.9 A˚ at pH 7, and 1.7 at pH 10,17 resonance Raman studies showed that Cpds I and II have FeẳO vibrations of comparable frequency, at 737 cm1and 776 cm1, respectively,18 consistent with the ferryl description.
The additional oxidizing equivalent in Cpd I is stored on the porphyrin ligand, as a-radical cation. This radical cation has a predominant2A2ucharacter, indicative of an electron abstraction from the a2u orbital of the porphyrin ligand,19 and is ferromagnetically coupled with the spin Sẳ1 of the ferryl state, as evidenced by EPR and Moăssbauer studies.20 With excess H2O2, an inactive intermediate compound III is formed, corresponding to an iron(III)–superoxo state and is equivalent to the addition of dioxygen to the ferrous state of HRP.22 The redox potentials (E00 values) of Cpds I and II are very similar and have been determined to be 0.88 V and 0.90 V, respectively for the couples Cpd I/Cpd II and Cpd II/ferric state.21Electrochemical data have been obtained on HRP by cyclic voltammetry, and provide new insights on the conversion of the catalytically active form to the inactive oxy-form (a superoxide anion bounded to an iron(III) center); these data are useful in biosensor applications for HRP.23
Figure 1 View of the active site of HRP with selected amino acids (for the complete X-ray structure, see ref.10).
His-170 FeIII
His-170 FeIV
His-170 FeIV
O AH + H+
+
A + H+
Compound I
Compound II Resting state
O H2O2 H2O
AH2
AH
Scheme 2
The high-valent iron–oxo species of the reactive intermediates of HRP are not directly access- ible for many different substrates. Early studies on the alkylation of the -meso position of the heme-group by alkyl hydrazines have indicated that substrates approach the active site of HRP from a particular edge of the prosthetic group.24 Substrate binding data have been obtained by proton NMR spectroscopy and molecular dynamics.25–27 The X-ray structure of the binary complex HRP–ferulic acid provided accurate data on the binding of the ferulic substrate (a naturally occurring phenol in plant-cell walls) near the distal Arg-38 via a hydrogen bond.10 This work also suggests that the water molecule produced from hydrogen peroxide after the generation of Cpd I remains in the active site, interacting via hydrogen bonds with ferulic acid, His-42, and Pro-139, and is liberated only after the oxidation of the phenolic substrate. This binding site of HRP must be considered as an open pocket that can accommodate a large range of substrates, explaining why this enzyme is such an efficient catalyst in the H2O2oxidation of many different electron-donating molecules, so long as their redox potentials are compatible with those of Cpds I and II. For example, HRP has been used in the modeling of extra-hepatic oxidations (not oxygenation) of exogens (including many different drugs).29
Studies on the mechanism of the O-demethylation of aromatic substrates catalyzed by HRP have shown that the methyl group is removed as methanol (and not as formaldehyde, as in cytochrome P450-mediated oxidations).30–32 This work has been useful in understanding the mechanism of lignin oxidation by ligninase, as some authors considered ligninase as an oxygen- ase, not as a peroxidase.
N-dealkylations are easily catalyzed by HRP, and its mechanism—whether or not an alpha- hydroxylamine is formed, as in P450-catalyzed dealkylations—has been controversial for many years.1,32Accurate kinetic isotope studies are in favor of a rate-determining electron-transfer step, followed by an H-atom transfer from the aminium radical-cation to the oxygen atom of the FeIVẳO species of Cpd II.33 The formation of the aminium radical-cation by HRP–Cpd I has been confirmed with the use of N-cyclopropylamine derivatives; the cyclopropyl ring of the aminium radical-cation undergoes a ring-opening rearrangement to generate a radical cation, with the positive charge on the nitrogen iminium and a radical on the terminal methylene group.34 HRP is unable to oxidize styrene, unlike chloroperoxidase, which is able to catalyze the oxidation of this olefin to a mixture of the corresponding epoxide and phenylacetaldehyde.35 How- ever, in the co-oxidation of styrene and 4-methylphenol, catalyzed by HRP, the epoxide results from the reaction with a hydroperoxide derived from the addition of dioxygen to a phenolic radical intermediate.36But a true peroxygenase activity can be obtained in the case of styrene with HRP mutants (His-42-Ala, His-42-Val, or Phe-41-Ala), indicating that substrate accessibility to the ferryl species is increased in these mutants.37,38 The asymmetric oxidation of sulfides to sulfoxides, catalyzed by HRP, has been described by Colonna et al.,39–41 with values of the enantiomeric excesses (ee) ranging from 30 to 68%. This reaction probably involves two one- electron oxidations rather than an oxygen-atom transfer from the high-valent iron–oxo intermedi- ates.42 The HRP-catalyzed asymmetric sulfoxidation can be improved by changing the reaction medium from aqueous solution to nearly (99.7%) anhydrous organic solvents (methanol or
isopropanol). The higher substrate solubility and enzyme access largely compensate for the reduced intrinsic activity of HRP in alcohols.43 However, the ee values are greatly enhanced when the sulfoxidation of thioanisole is performed in the presence of benzohydroxamic acid, a molecule forming a strong complex with HRP.44
Polyethylene-glycolated HRP has a reduced catalytic activity compared with the native enzyme, but this modification facilitates the study of the transient high-valent iron–oxo intermediates (the lifetime of Cpd I can be increased up to one hour at20C in chlorobenzene).45
The viability of a sol–gel-encapsulated HRP has been demonstrated.46The sonication of the sol during the initial hydrolysis and condensation steps allows the incorporation of fragile biomol- ecules into the sol–gel matrix. The diffusion of substrates within the sol–gel matrix limits the catalytic activity of the encapsulated enzyme.
Novel peroxidases can be generated by screening de novo heme-proteins derived from a designed combinatorial library.47Turnover numbers as high as 17,000 min1have been obtained.
8.11.2.2 Chloroperoxidase (CPO)
Among all fungal peroxidases, chloroperoxidase (CPO) is one of the few enzymes that are able to catalyze the oxidative chlorination of substrates using H2O2 and Claccording to Equation (2) (myeloperoxidase MPO is another example):48
AHỵH2O2ỵClỵHỵ CPO!AClỵ2H2O ð2ị This halogen-atom incorporation is catalyzed by CPO with substrates containing an activated carbon–hydrogen bond, such as -diketones (chlorodimedone is a classical substrate for CPO activity assays). This heme-enzyme can be easily obtained and purified from the fungusCaldar- iomyces fumago. The big difference between CPO and HRP is the presence of a cysteine residue as proximal ligand (Cys-29 in CPO; seeFigure 2).49,50
Furthermore, there is a manganese(II) ion bound to a heme-propionate and also surrounded by His-105, Glu-104, and Ser-108; this manganese ion might be the binding site for the chloride ion that has to be oxidized.51 The role of the manganese ion in CPO might be similar to that of calcium in myeloperoxidase, which is located not too far from the heme (seeSection 8.11.2.7for details on myeloperoxidase).52
The addition of hydrogen peroxide to the ferric state of the enzyme generates CPO–Cpd I, the only detectable intermediate having an FeIVẳO bond with a Raman stretching band at
Figure 2 View of the active site of CPO with selected amino acids (for the complete X-ray structure, see ref.51).
790 cm1.53,54 The radical-cation of Cpd I might be delocalized on the macrocycle or on the axial ligand (see ref. 55 for a proposal on the role of the S ligand in a high-valent cysteinato-heme- enzyme). The addition of chloride to Cpd I might generate an FeIII–OCl entity, which in turn may effect the chlorination of substrates.56 An alternative mechanism is the formation of free HOCl,48 since the same mixture of chloroanisole isomers was observed in the chlorination of anisole by free HOCl or by a CPO-catalyzed reaction. A third possible mechanism involves the manganese site of CPO as the binding site of Cl. Because of the short distance between this Mn site and the heme, the bound chloride ion could be oxidized to Clþ, which could attack the substrate present at the active site. Such a hypothesis would explain the absence of the formation of free HOCl during the CPO catalytic cycle, without invoking the formation of an iron(III) hypochlorito entity.
Chloroperoxidase is rendered inactive by the formation of an N-phenyl-heme in the oxidation of phenylhydrazine, or an N-alkyl-heme in the oxidation of terminal olefins.57,58 These data suggest that various substrates have relatively easy access to the active site of CPO. In fact, this enzyme is able to catalyze the oxidation of a large range of substrates (tertiary amines, olefins, heterocycles, etc.). The mechanism of theN-demethylation of tertiary amines, catalyzed by CPO, probably involves an electron abstraction from the nitrogen atom, followed by a proton elimin- ation of the methyl group (a primary isotope effect has been observed with akH/kD valueẳ2.5).
An alternative mechanism is the abstraction of an H atom from the methyl group, as observed for the hydroxylation reaction catalyzed by cytochrome P-450.48 Chloroperoxidase is also able to catalyze the epoxidation of different olefins (styrene, propylene, allyl chloride).45 The oxygen atom of the epoxide arises from the primary oxidant, as evidenced by the use of H218
O2.35It has been demonstrated that CPO-catalyzed epoxidations of cis-disubstituted olefins are highly enan- tioselective, with enantiomeric excesses (ee values) ranging from 33% to 85%.60 To avoid the irreversible degradation of CPO in the presence of H2O2, the concentration of the peroxide must be maintained as low as possible during the course of epoxidation. In these conditions, it has been possible to obtain 4,200 catalytic cycles with an ee value of 94% in the epoxidation of methylallyl propionate.61This CPO-catalyzed asymmetric epoxidation has been used in the synthesis of (R)- mevalonolactone,62 and also with!-bromo-2-methyl-1-alkenes.63
CPO is also able to catalyze the hydroxylation of allylic, benzylic, propargylic, and cis-cyclo- propylmethanol derivatives with ee values ranging from 60% to 95%.64–66 The CPO-catalyzed insertion of the oxygen atom into the CH bonds in these hydroxylations might involve a very short-lived, carbon-centered radical intermediate, as checked with a hypersensitive radical probe (the lifetime of such a radical intermediate must be less than 3 ps).67 CPO is also effective in converting oximes to halonitro compounds and ketones in a single step, in the presence of KCl or KBr and hydrogen peroxide.68No enantioselectivity has been observed during the formation of the nitro compounds, but the addition of co-solvents like dioxane or acetone increased the yield of the corresponding ketones.
The other useful reaction catalyzed by CPO is the enantioselective oxidation of sulfides to chiral sulfoxides with ee values of up to 90–95%.40,69,70Chiral sulfoxides are important synthons in the stereoselective synthesis of many natural products. Based on data obtained with a series of para-substituted thioanisoles, the proposed mechanism of the CPO-catalyzed S oxygenation involves an oxygen-atom transfer from compound I to the substrate, instead of a one-electron oxidation of the sulfide followed by the addition of H2O2on the intermediate S radical-cation, as is proposed for sulfide oxidations catalyzed by HRP.71
Two novel heme-containing peroxidases with CPO activity have been discovered in marine worms: Notomastus lobatus chloroperoxidase (NCPO), catalyzing the halogenation of phenols;
andAmphitrite ornata(DHP), performing the dehalogenation of halophenols to quinones.72Both enzymes contain a histidine residue as proximal ligand, and both are stable in their oxyferrous states (similarly to the Cpd III of HRP).73
8.11.2.3 Ligninase (LiP)
The possible development of the enzymatic degradation of lignin from wood pulp in the cellulose industry has stimulated many studies, since about 1980, on ligninolytic fungi. In particular, one fungus has focused the attention of several research groups in universities and industries:Phanero- chaete chrysosporium, which is able to degrade lignin by producing two extracellular peroxidases:
ligninase (LiP) and manganese peroxidase (MnP). Both enzymes are able to oxidize lignin (the
ultimate degradation product of lignin being CO2) via the formation of radical-cation intermedi- ates on the substituted phenyl rings of this natural polymer resulting from the polymerization of p-coumarylic, coniferyl, or sinapylic alcohols (all these cinnamic alcohols are derived from phenylalanine).74,75 The main target in lignin oxidation is the oxidation of the phenylpropanoid units, which constitute nearly 50% of the different linkages between the subunits of lignin. In particular, the oxidative cleavage of the CC bond of these arylglycerol--arylether motifs is the key step in both the enzymatic and the chemical degradation of lignin. The CC bond cleavage is triggered by the initial formation of radical-cation intermediates on the methoxylated aromatic rings constitutive of lignin.
Ligninase was independently purified by the groups of both Kirk and Gold in 1983.76,77Ligninase extracted from the extracellular fluid of ligninolytic cultures ofPhanerochaete chrysosporiumi s a mixture of six isozymes, the major one (H8) having been used for most of the physico-chemical studies. LiP has a molecular weight of 42,000 Da, and this glycoprotein contains one iron–
protoporphyrin IX entity as prosthetic group. Early studies on LiP, based on18O incorporation into oxidation products when performed under18O-labeled molecular oxygen, led to the classifi- cation of ligninase as an oxygenase.78 However, further studies indicated that 18O from labeled water was incorporated within the degradation products, as expected for a peroxidase-mediated oxidation (the initial radical-cation generated on methoxylated aromatic rings is highly electro- philic and reacts with water).79
The resting state of the enzyme is a high-spin iron(III) porphyrin with a histidine as proximal residue.80No water molecule is present as sixth ligand at room temperature. The X-ray structure of LiP confirms the presence of a proximal histidine (His-175), the conserved distal amino acids (His-47 and Arg-43), and a large pocket on the heme-edge.81 The structure at 1.7 A˚ resolution reveals a long Fe–N(His) bond compared to regular peroxidases, suggesting that this weak Fe–N bond reduces the electron density on the heme itself and consequently contributes to the higher redox potential of LiP.82The overall folding of ligninase resembles that of cytochromecperoxi- dase.
As for other peroxidases, a high-valent iron–oxo entity associated with a porphyrin-radical- cation, Cpd I, is generated by the addition of hydrogen peroxide (seeScheme 3for the catalytic cycle of LiP). This green compound (Soret band at 408 nm) has UV-visible and EPR characteristics similar to that of HRP–Cpd I.83
IV
FeIV N(His) O LiP
Fe O
O
N(His) Fe
N(His) O
III
(VA)
(VA)
AH2 AH2
AH2 H2O
H2O2
III N(His)
Fe
A (VA.+)
(VA.+)
O2.–
Inactivation Excess of H2O2
AH.
AH.
.+
Resting state
.
LiP II
LiP I LiP III
Scheme 3
The addition of 0.5 equivalent of a two-electron reducing agent like veratryl alcohol generates LiP–Cpd II (with a Soret band at 420 nm), which corresponds to the iron(IV)–oxo without oxidation of the porphyrin ligand. Site-directed mutations have demonstrated a key role for Trp-171 in veratryl alcohol oxidation, this amino acid being probably the location of an inter- mediate radical cation in the oxidation cascade.