Engineering a Bacterial DyP-Type Peroxidase for Enhanced Oxidation

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Engineering a Bacterial DyP-type Peroxidase for Enhanced Oxidation of Lignin-Related Phenolics at Alkaline pH Vania Brissos, Diogo Tavares, Ana Catarina Sousa, Maria Paula Robalo, and Ligia O. Martins ACS Catal., Just Accepted Manuscript • DOI: 10.1021/acscatal.6b03331 • Publication Date (Web): 07 Apr 2017 Downloaded from http://pubs.acs.org on April 7, 2017

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 Enzyme yields  Stability

 Activity DMP  pH optimum  Resistance to H2O2

A142V H125Y

2nd gen

3rd gen

 Activity ABTS  Redox potential

E188K 1st

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Epistatic interaction

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Engineering a Bacterial DyP-type Peroxidase for Enhanced Oxidation of Lignin-Related Phenolics at Alkaline pH

Vânia Brissos1, Diogo Tavares1, Ana Catarina Sousa2,3, Maria Paula Robalo2,3, Lígia O. Martins1

1

Instituto de Tecnologia Química e Biológica António Xavier, Universidade Nova de Lisboa, Av da República, 2780-157 Oeiras, Portugal 2 Área Departamental de Engenharia Química, ISEL - Instituto Superior de Engenharia de Lisboa, Instituto Politécnico de Lisboa, R. Conselheiro Emídio Navarro, 1, 1959-007 Lisboa, Portugal 3 Centro de Química Estrutural, Complexo I; Instituto Superior Técnico, Universidade de Lisboa, Av. Rovisco Pais, 1049-001 Lisboa, Portugal

Address correspondence: Lígia O. Martins, Instituto de Tecnologia Química e Biológica António Xavier, Universidade Nova de Lisboa, Av. da República, 2780-15 Oeiras, Portugal, E-mail: [email protected]

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ABSTRACT Dye-decolorizing peroxidases (DyPs) are a family of microbial heme-containing peroxidases that show important properties for lignocellulose biorefineries due to their ability to oxidize lignin-related compounds. Directed evolution was used to improve the efficiency of the bacterial PpDyP from Pseudomonas putida MET94 for phenolic compounds. Three rounds of random mutagenesis by error-prone PCR of the ppDyP-gene followed by high-throughput screening allow identifying 6E10 variant showing a 100-fold enhanced catalytic efficiency (kcat/Km) for 2,6-dimethoxyphenol (DMP), similar to those exhibited by fungal lignin peroxidases (∼105 M-1s-1). The evolved variant showed additional improved efficiency for a number of syringyl-type phenolics, guaiacol, aromatic amines, Kraft lignin and the lignin phenolic model dimer, guaiacylglycerol-β-guaiacyl-ether. Importantly, variant 6E10 displayed optimal pH at 8.5, an upshift of 4 units as compared to the wild-type, showed resistance to hydrogen peroxide inactivation, and was produced at 2-fold higher yields. The acquired mutations in the course of the evolution affected three amino acid residues (E188K, A142V and H125Y) situated at the surface of the enzyme, in the second shell of the heme cavity. Biochemical analysis of hit variants from the laboratory evolution, and single variants constructed using site-directed mutagenesis, unveiled the critical role of acquired mutations from the catalytic, stability and structural viewpoints. We show that epistasis between A142V and E188K mutations is crucial to determine substrate specificity of 6E10. Evidence suggests that ABTS and DMP oxidation occurs at the heme access channel. Details of the catalytic cycle of 6E10 were elucidated through transient kinetics providing evidences for the formation of a reversible enzyme-hydrogen peroxide complex (Compound 0) barely detected in the majority of heme peroxidases studied to date.

Key-words: directed evolution, dye-decolorizing peroxidases, ligninolytic enzymes, enzyme specificity, epistasis, Pseudomonas putida MET 94

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INTRODUCTION Lignocellulose biorefineries are promising alternative sources of renewable bulk and fine chemicals, materials, energy and fuels for sustainable development. Lignin provides strength, resistance to plants and a protective matrix surrounding the cellulose micro-fibrils of plant cell walls and therefore its removal facilitates the subsequent use of plant polysaccharides for biotechnological applications. In order to be economically feasible, future biorefineries need to overcome the recalcitrance of lignocellulose to degradation.1 This is largely determined by the molecular architecture of lignin that relies on phenylpropanoid units linked by a variety of ether and carbon-carbon bonds forming amorphous and insoluble three dimensional networks. Lignin itself is the most abundant aromatic polymer in nature and a key renewable source of high value chemicals and fuel precursors but is currently considered as bio-waste. Recent research has focused on its degradation and conversion using physico-chemical approaches including thermochemical and catalytic routes.1,2 However, at present, these strategies are not cost-effective and novel (bio)catalytic, solvent-free green processes need urgently to be optimized and implemented. Biocatalysis offers an environmentally friendly tool for lignin degradation, potentially holding the key for its successful valorisation. In nature, fungal laccases and peroxidases with high redox potential, lignin (LiP), manganese (MnP) and versatile (VP) peroxidases, are well recognized for playing a critical role in lignin depolymerization.3-6 In comparison, ligninolytic bacterial systems are far less understood7-10 but bacteria that are easy to culture, grow fast and for which molecular biology tools are well established, can be a valuable asset for exploring enzymatic strategies for lignocellulose biomass deconstruction and valorization. Dye-decolorizing peroxidases (DyPs) are a newly discovered family of bacterial and fungal heme peroxidases that show attractive catalytic properties for biotechnological purposes.11-13 These enzymes show activity for a wide range of substrates, from high redox synthetic dyes and aromatic sulphides to iron and manganese ions, phenolic and nonphenolic lignin units, using hydrogen peroxide as electron acceptor.14-18 Their ability to oxidize lignin-related compounds together with their abundancy in genomes of lignin degrading basidiomycetes (white-rot fungi)6,19,20 and widespread presence in fungal transcriptomes inhabiting forest soils21 support an active contribution of DyPs to lignin biodegradation enzymatic systems. DyPs have been classified in four phylogenetically distinct subfamilies based on their primary structure, with A-C subfamilies from bacterial and D subfamily from fungal origins, respectively.11 DyPs show no sequence homology to “classical” heme peroxidases, presenting divergence in the tertiary structure and catalytic residues in the heme pocket.11-13 They display a ferredoxin-like fold distinct from the motifs found in plant peroxidases with an αhelical structure, and have a strictly conserved aspartic acid (instead of the distal catalytic histidine), a residue proposed to act as the acid-base catalyst that promotes the heterolytic cleavage of hydrogen peroxide to water.22,23 A distal arginine is thought to be essential for proper coordination of the hydrogen peroxide molecule at the heme site and stabilization of the intermediate Compound I (Cpd I). Interestingly, in DyP members from class B, the distal aspartate seems not essential to the peroxidase activity, and instead the distal arginine 3


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emerges as the key acid-base catalyst.22,24 The reaction of Cpd I with one equivalent electron from a reducing substrate yields Compound II (Cpd II), which undergoes a second reaction by an equivalent electron to restore the resting state peroxidase.23,25 DyPs similarly to LiP or VP peroxidases, withdraw electrons from bulky substrates using long-range electron transfer routes from surface-exposed tryptophanyl or tyrosyl radicals to the heme.26,27 However, in contrast with ligninolytic fungal peroxidases, DyPs display a high number of exposed electron-rich aromatic side chain residues, challenging the unambiguous identification of radical-based catalytic sites.27,28 Additionally they possess, a rather flexible heme cavity providing alternative binding sites to small aromatic substrates.29,30 In a previous work, Pseudomonas putida MET94 was selected for its superior ability to decolorize a diverse array of high-redox synthetic dyes among a collection of bacterial strains.31 In order to characterize the dye degradation mechanisms of P. putida MET 94 the functional and structural characterization of a FMN-dependent azoreductase, PpAzoR, was first provided.31,32 PpAzoR was engineered towards improved thermostability33 and a wholecell two-step enzymatic system (with CotA-laccase) was optimized for the decolorization as well as detoxification of model wastewater dye-containing baths.34 Next, the biochemical fingerprints of PpDyP, a dye decolorising peroxidase from Class B, were characterized. 24,3537 The kinetic data showed PpDyP capability to oxidize with high efficiency, not only anthraquinonic and azo dyes, but also lignin-related compounds, such as the phenolics guaiacol, syringaldehyde or acetosyringone, the non-phenolic veratryl alcohol and also the metals manganese and ferrous ions.38 In spite of the broad substrate range exhibited, PpDyP activity for phenolics (kcat/Km ∼ 103 M-1 s-1) is far from that observed in fungal ligninolytic peroxidases (kcat/Km ∼ 105 M-1 s-1).39,40 Therefore, in the present work we set-up, for the first time, a directed evolution approach for improving a DyP enzyme for the oxidation of phenolics aiming to expand the biotechnological applications of the enzyme. Directed evolution constitutes the fastest and most effective methodology to tailor biocatalysts and was successfully used for enhancing the catalytic and stability properties of heme peroxidases, including LiPs and VPs, using yeast eukaryotic machinery.41-47 Herein the protocol for the production and screening of recombinant PpDyP and variants in Escherichia coli is discussed, together with the properties of an evolved DyP that exhibit (i) enhanced efficiency for guaiacyl and syringyl lignin-related phenolics and aromatic amines at alkaline pH, (ii) improved stability against inactivation by hydrogen peroxide and (iii) augmented production yields. A comprehensive biochemical analysis of selected variants from the in vitro evolution and variants constructed using site-directed mutagenesis guided the discussion of the structure-function relationships that correlated the enhanced properties of the 6E10 hit variant. MATERIALS AND METHODS Bacterial strains, plasmids and media. E. coli strain DH5α (Novagen) was used for routine cloning procedures, propagation and amplification of plasmid constructs. E. coli Tuner (DE3, Novagen), KRX (Promega) and BL21 star (DE3, Novagen) strains were used to express the 4


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ppDyP gene previously cloned in pET21-a (+) plasmid (Novagen) (pRC-1 plasmid38) or its evolved variants. In Tuner and BL21 star strains the target genes are under the control of T7 promoter and its expression is induced by isopropyl β-D-1-thiogalactopyranoside (IPTG), whereas in the KRX strain, the gene expression is under the control of rhaPBAD promoter, induced by rhamnose. Luria-Bertani medium (LB) or Terrific Broth medium (TB) were used for the growth of E. coli strains, supplemented with 100 µg·mL-1 ampicillin. ppDyP overexpression in different recombinant strains. E. coli Tuner (DE3, Novagen), KRX (Promega) and BL21 star (DE3, Novagen) strains were transformed with plasmid pRC138, containing the ppDyP gene. Recombinant strains were cultivated in 100 mL LB medium in 500 mL-Erlenmeyer flasks and cells were disrupted as previously described.38 cell crude extracts were used for enzymatic activity measurements. The protein concentration was determined using the Bradford assay with bovine serum albumin (BSA) as standard. SDSPAGE electrophoresis was used to visualize protein overproduction in cell crude extracts. The enzymatic activity of PpDyP was measured by following at the maximum absorption wavelengths the products of oxidation of 1 mM of either 2,2’-azino bis (3ethylbenzthiazoline-6-sulfonic acid) (ABTS) (ε420nm = 36,000 M-1 cm-1), guaiacol (Ɛ470nm = 26,600 M-1 cm-1) or DMP (Ɛ468nm = 49,600 M-1 cm-1) in 20 mM acetate buffer at pH 4.3 in the presence of 0.2 mM H2O2 at 25°C using a Nicolet Evolution 300 spectrophometer (Thermo Industries). The KRX strain was selected for further studies based on the highest observed specific activities among the strains tested (Table S2). Libraries construction. Libraries were generated by random mutagenesis with error-prone PCR. Primers 5’-GGATTAGCCTCATATGCCGTTCCAGCAAGG-3’ (PpDyP Forward) and 5’-GTGTTTCTGTATCTGGATCCTTAGAGATCAGGCCCGC-3’ (PpDyP Reverse) were used for amplification (NdeI and BamHI restriction sites are underlined). ep-PCR was performed in a 50 µL reaction using 3 ng of DNA template, 0.5 µM of primers, 200 µM of dNTPs, 7 mM MgCl2, Taq polymerase buffer and 5 U of Taq polymerase (Fermentas). In order to select the concentration of MnCl2 in the ep-PCR protocol seven libraries of ppDyP mutant genes were constructed with Taq polymerase in the presence of 0.01 - 0.2 mM MnCl2 (Figure S1). After an initial denaturation period of 10 min at 94°C, the reaction was thermally cycled 28 times (1 min at 94°C, 1 min at 57°C and 1.5 min at 72°C; MyCyclerTM thermocycler, Biorad) followed by 10 min at 72°C. The resulting gene libraries were purified, (GFX PCR DNA and the Gel Band Purification kit (GE Healthcare)) cloned into pET-21a (+) (Novagen) and transformed into E. coli KRX by electroporation. The concentration of 0.01 mM MnCl2, which leads to 30% of the total number of clones with less than 10% activity of wild type (Figure S1) and putatively result in a mutation rate of 1-3 amino acid substitutions per gene48, was chosen for the construction of libraries of mutants; indeed, the DNA sequence revealed that 5 out 6 clones show one mutation and 1 out 6 has two mutations. Expression of ppDyP wild-type and variants. Agar colonies were individually picked and transferred to 96-well plates and growth as previously described.49 Induction of ppDyP gene was performed after 4 h of growth by adding 0.2% rhamnose and 15 µM hemin. Cultures 5


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were harvested by centrifugation after 24 h of cultivation (2,750g for 30 min at 4°C) and were disrupted through (i) addition of 100 µL of a 40% Bacterial Protein Extraction Reagent (BPER®) lysis solution (Thermo Fisher), (ii) 3 freeze and thaw cycles (immersion in liquid nitrogen/ room temperature (5 min)) followed by suspension in 100 µL of 20 mM Tris-HCl, pH 7.6 and, (iii) a combination of freeze and thaw cycles (- 80°C (15 min)/ 30°C (5 min)) followed by suspension in 100 µL of 20 mM Tris-HCl, pH 7.6 supplemented with lysozyme (2 mg·mL-1). After centrifugation cell supernatants were used for activity measurements. The lowest coefficient of variance (CV = standard deviation/ mean × 100%) for protein content (17%) and activity (21%) were achieved using the combined physical and enzymatic procedure (Table S3). Activity screenings in 96-well plates. Enzymatic activities were performed in 200-µL reactions containing 180 µL of 20 mM acetate buffer at pH 4.3 containing 0.2 mM H2O2, 1 mM ABTS or 1 mM DMP and 20 µL crude extracts. The reactions were monitored in a Synergy 2 (BioTek) microtiter plate reader at 420 nm and at 468 nm for ABTS and DMP oxidation, respectively. DNA was extracted (GeneJET Plasmid Miniprep Kit (Thermo Scientific)) and the presence of mutations were verified by DNA sequencing analysis using T7 terminator universal primers. The variant showing the highest activity for ABTS or DMP oxidation in each generation was selected for parenting the next generation. Construction of the A142V, T134V, T138V and M212L variants using site directed mutagenesis. The Quick change mutagenesis protocol (Stratagene) was used to introduce single amino acid replacements in the ppDyP gene. Plasmid pRC-1, with ppDyP, was used as template in the PCRs reactions (50 µL): 3 ng of DNA template, 2 µM of primers (Table S1), 200 µM of dNTPs, NZYProof polymerase buffer, and 1.25 U of NZYProof polymerase (NZYTech). After 4 min at 95°C, the reaction was thermally cycled 20 times (1 min at 95°C, 1.5 min at 62-71°C and 7 min at 72°C) followed by 10 min at 72°C. The amplified products were digested with the endonuclease DpnI and transformed into E. coli DH5α cells. DNA sequencing was used to confirm the presence of the desired mutations. Production and purification of wild-type and variants. The genes coding for wild-type enzyme, hit variants of each evolution generation (9F6, 31F3, 17F11, 21G11, 6E10) and variants obtained by site-directed mutagenesis (A142V, T134V, T138V and M212L), were cloned into pET21a (+) plasmid and introduced into the host E. coli Tuner (DE3, Novagen). Recombinant enzymes were produced in 1L of LB medium in 5L-Erlenmeyers and purified as previously described.38 Variants T134V, T138V and M212L were purified using only one chromatographic step (Q-Sepharose). The concentration of purified proteins was estimated using the molar absorption coefficient of PpDyP (ε280 = 34,850 M-1·cm-1), calculated from the protein sequence using the ExPASy Bioinformatics Resource Portal (http://web.expasy.org). Biochemical analysis. UV-visible absorption spectra, heme content and cycle voltammetry measurements of purified enzymes were performed following previously optimized procedures.24,38 6


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Apparent steady-state kinetic analysis. Enzymatic activities of PpDyP wild-type and variants were spectrophotometrically monitored at 25°C. The activity dependence on pH was measured by monitoring the oxidation of reduced substrates at their maximal absorption wavelengths at 25°C using Britton-Robinson buffer (100 mM phosphoric acid, 100 mM boric acid, and 100 mM acetic acid mixed with NaOH to the desired pH in the range 2 - 11). Apparent steady-state kinetic parameters for substrates were measured in 100 mM sodium acetate or phosphate buffer at the optimal pH of the enzymes; for H2O2 (0.005 - 5 mM) in the presence of the 10 mM ABTS for wild-type and 3 mM for the variants; for ABTS (0.1 - 10 mM), guaiacol (0.001 - 3 mM), DMP (0.01 - 2 mM), syringaldehyde (SA) (0.02 - 0.4 mM; ε300nm pH 4.3 = 6,923 M-1 cm-1, ε360nm pH 8.5 = 16,000 M-1 cm-1)50, acetosyringone (AS) (0.02 0.4 mM; ε300nm pH 4.3 = 7,076 M-1cm-1, ε360nm pH 8.5 = 12,000 M-1 cm-1)50, methyl syringate (MS) (0.02 - 0.4 mM; ε275nm pH 4.3 = 7,846 M-1 cm-1, ε320nm pH 8.5 = 12,461 M-1 cm-1)50, 4aminodiphenylamine (4-ADA) (0.01 - 2 mM; ε510nm = 1,046 M-1 cm-1), 2,5diaminobenzenesulfonic acid (2,5-DABSA) (0.02 - 3 mM; ε470nm = 2,372 M-1 cm-1), 1,2phenylenediamine (1,2-PDA) (0.01 - 10 mM; ε440nm = 18,387 M-1 cm-1) and 1,4phenylenediamine (1,4-PDA) (0.01 - 10 mM; ε510nm pH 5 = 3,467 M-1 cm-1, ε460nm pH 8 = 3,408 M-1 cm-1) in the presence of 0.5 mM H2O2. Enzymatic oxidation of 0.015-0.4 mM alkali Kraft lignin (Sigma-Aldrich) was performed with 2.5 mM H2O2 and 10 U of enzyme in 100 mM sodium acetate or phosphate buffer and monitored at 465 nm,15,18 the addition of 2 mM MnCl2 or 2 mM methylsyringate were also tested. The molecular concentration of Kraft lignin was calculated using an average molecular mass of 10,000 Da. Kinetic data was fitted directly using the Michaelis-Menten equation or the equation for non-linear curve that fits enzyme kinetics affected by substrate inhibition (v = Vmax[S]/(Km+[S](1+[S]/Ki))) (Origin software). Products from the oxidation of aromatic amines. The reaction mixtures contained 5 mM of the primary aromatic amines, 4-ADA, 2,5-DABSA, 1,4-PDA or 1,2-PDA in 50 mM acetate buffer pH 4.5, 0.2 mM of H2O2 and 1 U·mL-1 of wild-type PpDyP. After 24 h at room temperature reaction mixtures were diluted 1:1 in methanol and 20 µL of the resultant solution were injected and analysed by ESI-MS. LR ESI mass spectra were obtained on a LCQ Fleet mass spectrometer operated in the ESI positive/negative ion modes (Thermo Scientific) as described previously.51 Oxidation of guaiacylglycerol-β β-guaiacyl ether (GGE). The enzymatic oxidation of GGE was monitored by high-performance liquid chromatography (HPLC). The reaction mixtures (0.5-1 mL) contained 2.5 mM of GGE dissolved in 1:1 ethanol/buffer, 2.5 mM H2O2 and 10 U of enzyme in sodium acetate pH 5 for PpDyP and sodium phosphate buffer pH 7.5 for 6E10. Reactions in the presence of 2 mM MnCl2 were also performed. After different times, aliquots were withdrawn and diluted 1:1 in methanol before injection and analyzed in a HPLC Merck Hitachi with a diode-array (DAD) system, using a reverse phase C-18 column (LiChrospher 100 RP-18, Merck, Darmstad, Germany) with a pre-column at 40ºC and a flow rate of 1 mL·min-1. Reaction products were eluted with a linear gradient of H2O:methanol plus 0.5% v/v of acetic acid from 30% to 80% of methanol over 7 min, and maintained 7


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isocratic at 80% for 2 min. The starting conditions were restored maintained isocratic for 9 min. Samples were also analyzed by HPLC ESI-MS using Dionex Ultimate 3000 composed of a binary pump HPG3200, an autosampler WPS300 and a column oven TCC3000 coupled in-line to a LCQ Fleet ion trap mass spectrometer equipped with an ESI ion source (Thermo ScientificTM). Separations were carried out with a Purespher Star C18 (250 x 4.6 mm. 5 µm, Merck) at 30ºC, using a flow rate of 0.3 mL·min-1, a mobile phase of 0.5% of acid formic in water (v/v, eluent A) and methanol (eluent B), with a linear gradient from 50 to 80% B over 3 min followed by a 9 min isocratic 80% B, and then the column was re-equilibrated with 50% B for 10 min. The mass spectrometer was operated in the ESI positive and negative ion modes as described previously.51 The Xcalibur software was used for data acquisition and processing. Kinetic stability. Thermal inactivation assays were performed as previously described.38 In brief, enzyme solutions were incubated at 40°C in 20 mM Tris-HCl buffer, pH 7.6, and at fixed time intervals, sample aliquots were withdraw and tested for activity following the ABTS oxidation at 25°C. Thermal inactivation appears to obey first-order kinetics and the half-life value (t1/2) was calculated using t1/2= ln2/kin. Stopped-flow experiments. Transient-state kinetics were performed by following previously optimized methodologies.23 In brief, 6E10 preparations were mixed with H2O2 or guaiacol at pH 8.5 in 0.1 M sodium phosphate buffer at 25°C in a Hi-Tech SF-61DX2 stopped-flow apparatus coupled to a diode array. A 2 µM of enzyme (final concentration) was mixed with 0 - 200 µM H2O2. UV-Vis spectra (350-700 nm) were recorded at different time scales. The rate constants for Cpd I formation (k1obs) were obtained from the exponential fit of the absorbance decrease measured at 404 nm. Cpd II formation (k2obs) and conversion to the resting enzyme (k3obs) in the presence of guaiacol were obtained from the exponential fit of the absorbance increase measured at 450 nm and 415 nm, respectively. In these experiments, a preparation of PpDyP 6E10 in the presence with 1 equivalent of H2O2 was mixed with increasing concentrations of guaiacol. The second-order rate constants k2’ (Cpd I decay to Cpd II) and k3´ (Cpd II decay to the resting enzyme) were obtained from the slope of the plots k2obs vs [guaiacol] and k3obs vs [guaiacol], respectively.23 RESULTS AND DISCUSSION Directed evolution of PpDyP Three rounds of evolution through ep-PCR were performed and a total of approximately 6,000 clones were screened to find variants with improved catalytic efficiency for DMP (Figure 1). In the 1st generation, the library of variants was screened for ABTS oxidation since activity for DMP was not detected in crude extracts of E. coli cells grown in 96-well plates; after screening ~2,000 clones two variants, 31F3 (with single mutation H125R) and 9F6 (harboring the single mutation E188K), were identified showing 3- to 4-fold higher activity for ABTS when compared to the parent wild-type. Variant 9F6 was selected as the parent for the 2nd generation since residual activity for DMP was observed in this variant, in 8


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contrast to 31F3 where no activity for the phenolic substrate was detected. A total of ~2,000 clones were screened in the 2nd generation using both ABTS and DMP as substrates and 2 variants selected: the 21G11 variant (with mutations E188K and H125R) showing 6-fold higher activity for ABTS and 1.5-fold for DMP, and 17F11 variant (with mutations E188K and A142V) with a 3-fold higher activity for DMP and 2-fold higher for ABTS as compared to their parental strain (9F6). Variant 17F11 was used as the parent for the 3rd generation even though a reproducible high variation associated to this mutant (CV ~ 45%) was observed in the activity screening (data not shown). After screening ~2,000 clones, the 6E10 hit variant (with mutations E188K, A142V and H125Y) was identified showing 3.5- and 2-fold higher activities for DMP and ABTS, respectively, as compared to 17F11. The three acquired mutations in the course of the evolution (E188K, A142V and H125Y) affect amino acid residues situated at the surface of the PpDyP enzyme, in the second shell of the heme cavity accordingly to the PpDyP model structure (Figure 2A).38 This model structure was derived from the Phyre2 server (99% modelled at >90% confidence); six templates (DyPB from Rhodococcus jostii RHA1 (PDB 3qns; 29% identity), TyrA from Schewanella oneidensis (PDB 2iiz; 27% identity), VcDyP from Vibrio choleroe (PDB 5de0, 27% identity), DyP from Thermobifida cellulosilytica (PDB 4gs1, 28% identity), BtDyP from Bacteroides thetaiotaomicron (PDB 2gvk, 25% identity) and DyP from Streptomyces coelicolor (PDB 4gt2; 27% identity)) were selected to model PpDyP based on heuristics to maximize confidence, percentage identity and alignment coverage.52 The position E188K is located at ∼ 5 Å to the heme 6-propionate side chain, in the surface loop I179-A196 at the entry to the heme access channel, a putative binding site for small substrate molecules29,30; A142V, affects a small α-helix E140-A146, close to the loop D117-D139, that contains the catalytic distal residues D132 and N136 and finally position H125Y is located in the loop D117-D139, at ∼ 4 Å from D132. Biochemical and structural analysis of the hit variants 9F6, 17F11 and 6E10 In order to characterize the laboratory evolution of P. putida MET94 PpDyP and assess the structural basis of the improved catalytic efficiency, variants 9F6, 17F11 and 6E10, were overproduced in E. coli in a batch scale, purified and characterized. The Reinheitszahl values of purified enzymes were between 1.3 and 1.9 and the heme b content was ~ 1 mol per mole of protein, similarly to the wild-type enzyme (Table 1). The overall electronic absorption features of PpDyp variants as assessed by UV-vis revealed the characteristic Soret band at ~ 400 nm, Q bands at ~ 502 and 545 nm, and a charge transfer band at ~ 630 nm (Figure S2).38 The formation of an intermediate with spectral features of Cpd I (decrease in intensity of a Soret band, disappearance of Q bands and appearance of a band ~ 550-600 nm) was observed upon addition of 1 equivalent of hydrogen peroxide (Figure S2). The catalytic properties of purified variants were determined at the optimal pH using H2O2, ABTS and DMP as substrates (Table 2 and 3, Figure S3). The replacement of E188K, situated at ∼ 5 Å to the 6-heme propionate (Figure 2A), in the 9F6 variant, selected after a screening for ABTS oxidation, resulted in a 7-fold higher turnover rate (kcat) for H2O2 but to a 9


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4-fold higher Km, resulting in a catalytic efficiency ((kcat/Km) only slightly higher than the wild-type (Table 2). Notably, one order of magnitude higher efficiency for ABTS was observed when compared to the wild-type as consequence of a ∼ 6-fold increased kcat and ~ 10 times decreased Km. E188 is close to several acidic residues at the main heme access channel (Figure 2B) and its replacement by a positively charged lysine likely favors accommodation of the anionic substrate ABTS, facilitating its oxidation to the 6-heme propionate. The structure of DyPDec1 solved in complex with 2,6-dimethoxyphenol and ascorbate show both substrates interacting with the heme propionate through hydrogen bonding networks.30 The kinetic results agree well to the significantly enhanced redox potential of the Fe2+/Fe3+ couple (E0´ = -110 mV) in the 9F6 variant in comparison to the wild-type enzyme (E0´ = -260 mV) (Table 1) which can be explained on a purely electrostatic basis: the removal of a negative charge in the vicinity of the heme should facilitate its reduction. Likewise, the replacement of the distal residues D132 with an asparagine or N136 and R214 with a leucine in PpDyP resulted in significantly increased E0´ by 140-160 mV.24 The E188 residue, as well as distal Asp, Asn and Arg residues participate in an extensive network of hydrogen bonds interacting with the distal and proximal heme sites, influencing the polarity of the heme environment. This view is supported by the highly flexible heme cavity of PpDyP35,36, pointing to the importance of electrostatics and the hydrogen bond network of the second coordination sphere of the active site of PpDyP. Variant 17F11 (E188K and A142V) selected in the second generation, after a screening for DMP oxidation, exhibits a two order of magnitude enhanced catalytic efficiency (kcat/Km = 0.9 × 105) for DMP oxidation as compared to 9F6 or wild-type (Table 3, Figure S3B) and exhibited values similar to those of Phanerochaete crysosporium LiP and MnP53,54 Pleurotus eryngii VP39 and Irpex lacteus DyP.40 In contrast, the observed efficiency for ABTS resembles that of the wild-type and is 10-fold lower than that of the 9F6 variant from the 1st generation due to a 4-fold higher Km value (Table 2, Figure S3A). These results show that introduction of A142V mutation improve the enzyme specificity for DMP at the expense of that for ABTS, i.e. a trade-off in the oxidation of the two substrates occurred suggesting that both substrates share the same interaction site on the enzyme surface, at the heme access channel, but e.g. their modes of binding differ. Weak trade-offs have been related with the conformational flexibility of proteins allowing for alternative conformations to mediate new functions without severely compromising the conformation that mediates the existing function.55 Variant 17F11 harbors two additional notable features: i) a higher pH optima for both ABTS (pH 5.5) and DMP (pH 8.5) as compared to wild-type (4.3) (Figures S4 A,B), and ii) a 4-fold higher Ki for H2O2, (~ 2.3 mM) (Figure 4C). Therefore mutation A142V, introduced in the 2nd generation is responsible, alone or in combination with E188K, for the observed remarkable properties. In order to confirm this hypothesis, the single variant A142V was constructed using sitedirected mutagenesis, overproduced in E. coli, purified and characterized. The single substitution A142V in the wild-type led to 5-fold lower catalytic efficiency to H2O2 which may likely explain the 7-fold inferior efficiency for hydrogen peroxide exhibited by the 10


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17F11 variant as compared to the 9F6 variant (Table 2). In contrast, while the replacement of Ala142 by a Val is beneficial for both ABTS and DMP oxidation (2- and 8-fold enhanced kcat/Km) in the wild-type enzyme, opposite effects are observed in the 9F6 background; it is detrimental for ABTS oxidation (40-fold lower catalytic efficiency) and beneficial for DMP oxidation (60-fold higher efficiency) as observed in the 17F11 variant (Table 2 and 3). These results suggest that the contribution of A142V for the enzyme fitness, i.e. its catalytic efficiency (kcat/Km), is dependent on the genetic background and therefore, its effect in the 17F11 variant depends on the presence of E188K mutation, a phenomenon known as intragenic epistasis.56 The level of pairwise epistasis between the mutations A142V and E188K was measured as the difference between the observed fitness (W) of the double mutant, variant 17F11 (harboring A142V and E188K mutations) and the expected fitness based on the effects of each individual mutation (variant A142V and 9F6 with E188K mutation) relative to the wild-type: ε =W17F11 – WA142V*W9F6. Epistasis is said to be positive when ε > 0, and negative when ε < 0. Therefore, negative and positive epistasis between A142V and E188K residues is detected for the catalytic efficiency of ABTS (ε = -124 ± 0.5) and DMP (ε = 106 ± 9), respectively. Discrimination is mainly manifested in the Km factor, since introduction of A142V mutation in the wild-type led to a 10-fold decreased Km value for ABTS and 4-fold increased Km for DMP, while the opposite is observed in the 9F6 background (5-fold increase for ABTS and 1.5-fold decrease for DMP), whereas kcat values consistently decreased for ABTS and increased for DMP upon replacement of A142V in both backgrounds (Table 2 and 3). Previous studies suggest that a significant number of epistatic effects stem from long-range interactions and could be mediated by loop repositioning, protein dynamics, or through displacement of active site residues or co-factors.56 Residues A142V and E188K are located ∼ 20 Å apart, in opposite sides of the heme group, and are not expected to directly interact. The replacement of alanine, considered the finest helix-forming residue, by a valine, might have decreased the helical propensity of secondary α-helix segment E140-A146, affecting the local structure of loop D117-D139 leading to its conformational repositioning (Figure 2A). This in turn may impacted interactions with the two other flexible loops close to the heme group; loop A202-G226, which is linked by the small α-helix H197-T201, to loop I179-A196, harboring E188K. Note that charged residues at the heme surface are part of the three loops (Figures 2A,B). Together, these changes would alter the physical properties of surface accessible side chains in the heme channel access of variant 17F11 directly impacting its enzyme efficiency for substrates. Interestingly, A142V variant shows, similarly to 17F11, increased pH optima for both ABTS (5.5) and DMP (8.5) and resistance to hydrogen peroxide (Figures S5 A-C). In a previous work, we employed site directed mutagenesis to investigate the role of distal residues (D132, N136 and R214) in the peroxidative cycle of PpDyP.24 Variants N136L and D132N showed an optimal pH at 5.6 and 7.4, respectively, which corresponds to 2 and 3 units up-shifted as compared to wild-type (pHop = 4.3), while variant R214L showed an optimum at pH 3.6. The opposite pH shifts in the analyzed variants was correlated to a cooperative action between the Asp and Arg residues in the acid-base mechanism involved in H2O2 binding and fission in the heme pocket of PpDyP; in the absence of Asp (D132N), the optimal pH shifted to values 11


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ACS Catalysis

closer to the pKa value of Arg (12.48) and in the absence of Arg (R214L), a down-shift to values lower than the pKa value of Asp (3.9).24 Therefore the increased optimal pH exhibited by 17F11 variant may at least partially, reflected structural reorganization introduced by the A142V mutation that affected the relative position (and eventual displacement) of the catalytic residues to the heme site and/or eventually changes of the pKa value of the Asp132 that is sensitive to local conformational fluctuations. Nevertheless, neither of these hypotheses could be evaluated here. The inhibition of peroxidative activity in the presence of high concentrations of H2O2 has been related to the oxidation of sensitive residues (Arg, Asn, Met, Lys, Thr and Trp) located close to the active site, or at the surface of the peroxidases.41,45,57-61 Analysis of the PpDyP heme cavity showed the presence of three solvent-exposed residues T138, T134 and M212 (Figure 2C), which are putatively exposed to the oxidizing effects of H2O2. To test this hypothesis, single variants T134V, T138V and M212L were constructed by site-directed mutagenesis, overproduced in E. coli, purified and characterized. The T138V variant exhibits a 3-fold higher Ki for H2O2, (~ 1.9 mM) as compared to the wild-type (0.7 mM) whereas T134V and M212L variants, located closer (∼ 4Å) to the heme, show resistance to hydrogen peroxide inhibition (Figure S5D). Therefore, our hypothesis is that introduction of the A142V mutation followed by the putative conformational rearrangements in the 17F11 variant, as discussed above, protected sensitive amino acid residues close to the heme cavity from H2O2 oxidation. The biochemical analysis of variant 6E10 (E188K, A142V and H125Y) showed that substitution H125Y led to similar catalytic efficiency for both ABTS and DMP as compared to 17F11 variant (Table 3) (Figures S4 A, B). The observed 2-fold increased production levels of the 6E10 variant (Table 1) is a direct consequence of mutation H125Y and is corroborated by the characterization of variants 31F3 (H125R) and 21G11 (E188K and H125R), identified during the course of the evolution experiment (Figure 1). These variants exhibits similar protein yields (12-15 mg·mL-1) to those observed in 6E10 variant (12 mg·mL-1) (Table 1). Therefore residue His125, located in the loop D117-D139 seems to represent a structural hot-spot involved in the modulation of the levels of soluble PpDyP as a consequence for example, of a reduced proteolysis in the cytoplasm of E. coli host strain. This hypothesis is based in the thermal inactivation profiles of the wild-type and variant enzymes indicating that variant 9F6 showed a lower half-life inactivation at 40°C (52 ± 8 min-1, Figure S6) when compared with the wild-type (110 ± 17 min-1) also observed in the variant 17F11 (36 ± 5 min-1). However, the 6E10 variant, harboring the additional mutation H125Y presented a half-life value (114 ± 12 min-1) similar to the wild-type. Substrate range of the 6E10 variant The substrate range of the 6E10 variant was further investigated using a variety of phenolic, Kraft lignin and aromatic amine substrates (Table 4 and Figure S7); this variant exhibited for all substrates tested, 2 to 100-fold improved oxidation efficiencies. Interestingly and in contrast to the wild-type PpDyP38, 6E10 was not inhibited at high substrate concentrations of 12


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the phenolics tested (above 0.25 - 0.3 mM) (data not shown). The products obtained from the oxidation of the aromatic amines were identified using ESI-MS (Table 5). The reactions of para-substituted primary aromatic amines with electron donor groups, 1,4-PDA and 4-ADA, generated the colored di-substituted benzoquinonediimine trimeric structures, identical to those previously obtained using CotA-laccase from B. subtillis; the product of 1,4-PDA is known as the Bandrowski’s base, with a well-established application in hair and fur dyeing.62 The enzymatic oxidation of ortho and meta,para-disubstituted aromatic amines, 1,2-PDA and 2,5-DABSA, produce the phenazine cores, heterocyclic nitrogen containing compounds that are important biological active motifs of antibiotics and antibacterial agents, anti-tumor agents, pesticides, dyestuffs, biosensors and are the building blocks for the synthesis of organic semiconductors or electrical-photochemical materials.63 The transformation of alkali Kraft lignin was investigated by monitoring at 465 nm the time course of reactions.15,18 The catalytic efficiency (kcat/Km) of 6E10 variant (4.1 × 103 s-1·M-1) was 2.5-fold higher than the wild-type (Table 4) but two-order of magnitude lower than reported for P. fluorescens DyP1B and DyPA.64 Addition of MnCl2 to reaction mixtures resulted in similar catalytic efficiencies but methyl syringate led to one order of magnitude increased kcat/Km (Table 4). The oxidation of guaiacylglycerol-β-guaiacyl ether, a phenolic type model compound containing the β-O-4 bond which constitutes 50 to 70% of intersubunit bonds in lignin, by the 6E10 variant resulted in a 70% conversion yield after 1 h of reaction at pH 7.5 (Figure 3A), contrasting with only 20% conversion observed using the wild-type enzyme at pH 5 (Figure 3B). These values remain unchanged up to 96 h of reaction. The differences in the conversion yields between the two enzymes are likely due to the differences in the pH of reactions. Addition of MnCl2 led to analogous results (data not shown). Similar yields of GGE conversion (65%) were observed with lignin peroxidase from Pleurotus eryngii.65 The measured specific activities are 22 ± 1 and 12 ± 4 U·mg-1, for 6E10 and wild-type, respectively, 1000-fold higher as compared to the only value reported in the literature (0.017 U·mg-1 for R. jostii DyP).16 Analysis of the reaction products of GGE conversion by liquid HPLC-mass spectrometry in positive mode, yielded m/z 661 [M+Na]+ and 635 [M+H]+, consistent with molecular formulas C34H38O12 and C34H34O12, respectively, and the structures presented in Figure 3C. This data suggests that the main GGE degradation patterns performed by PpDyP resulted from oxidative dimerization, probably formed through C-C coupling of the phenolic unit, and a Cα-oxidation. To test whether the GGE dimerization is dependent on substrate concentration assays were performed at lower concentrations of substrate (0.5-2.5 mM), but no differences were detected in the nature of products formed. The C-C coupling of the phenolic unit was previously observed in reactions using the enzyme TfuDyP from T. fusca.66 In contrast, DyP from R. jostii, cleaves the Cα−C β bond similarly to what was observed with fungal LiPs.15,67 The reasons behind the differences in the mechanism of degradation of the lignin phenolic model compounds among the different DyPs are not clear at present. Catalytic mechanism of PpDyP 13


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PpDyP shows concentration and pH-dependent aggregation which prevents investigating its catalytic cycle at pH 4.3, the optimal for activity.24,38 Variant 6E10 shows a similar aggregation profile as the wild-type enzyme, but at pH 8.5, the optimal for phenolics oxidation, aggregation is not observed. Therefore its catalytic cycle was investigated using transient kinetics. The addition of 5 µM H2O2 to 6E10 at pH 8.5, revealed spectral changes indicative of Cpd I formation (Figure 4A). The rate constants of Cpd I formation (k1obs) are linearly proportional to the H2O2 concentration up to 30 µM, whereas at higher concentrations saturation occurs (Figure 4B). This is consistent with a two-step mechanism as first suggested by Poulos and Kraut 68: (i) formation of a a complex E-H2O2, often called Compound 0, and (ii) the heterolytic O-O bond cleavage producing Cpd I and water: K

1

k1

E + H2O2 ⇋ E - H2O2 → Cpd I + H2O

(1)

In which the K1 (M) is an apparent dissociation constant and k1 (s-1) is a first order catalytic rate constant. The kinetic evidence for a two-step mechanism was recently reported for Bacillus subtilis BsDyP.23 Prior to these studies the formation of Compound 0 was only described in low-temperature studies of horseradish peroxidase and variants, and lactoperoxidase using cryoradiolysis and RR spectroscopy.69-75 The constants K1 and k1 are (44 ± 3) × 106 M and 133 ± 4 s-1, respectively. These data pass through the origin, indicating that the second step of the reaction is irreversible. The second-order rate constant of Cpd I formation (k1’ = k1/K1 = (3 ± 0.3) × 106 M-1 s-1) is of the same order of magnitude as those previously reported for BsDyP 23, lignin and manganese peroxidase from the white-rot fungus Phanerochaete chrysporium 76 and versatile peroxidase from P. eryngii. 77 Next, Cpd I was generated by mixing 6E10 with H2O2 (1:1) and reacted with increasing concentrations of guaiacol. The formation of Cpd II was completed in less than 200 ms and the second-order rate constant for the Cpd I reduction to Cpd II (k2’ = (6.1 ± 0.3) × 105 M-1 s1 ) was estimated from the slope of the plot in Figure 4C assuming the kinetic scheme: k2 ’

Cpd I + AH → Cpd II + A

·

(2)

This value is of the same order of magnitude as those previously reported for BsDyP from B. subtilis23 and lignin and manganese peroxidases from P. chrysporium.78 The observed pseudo first order rate constant (k3obs) for Cpd II reduction was linearly proportional to the guaiacol concentrations used in the reactions, indicating that the reaction follows second order kinetics (Figure 4D) in contrast with results obtained with BsDyP23 and in accordance with data obtained for P. chrysporium manganese peroxidase.54,78,79 The apparent second order rate constant for the reduction of Cpd II to the resting enzyme by guaiacol (k3’) was (1.2 ± 0.3) × 102 M-1 s-1, the rate-limiting step in the PpDyP catalytic reaction, as typically observed in peroxidases. The obtained results allow for the proposal of a catalytic cycle for PpDyP as shown in Figure 5.

14


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CONCLUSIONS DyPs are potentially important biocatalysts for industrial oxidations and redox conversion processes in particular for lignin deconstruction. This is considered a key process for utilization of lignocellulosic biomass for bulk and fine chemicals, materials and biofuels. In this study we provide the first laboratory evolution study of a bacterial dye-decolorizing peroxidase. The evolved variant of P. putida MET94 PpDyP has improved catalytic efficiencies at alkaline pH for phenolic and aromatic amines substrates, including Kraft lignin and the model lignin dimer guaiacylglycerol-β-guaiacyl ether, representative of the main chain linkage type in lignin. This variant was also shown to be resistant to high hydrogen peroxide concentrations, one of the main limitations to the biotechnological applications of peroxidases, and provide clear advantages for application in bioprocesses that occur at the neutral to alkaline conditions, for example those used in lignocellulose processing. Fungal ligninolytic peroxidases, despite their biotechnological interest, are not functional at basic pH due to the loss of calcium ions that affects their structural integrity and functional performance. This report highlights the value of directed evolution, not only to generate enzymes with improved properties for applications in organic synthesis, pulp bleaching or lignin valorization, but also for studying protein structure-function relationships. This study opens perspectives for further evolution of these enzymes for new properties and for studying protein structure and interactions within the DyP-type peroxidase family of enzymes. ACKNOWLEDGMENTS The authors thank P. Durão, M. Conceição Oliveira and S. Todorovic for helpful discussions. This work was supported by Fundação para a Ciência e Tecnologia (FCT), Portugal (PTDC/BBB-EBB/0122/2014, RECI/QEQ-QIN/0189/2012 and REM2013) and Research Unit GREEN-it "Bioresources for Sustainability" (UID/Multi/04551/2013). V.B. holds a Post-doc fellowship (SFRH/BPD/109431/2015) from FCT, Portugal. SUPPORTING INFORMATION AVAILABLE: Supporting information consists of 3 additional tables (Tables S1-S3) and 8 additional figures (Figures S1-S8). REFERENCES (1) (2) (3) (4) (5)

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Syed, K.; Yadav, J. S.; Doddapaneni, H.; Subramanian, V.; Lavin, J. L.; Oguiza, J. A.; Perez, G.; Pisabarro, A. G.; Ramirez, L.; Santoyo, F.; Master, E.; Coutinho, P. M.; Henrissat, B.; Lombard, V.; Magnuson, J. K.; Kues, U.; Hori, C.; Igarashi, K.; Samejima, M.; Held, B. W.; Barry, K. W.; LaButti, K. M.; Lapidus, A.; Lindquist, E. A.; Lucas, S. M.; Riley, R.; Salamov, A. A.; Hoffmeister, D.; Schwenk, D.; Hadar, Y.; Yarden, O.; de Vries, R. P.; Wiebenga, A.; Stenlid, J.; Eastwood, D.; Grigoriev, I. V.; Berka, R. M.; Blanchette, R. A.; Kersten, P.; Martinez, A. T.; Vicuna, R.; Cullen, D. PNAS 2012, 109, 5458-5463. Floudas, D.; Binder, M.; Riley, R.; Barry, K.; Blanchette, R. A.; Henrissat, B.; Martinez, A. T.; Otillar, R.; Spatafora, J. W.; Yadav, J. S.; Aerts, A.; Benoit, I.; Boyd, A.; Carlson, A.; Copeland, A.; Coutinho, P. M.; de Vries, R. P.; Ferreira, P.; Findley, K.; Foster, B.; Gaskell, J.; Glotzer, D.; Gorecki, P.; Heitman, J.; Hesse, C.; Hori, C.; Igarashi, K.; Jurgens, J. A.; Kallen, N.; Kersten, P.; Kohler, A.; Kues, U.; Kumar, T. K.; Kuo, A.; LaButti, K.; Larrondo, L. F.; Lindquist, E.; Ling, A.; Lombard, V.; Lucas, S.; Lundell, T.; Martin, R.; McLaughlin, D. J.; Morgenstern, I.; Morin, E.; Murat, C.; Nagy, L. G.; Nolan, M.; Ohm, R. A.; Patyshakuliyeva, A.; Rokas, A.; Ruiz-Duenas, F. J.; Sabat, G.; Salamov, A.; Samejima, M.; Schmutz, J.; Slot, J. C.; St John, F.; Stenlid, J.; Sun, H.; Sun, S.; Syed, K.; Tsang, A.; Wiebenga, A.; Young, D.; Pisabarro, A.; Eastwood, D. C.; Martin, F.; Cullen, D.; Grigoriev, I. V.; Hibbett, D. S. Science 2012, 336, 1715-1719. Masai, E.; Katayama, Y.; Fukuda, M. Biosci Biotech Bioch 2007, 71, 1-15. Bugg, T. D.; Ahmad, M.; Hardiman, E. M.; Rahmanpour, R. Nat Prod Rep 2011, 28, 18831896. Brown, M. E.; Chang, M. C. Curr Opin Chem Biol 2014, 19, 1-7. de Gonzalo, G.; Colpa, D. I.; Habib, M. H.; Fraaije, M. W. J Biotechnol 2016, 236, 110-119. Sugano, Y. Cell Mol Life Sci 2009, 66, 1387-1403. Hofrichter, M.; Ullrich, R.; Pecyna, M. J.; Liers, C.; Lundell, T. Appl Microbiol Biotechnol 2010, 87, 871-897. Colpa, D. I.; Fraaije, M. W.; van Bloois, E. J Ind Microbiol Biotechnol 2014, 41, 1-7. Liers, C.; Bobeth, C.; Pecyna, M.; Ullrich, R.; Hofrichter, M. Appl Microbiol Biotechnol 2010, 85, 1869-1879. Ahmad, M.; Roberts, J. N.; Hardiman, E. M.; Singh, R.; Eltis, L. D.; Bugg, T. D. Biochemistry 2011, 50, 5096-5107. Brown, M. E.; Barros, T.; Chang, M. C. ACS Chem Biol 2012, 7, 2074-2081. Singh, R.; Grigg, J. C.; Qin, W.; Kadla, J. F.; Murphy, M. E.; Eltis, L. D. ACS Chem Biol 2013, 8, 700-706. Rahmanpour, R.; Rea, D.; Jamshidi, S.; Fulop, V.; Bugg, T. D. Arch Biochem Biophys 2016, 594, 54-60. Ruiz-Duenas, F. J.; Lundell, T.; Floudas, D.; Nagy, L. G.; Barrasa, J. M.; Hibbett, D. S.; Martinez, A. T. Mycologia 2013, 105, 1428-1444. Linde, D.; Ruiz-Duenas, F. J.; Fernandez-Fueyo, E.; Guallar, V.; Hammel, K. E.; Pogni, R.; Martinez, A. T. Arch Biochem Biophys 2015, 574, 66-74. Kellner, H.; Luis, P.; Pecyna, M. J.; Barbi, F.; Kapturska, D.; Kruger, D.; Zak, D. R.; Marmeisse, R.; Vandenbol, M.; Hofrichter, M. PLOs ONE 2014, 9, e95557. Singh, R.; Eltis, L. D. Arch Biochem Biophys 2015, 574, 56-65. Mendes, S.; Catarino, T.; Silveira, C.; Todorovic, S.; Martins, L. O. Cat Sci & Technol 2015, 5, 5196-5207. Mendes, S.; Brissos, V.; Gabriel, A.; Catarino, T.; Turner, D. L.; Todorovic, S.; Martins, L. O. Arch Biochem Biophys 2015, 574, 99-107. Chen, C.; Shrestha, R.; Jia, K.; Gao, P. F.; Geisbrecht, B. V.; Bossmann, S. H.; Shi, J.; Li, P. J Biol Chem 2015, 290, 23447-23463. Strittmatter, E.; Liers, C.; Ullrich, R.; Wachter, S.; Hofrichter, M.; Plattner, D. A.; Piontek, K. J Biol Chem 2013, 288, 4095-4102. Linde, D.; Pogni, R.; Canellas, M.; Lucas, F.; Guallar, V.; Baratto, M. C.; Sinicropi, A.; SaezJimenez, V.; Coscolin, C.; Romero, A.; Medrano, F. J.; Ruiz-Duenas, F. J.; Martinez, A. T. Biochem J 2015, 466, 253-262. 16


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(28) (29) (30) (31) (32) (33) (34) (35) (36) (37) (38) (39) (40) (41) (42) (43) (44) (45) (46) (47) (48) (49) (50) (51) (52) (53) (54) (55) (56) (57) (58) (59) (60)

Baratto, M. C.; Sinicropi, A.; Linde, D.; Saez-Jimenez, V.; Sorace, L.; Ruiz-Duenas, F. J.; Martinez, A. T.; Basosi, R.; Pogni, R. J.Phys Chem. B 2015, 119, 13583-13592. Strittmatter, E.; Serrer, K.; Liers, C.; Ullrich, R.; Hofrichter, M.; Piontek, K.; Schleicher, E.; Plattner, D. A. Arch Biochem Biophys 2015, 574, 75-85. Yoshida, T.; Tsuge, H.; Hisabori, T.; Sugano, Y. FEBS Lett 2012, 586, 4351-4356. Mendes, S.; Pereira, L.; Batista, C.; Martins, L. O. Appl Microbiol Biotechnol 2011, 92, 393405. Goncalves, A. M.; Mendes, S.; de Sanctis, D.; Martins, L. O.; Bento, I. FEBS J 2013, 280, 6643-6657. Brissos, V.; Goncalves, N.; Melo, E. P.; Martins, L. O. PLOs ONE 2014, 9, e87209. Mendes, S.; Farinha, A.; Ramos, C. G.; Leitao, J. H.; Viegas, C. A.; Martins, L. O. Bioresource Technol 2011, 102, 9852-9859. Sezer, M.; Genebra, T.; Mendes, S.; Martins, L. O.; Todorovic, S. Soft Matter 2012, 8, 1031410321. Sezer, M.; Santos, A.; Kielb, P.; Pinto, T.; Martins, L. O.; Todorovic, S. Biochemistry 2013, 52, 3074-3084. Todorovic, S.; Hildebrandt, P.; Martins, L. O. Phy Chem Chem Phys 2015, 17, 11954-11957. Santos, A.; Mendes, S.; Brissos, V.; Martins, L. O. Appl Microbiol Biotechnol 2014, 98, 2053-2065. Morales, M.; Mate, M. J.; Romero, A.; Martinez, M. J.; Martinez, A. T.; Ruiz-Duenas, F. J. J Biol Chem 2012, 287, 41053-41067. Salvachua, D.; Prieto, A.; Martinez, A. T.; Martinez, M. J. Appl Environ Microbiol 2013, 79, 4316-4324. Cherry, J. R.; Lamsa, M. H.; Schneider, P.; Vind, J.; Svendsen, A.; Jones, A.; Pedersen, A. H. Nature Biotechnol 1999, 17, 379-384. Morawski, B.; Quan, S.; Arnold, F. H. Biotechnol Bioeng 2001, 76, 99-107. Bulter, T.; Alcalde, M.; Sieber, V.; Meinhold, P.; Schlachtbauer, C.; Arnold, F. H. Appl Environ Microbiol 2003, 69, 987-995. Ryu, K.; Hwang, S. Y.; Kim, K. H.; Kang, J. H.; Lee, E. K. J Biotechnol 2008, 133, 110-115. Garcia-Ruiz, E.; Gonzalez-Perez, D.; Ruiz-Duenas, F. J.; Martinez, A. T.; Alcalde, M. Biochem J 2012, 441, 487-498. Gonzalez-Perez, D.; Garcia-Ruiz, E.; Ruiz-Duenas, F. J.; Martinez, A. T.; Alcalde, M. ACS Catalysis 2014, 4, 3891-3901. Gonzalez-Perez, D.; Mateljak, I.; Garcia-Ruiz, E.; Ruiz-Duenas, F. J.; Martinez, A. T.; Alcalde, M. Cat Sci & Technol 2016, 6, 6625-6636. Salazar, O.; Sun, S. In Directed enzyme evolution: screening and selection methods; Arnold, F. H., Georgiou, G., Eds.; Humana Press: Totowa, New Jersey, 2003; Vol. 230, p 85-97. Brissos, V.; Ferreira, M.; Grass, G.; Martins, L. O. ACS Catalysis 2015, 5, 4932-4941. Rosado, T.; Bernardo, P.; Koci, K.; Coelho, A. V.; Robalo, M. P.; Martins, L. O. Bioresource Technol 2012, 124, 371-378. Sousa, A. C.; Oliveira, M. C.; Martins, L. O.; Robalo, M. P. Green Chem. 2014, 16, 41274136. Kelley, L. A.; Mezulis, S.; Yates, C. M.; Wass, M. N.; Sternberg, M. J. Nature Protoc 2015, 10, 845-858. Koduri, R. S.; Tien, M. J Biol Chem 1995, 270, 22254-22258. Gelpke, M. D.; moenne-Loccoz, P.; Gold, M. H. Biochemistry 1999, 38, 11482-11489. Soskine, M.; Tawfik, D. S. Nature Rev Genetics 2010, 11, 572-582. Miton, C. M.; Tokuriki, N. Protein Sci 2016, 25, 1260-1272. Miyazaki, C.; Takahashi, H. FEBS Lett 2001, 509, 111-114. Ogola, H. J. O.; Hashimoto, N.; Miyabe, S.; Ashida, H.; Ishikawa, T.; Shibata, H.; Sawa, Y. Appl Microbiol Biotech 2010, 87, 1727-1736. Ryan, B. J.; O'Fagain, C. Biochimie 2007, 89, 1029-1032. Valderrama, B.; Garcia-Arellano, H.; Giansanti, S.; Baratto, M. C.; Pogni, R.; VazquezDuhalt, R. FASEB J 2006, 20, 1233-+. 17


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(61) (62) (63) (64) (65) (66) (67) (68) (69) (70) (71) (72) (73) (74) (75) (76) (77) (78) (79)

Morawski, B.; Quan, S.; Arnold, F. H. Biotechnol Bioeng 2001, 76, 99-107. Sousa, A. C.; Martins, L. O.; Robalo, M. P. Adv Synth Catal 2013, 355, 2908-2917. Sousa, A. C.; Oliveira, M. C.; Martins, L. O.; Robalo, M. P. Green Chem 2014, 16, 41274136. Rahmanpour, R.; Bugg, T. D. Arch Biochem Biophys 2015. Caramelo, L.; Martinez, M. J.; Martinez, A. T. Appl Environ Microb 1999, 65, 916-922. Loncar, N.; Colpa, D. I.; Fraaije, M. W. Tetrahedron 2016. Wong, D. W. Appl Biochem Biotechnol 2009, 157, 174-209. Poulos, T. L.; Kraut, J. J Biol Chem 1980, 255, 8199-8205. Balny, C.; Travers, F.; Barman, T.; Douzou, P. Eur Phys J 1987, 14, 375-383. Baek, H. K.; Van Wart, H. E. J Am Chem Soc 1992, 114, 718-725. Denisov, I. G.; Makris, T. M.; Sligar, S. G. J Biol Chem 2002, 277, 42706-42710. Tanaka, M.; Matsuura, K.; Yoshioka, S.; Takahashi, S.; Ishimori, K.; Hori, H.; Morishima, I. Biophys J 2003, 84, 1998-2004. Rodriguez-Lopez, J. N.; Smith, A. T.; Thorneley, R. N. J Biol Inorg Chem 1996, 1, 136-142. Rodriguez-Lopez, J. N.; Smith, A. T.; Thorneley, R. N. J Biol Chem 1996, 271, 4023-4030. Mak, P. J.; Thammawichai, W.; Wiedenhoeft, D.; Kincaid, J. R. J Am Chem Soc 2015, 137, 349-361. Wariishi, H.; Dunford, H. B.; MacDonald, I. D.; Gold, M. H. J Biol Chem 1989, 264, 33353340. Ruiz-Duenas, F. J.; Morales, M.; Perez-Boada, M.; Choinowski, T.; Martinez, M. J.; Piontek, K.; Martinez, A. T. Biochemistry 2007, 46, 66-77. Banci, L.; Ciofi-Baffoni, S.; Tien, M. Biochemistry 1999, 38, 3205-3210. Wariishi, H.; Valli, K.; Gold, M. H. J Biol Chem 1992, 267, 23688-23695.

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ACS Catalysis

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

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FIGURE LEGENDS Figure 1 – Lineage of PpDyP variants generated in this study. Variants with higher activity for ABTS are shown in light grey and variants with higher activity for DMP are in white. nd - not detected. iA – initial activity. Figure 2 – Map of mutations (in green) present in the 6E10 variant using the PpDyP model structure.38,52 (A) Overall fold of PpDyP showing the typical ferredoxin-like fold with the heme cofactor sandwiched between distal and proximal sides. Detail view of the PpDyP active site; the conserved proximal axial H197 residue and the distal D132, N136 and R214 residues are shown in cyan.38 The E188K mutation is located in the surface loop I179-A196, at ∼ 5Å of the heme propionate. The A142V mutation is in a small α-helix E140-A146, in close contact with loop D117-D139 (in red) that contains the distal residues N136 and D132, and mutation H125Y. (B) Heme access channel represented as electrostatic surface (stick model). (C) Relative position of T134, T138 and M212 putative H2O2-sensitive residues (in yellow); their distances to the A142V mutation are 5 (T138), 8 (M212) and 11 Å (T134). Figure 3 – HPLC traces, monitored at 280 nm, showing the oxidation of 2.5 mM of guaiacylglycerol-β-guaiacyl ether (GGE) using 10 U of 6E10 (A) or wild-type (B) in the presence of 2.5 mM H2O2, in sodium phosphate buffer pH 7.5 and pH 5, respectively. (C) Analysis of products by liquid HPLC-mass spectrometry in positive mode, show m/z 661 [M+Na]+ and 635 [M+H]+, consistent with molecular formulas C34H38O12 and C34H34O12, respectively. Figure 4 – Stopped-flow analysis of the reaction of PpDyP 6E10 in sodium phosphate buffer, pH 8.5 at 25°C. (A) 6E10 (2 µM) was mixed with ~2.5 equivalents of H2O2. The time-course spectra reveal the formation of an intermediate with the characteristics of Compound I. The inset shows the enlarge region between 450 and 600 nm. (B) Reaction of 6E10 with H2O2. Compound I formation as a function of time was monitored at 404 nm and the rates were obtained from single exponential fits. In (C) Reactions of Compound I with guaiacol and (D) Reactions of Compound II with guaiacol. One syringe contained 6E10 mixed with H2O2 (1:1) and the second contained guaiacol. Compound II formation (C) and Compound II decay (D) were monitored at 450 nm, the isosbestic of resting enzyme and Cpd I, and at 415 nm, the isosbestic of Cpd I and Cpd II, respectively (Figure. S8). The traces were exponential in character. Figure 5– Schematic representation of the catalytic cycle of 6E10 at pH 8.5. The resting state enzyme, E, Compound I (Cpd I), Compound II (Cpd II) were spectroscopically identified; the presence of the intermediate E-H2O2 is inferred from kinetic analysis. AH · represents the reducing substrate and A indicates the formed radical product.

19


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1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

ACS Catalysis

Table 1- PpDyP wild-type and variants production levels, spectroscopic and redox properties. Enzyme production (mg·L-1)

Rz

λmax (nm)

wild-type

6

1.9

9F6

7

17F11

(mM-1 cm-1)

Heme content

E0´ (mV)

402

40

1.0 ± 0.1

-260 ± 10a

1.9

400

59

0.9 ± 0.1

-110 ± 10

7

1.3

401

57

0.9 ± 0.1

-105 ± 10

6E10

12

1.6

400

58

0.9 ± 0.1

-60 ± 10

A142V

7

1.8

405

94

0.9 ± 0.1

ND

31F3

12

2.0

405

49

0.9 ± 0.1

ND

21G11

15

1.4

405

35

0.8 ± 0.1

ND

ε

a 36

ND – not determined

20


ACS Catalysis

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Table 2 – Apparent steady-state kinetic parameters for reduction of H2O2, using ABTS as reducing substrate, and oxidation of ABTS of PpDyP wild-type and variants, in the presence of 0.5 mM H2O2, at 25°C and the optimal pH of the enzymes. H2O2

ABTS

mutations

pH

kcat (s-1)

Km (µM)

kcat/Km (M-1s-1)

Ki (mM)

Km (µM)

kcat/Km (M-1s-1)

WT

-

4.3

23 ± 2

60 ± 10

(3.8 ± 0.5) × 105

0.7 ± 0.1

2500 ± 300

(0.9 ± 0.1) × 104

9F6

E188K

4.3

128 ± 6

200 ± 60

(6.4 ± 0.9) × 105

0.8 ± 0.2

220 ± 60

(6.0 ± 0.6) × 105

17F11

E188K/A142V

5.5

15 ± 3

160 ± 60

(0.9 ± 0.3) × 105

2.3 ±1.1

1000 ± 300

(1.5 ± 0.4) × 104

6E10

E188K/A142V/H125Y

5.5

15 ± 1

44 ± 5

(3.4 ± 0.3) × 105

nd

1100 ± 200

(1.4 ± 0.1) × 104

A142V

5.5

4.4 ± 0.6

47 ± 9

(0.9 ± 0.2) × 105

nd

260 ± 40

(1.7 ± 0.3) × 104

H125R

4.3

26 ± 2

220 ± 40

(1.2 ± 0.1) × 105

5±1

500 ± 100

(5.2 ± 0.7) × 104

4.3

69 ± 7

200 ± 70

(3.5 ± 0.9) × 105

0.8 ± 0.3

280 ± 50

(2.5 ± 0.3) × 105

31F3

21G11 E188K/H125R nd - not detected

21


Page 23 of 30

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

ACS Catalysis

Table 3 – Apparent steady-state kinetic parameters for oxidation of DMP of PpDyP wild-type and variants, in the presence of 0.5 mM H2O2, at 25°C and the optimal pH of the enzymes. DMP mutations

pH

kcat (s-1)

Km (µM)

kcat/Km (M-1s-1)

WT

-

4.3

0.05 ± 0.01

70 ± 20

(0.7 ± 0.2) × 103

9F6

E188K

4.3

0.09 ± 0.01

60 ± 20

(1.5 ± 0.3) × 103

17F11

E188K/A142V

8.5

3.6 ± 0.1

42 ± 4

(0.9 ± 0.1) × 105

6E10

E188K/A142V/H125Y

8.5

6±2

58 ± 6

(1.1 ± 0.4) × 105

A142V

8.5

1.8 ± 0.2

250 ± 40

(7.2 ± 0.9) × 103

31F3

H125R

4.3

0.03 ± 0.003

80 ± 20

(0.4 ± 0.1) × 103

21G11

E188K/H125R

4.3

0.02 ± 0.002

23 ± 6

(0.6 ± 0.1) × 103

22


ACS Catalysis

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Page 24 of 30

Table 4 – Catalytic efficiencies of wild-type and 6E10 variant for various lignin-related phenolic compounds, Kraft lignin (in the absence and presence of Mn2+ or methyl syringate) and aromatic amines at 25°C and at the optimal pH of the reaction; 4-ADA, 4-aminodiphenylamine, 2,5-DABSA, 2,5-diaminobenzenesulfonic acid, 1,2 PDA, 1,2phenylenediamine and 1,4-PDA, 1,4-phenylenediamine.

Wild-type Phenolics Guaiacol Syringaldehyde Acetosyringone Methyl syringate Kraft lignin + Mn2+ + Methyl syringate Aromatic 4-ADA amines 2,5-DABSA 1,2-PDA 1,4-PDA

pH 4.3 4.3 4.3 4.3 4.3 4.3 4.3 5 5 5 5

6E10 -1

-1

kcat/Km (s ·M ) 1.2 × 103 1.2 × 103 2.6 × 103 1.6 × 102 1.6 × 103 2.1 × 103 1.1 × 104 2.2 × 105 2.9 × 103 4.5 × 102 2.5 × 103

pH 8.5 8.5 8.5 8 8.5 8.5 8.5 5 8 8 8

kcat/Km (s-1·M-1) 2.2× 104 3.3 × 104 5.0 × 103 1.4 × 104 4.1 × 103 4.6 × 103 1.2 × 104 4.2 × 105 3.8 × 103 1.1 × 103 1.8 × 104

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Page 25 of 30

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ACS Catalysis

Table 5– ESI-MS of the products obtained in the enzymatic oxidation of the aromatic amines with 1 U·mL-1 of wild-type PpDyP, in 50 mM acetate buffer, pH 4.5, with 0.2 mM of H2O2. Aromatic amines

Products

m/z

Ref.

547 [M+H]+

62

319 [M+H]+

62

211 [M+H]+

63

369 [M-H]-

63

4-aminodiphenylamine

C36H30N6 (MW = 546.66 g·mol-1) 1,4-phenylenediamine

C18H18N6 (MW = 318.38 g·mol-1) 1,2-phenylenediamine

C12H10N4 (MW = 210.23 g·mol-1) 2,5-diaminobenzene sulphonic acid

C12H10N4O6S2 (MW = 370.36 g·mol-1)

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ACS Catalysis

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Figure 1

3rd iAABTS= 15× iADMP= 11×

2nd iAABTS= 7× iADMP= 3× 1st iAABTS= 4× iADMP= r

6E10 E188K A142V H125Y

17F11

21G11

E188K A142V

E188K H125R

9F6 E188K

2nd iAABTS= 23× iADMP= 1.5×

31F3 H125R

1st iAABTS= 3× iADMP= nd

Wild-type -type Figure 1 – Lineage of PpDyP variants generated in this study. Variants with higher activity for ABTS are shown in light grey and variants with higher activity for DMP are in white. nd - not detected. iA - initial activity.

25


Page 27 of 30

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

ACS Catalysis

Figure 2

Figure 2 – Map of mutations (in green) present in the 6E10 variant using the PpDyP model structure. (A) Overall fold of PpDyP shows the typical ferredoxin-like fold with the heme cofactor sandwiched between distal and proximal sides. Detail view of the PpDyP active site; the conserved proximal axial H197 residue and the distal D132, N136 and R214 residues are shown in cyan.38 The E188K mutation is located in the surface loop I179-A196, at the entry to the heme channel at ∼ 5Å of the heme propionate. The A142V mutation is in a small α-helix E140-A146, in close contact with loop D117D139 (in red) that contains the distal residues N136 and D132, and mutation H125Y. (B) Heme access channel represented as electrostatic surface (stick model). (C) Relative position of T134, T138 and M212 putative H2O2-sensitive residues (in yellow); their distances to the A142V mutation are 5 (T138), 8 (M212) and 11 Å (T134).

26


ACS Catalysis

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 28 of 30

Figure 3

A

B

t96h t48h

t0 4

5

C

6

7 8 9 Minutes

t1h

t96h t48h

t24h

t0

10

4

6

5

7 8 9 Minutes

t1h

t24h

10

OH H3CO

HO

OH O

m/z 343 [M+Na]+

H3CO

m/z 661 [M+Na]+

OCH3 HO OH H3CO

OH

O O HO

OCH3

HO OH

m/z 635 [M+H]+

OCH3

H3CO

O OH H CO 3

OH

O O HO

OCH3

HO O

(8 )

H CO 3

1h time 0 4

5

6

7 8 Time (min)

9

10

Figure 3 – HPLC traces, monitored at 280 nm, showing the oxidation of 2.5 mM of guaiacylglycerolβ-guaiacyl ether (GGE) using 10 U of 6E10 (A) or wild-type (B) in the presence of 2.5 mM H2O2, in sodium phosphate buffer pH 7.5 and pH 5, respectively. (C) Analysis of products by liquid HPLCmass spectrometry in positive mode, show m/z 661 [M+Na]+ and 635 [M+H]+, consistent with molecular formulas C34H38O12 and C34H34O12, respectively. 27


Page 29 of 30

Figure 4

20

80 k1obs (s-1)

30

B

100

k1obs (s-1)

ε (mM-1 cm-1)

120

0.00075 s 0.015 s 0.03 s 0.05 s 0.075 s 0.1 s 0.4 s

A

40

60 40

10

450

550

650

20

60 50 40 30 20 10 0

0

350

450

550

0

650

10

20

30

[H2O2] (µ µM)

0

0

50

100

Wavelength (nm)

150

200

[H2O2] (µ µM) 0.8

C

6

D k3obs (s-1)

0.6

k2obs (s-1)

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

ACS Catalysis

4

2

0.4

0.2

0

0 0

2

4

6

8

10

0

1

Guaiacol (µ µM)

2

3

4

5

6

Guaiacol (mM)

Figure 4 – Stopped-flow analysis of the reaction of PpDyP 6E10 in sodium phosphate buffer, pH 8.5 at 25°C. (A) 6E10 (2 µM) was mixed with ~2.5 equivalents of H2O2. The time-course spectra reveal the formation of an intermediate with the characteristics of Compound I. The inset shows the enlarge region between 450 and 600 nm. (B) Reaction of 6E10 with H2O2. Compound I formation as a function of time was monitored at 404 nm and the rates were obtained from single exponential fits. In (C) Reactions of Compound I with guaiacol and (D) Reactions of Compound II with guaiacol. One syringe contained 6E10 mixed with H2O2 (1:1) and the second contained guaiacol. Compound II formation (C) and Compound II decay (D) were monitored at 450 nm, the isosbestic of resting enzyme and Cpd I, and at 415 nm, the isosbestic of Cpd I and Cpd II, respectively (Figure. S8). The traces were exponential in character.

28


ACS Catalysis

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Page 30 of 30

Figure 5

H2O2

K1 (44 ± 3) × 10-6 M

E A AH

E - H2O2

k1 133 ± 4 s-1 H2O

Cpd I

Cpd II A

AH

k´2 (6.1 ± 0.3) × 105 M-1 s-1

Figure 5– Schematic representation of the catalytic cycle of 6E10 at pH 8.5. The resting state enzyme, E, Compound I (Cpd I), Compound II (Cpd II) were spectroscopically identified; the presence of the intermediate E-H2O2 is inferred from kinetic analysis. AH represents the reducing substrate and A· indicates the formed radical product.

29


Engineering a Bacterial DyP-Type Peroxidase for Enhanced Oxidation

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