pH-Dependence of Manganese (II) Oxidation Reaction by Novel Wild-Type and Mutants Recombinant <i>Phlebia radiata</i> Manganese Peroxidase 3 (rPr-MnP3) Enzymes

The goal of this study was to determine whether mutation of the Mn-binding site of wild-type recombinant Phlebia radiata manganese peroxidase 3 affected the pH-dependence kinetic parameters. pH range investigated was 2.5 – 12.0. The catalytic efficiency of the mutant enzymes at high and low pH in comparison to the wild-type was investigated using standard rPr-MnP3 protocol. Wild-type recombinant Phlebia radiata MnP3 enzyme showed optimal activity with Mn (II) as substrate at pH 5.0 and remained moderately active (approximately 40%) in the pH range of 6.0 - 9.0. The rPr-MnP3 mutants’ maximum activity ranged between 5.5 and 8.0. Wild-type and mutants rPr-MnP3 enzymes exhibited a similar pH profile with optimum pH of 3.0 for ABTS oxidation. Mutation has severely decreased the catalytic efficiency for Mn (II) oxidation at pH 5.0. The rPr-MnP3 enzymes showed enhanced affinity for Mn (II) at alkaline pH and a more alkaline range for catalysis than ever reported for any Manganese Peroxidase. This study reveals that at higher pH, rPr-MnP3 can function with alternative ligands in the Mn (II) site and does not have an absolutely obligate requirement for an all carboxylate ligand set. These results further strongly confirm that Mn ²⁺ binding site is the only productive catalytic site for Mn (II) oxidation.

• pH-Dependence
• <i>Phlebia radiata</i>
• Manganese Peroxidase
• Wild-Type
• Mutants
• Recombinant Enzyme

1. Introduction

Manganese peroxidase (MnP) (EC 1.11.1.13) is a class II extracellular fungal haem-containing peroxidase belonging to the plant peroxidase-like protein superfamily [1] [2]. Manganese peroxidase is still considered the most ubiquitous ligninolytic enzyme among white-rot fungi and was first discovered in Phanerochaetechrysosporium [3] [4] and thereafter in most of the wood lignin and litter-decaying basidiomycetes [5]. MnP has been isolated or expressed, purified and characterized from various white rot fungi [6] - [11].

In plants, MnP, a component of ligninolytic enzymes complex catalyzes lignin de-polymerization. The enzyme has been found to catalyze the breakdown of lignin and phenolic lignin model compounds, as well as the non-phenolic ones in the presence of certain compounds to polycyclic aromatic hydrocarbons by oxidation of Mn²⁺ to Mn³⁺ with H₂O₂ as an oxidant [1] [12]. Manganese peroxidase remains one of the most common lignin degradation enzymes and has great potential in the field of agriculture for degradation of some cellulose, hemicellulose, lignin, etc. In the environment, this enzyme has been widely employed in the degradation of some recalcitrant organic pollutants such as industrial dyes, polycyclic aromatic hydrocarbons, chlorophenols; and nitroaromatic compounds, known to be very harmful to human health [13]. Several researches have suggested that some azo dyes could be efficiently degraded by the purified MnPs, which were isolated from P.chrysosporium,Lentinula edodes,Trametes versicolor,Dichomitus squalens,Stereum ostrea and Irpex lacteus [13] - [18].

The key physiological role of MnP is the oxidation of Mn²⁺ to Mn³⁺ ions [4] [19] [20]. The MnP oxidizes the one-electron donor Mn²⁺ to Mn³⁺. Manganese (III) ion manufactured enzymatically is assumed to form a diffusible oxidant complex with dicarboxylic acid chelators such as oxalate which is also secreted by the fungus [20] [21] [22] [23]. The Mn³⁺ organic acid complex then oxidizes phenolic substrates, including lignin substructure model compounds [24] and aromatic pollutants [4] [25] [26] [27] as well as possible mediator molecules [26] [27].

A unique characteristic of MnP is that the best reducing substrate for Compounds I and II is Mn (II), a metal ion naturally present in wood. However, compound II of MnP exhibits an absolute requirement for Mn (II) as an exclusive reductant essential for the completion of the catalytic cycle of the enzyme [22] [28]. Only Mn (II) is capable of very rapidly reducing MnP compound II to its original state, thereby allowing the enzyme to initiate new substrate oxidations [22] [29] [30]. Not only is Mn essential for the activity of MnP [31], but it also regulates the production of MnP [32] [33], laccase and Lignin peroxidase (LiP) in P.chrysosporium and in other white rot fungi [34] ).

Ionic strength and pH are very important factors affecting enzyme activity. The protein surface is the point of contact with the solvent and media, thus the physical and chemical environment are sensed through the surface residues. However, changes in the ionization state of the polar residues of protein may interfere with enzymatic activity.

2, 2’-azino-bis (3-ethylbenz-thiazoline-6-sulfonic acid) (ABTS) is a well-known substrate for peroxidases [35] and has been shown to be rapidly oxidised by MnPs [36] [37] [38]. The chemical properties of ABTS and its oxidation by peroxidases, particularly HRPC; have been intensively studied [39] [40]. 2, 2’-azino-bis (3-ethylbenz-thiazoline-6-sulfonic acid) (ABTS) undergoes a single electron oxidation in the presence of peroxidase and hydrogen peroxide to produce a long-lived cation radical. This radical product has an intense blue/green colour and thus its accumulation can be measured spectroscopically [35]. Note that MnP does not require a functional Mn (II) oxidation site for ABTS to be a good substrate as its oxidation is more non-specific in nature.

The functions and applications of MnP enzyme stimulate interest in studying and understanding enzyme structure, biochemical characteristics, and genes. Therefore, searching for new MnP that is highly effective from widely distributed sources worldwide is crucial for its application in agricultural productions, medical, industrial and environmental protection. Recently, more and more attention has been paid to the value of bioremediation of this enzyme. The focus of this study was to determine the optimum conditions of pH to maintain recombinant Phlebiaradiata MnP3 activities. Furthermore, the work also aimed at investigating if mutation of the Mn-binding site of wild-type recombinant Phlebiaradiata manganese peroxidase 3 enzyme has affected the pH dependence kinetic parameters and to also ascertain the effectiveness of the mutant enzymes at high and low pH in comparison to the wild-type. This involved heterologous expression and purification of recombinant MnP3 from Phlebiaradiata, followed by Manganese dependent peroxidase assays using the required substrates and reagents.

2. Materials and Methods

2.1. Materials

The complete MnP3 gene of Phlebia radiata strain 79 (ATCC 64658) was provided by Dr. Taina Lundell, Department of Food and Environmental Sciences, Division of Microbiology, University of Helsinki, Finland. The Gene Bank accession number for the cDNA encoding peroxidase Pr-MnP3 is AJ566200. The Pr-MnP3 cDNA was present in vector pCR2.1.TOPO. The Escherichia coli expression vector pFLAG1 was obtained from International Biotechnologies Inc, UK. Peroxidases, wild-type, E40H, E44H, E40H/E44H, DI86H and 186N (RZ = 5.6, 2.1, 4.5, 5.2, 4.4 and 5.0, respectively) were produced, activated and purified based on the procedure previously described [41]. All chemicals used in this study were obtained from Sigma–Aldrich, UK and Fisher Scientific, UK. Restriction enzymes were supplied by NEBiolabs, UK. All spectroscopic measurements in this study were carried out using UV/Vis spectrophotometer (UV-2401 PC, Shimadzu Scientific Instruments, Addison, IL).

2.2. Steady-State Kinetic Analysis of rPr-MnP3 Activity with Manganese (II)

Mn (II) is assumed to be the in vivo substrate for manganese peroxidase. A set of apparent steady-state kinetic constants for Mn (II) to Mn (III) oxidation for each of the engineered Phlebia radiata MnP3 enzymes was obtained by measuring the initial rates of assays at 238 nm for varying MnSO₄ concentrations: 0.02 - 1.0 mM for wild-type, D186H and D186N mutants and 1 - 35 mM for E40H, E44H, and E40H/E44H mutants. The concentration of MnSO₄ was increased for E40H, E44H and E40H/E44H enzymes since the concentration used for the wild-type, D186H and D186N could not produce measurable activity. Enzyme concentrations of 0.8 nM and 0.2 μM and the assay buffer was 100 mM Na-tartrate, pH 5.0 and 8.0 for wild-type and mutants, respectively.

All assays were performed with a fixed 0.1 mM hydrogen peroxide at 25˚C using ultraviolet spectrophotometer (UV-2401 PC, Shimadzu Scientific Instruments, Addison, IL, USA). In this study, the product being detected was actually a Mn³⁺-tartrate complex with ε238 = 6.5 mM⁻¹ cm⁻¹ [41]. To ensure quality control, best laboratory practices were adopted and all measurements were carried out in triplicate.

2.3. Steady-State Kinetic Analysis of Recombinant Phlebiaradiata Manganese Peroxidase 3(rPr-MnP3) Activity with ABTS

2, 2’-azino-bis (3-ethylbenzthiozoline-6-sulphonic acid) (ABTS) is a commonly used peroxidase substrate with very high levels of activity as seen in plant peroxidases [35]. 2, 2’-azino-bis (3-ethylbenzthiozoline-6-sulphonic acid) assays for wild-type rPr-MnP3 were carried out in 100 mM Na-tartrate buffer, pH 3.0, with a fixed H₂O₂ concentration (0.1 mM), enzyme concentration (0.01 uM) and the ABTS final concentration of 3.5 mM for the wild-type enzyme while ABTS concentration range 1 - 6.5 mM was used for the mutants variants. Formation of the radical product as a function of time was measured at 25˚C by monitoring the increase in absorbance at 414 nm. The initial rates were determined using ε414 = 36.8 mM⁻¹·cm⁻¹ [34] and all ABTS assays were in triplicate.

2.4. Determination of the pH Optima for Mn 2+ and ABTS Oxidation by rP.radiata MnP3 Enzymes

The optimum pH values for the oxidation of Manganese (II) and 2, 2’-azino-bis (3-ethylbenz-thiazoline-6-sulfonic acid) (ABTS) by wild-type rPr-MnP3 were determined by assaying enzyme activity in 100 mM tartrate buffers of varying pH. For Mn (II) oxidation, assay mixtures contained 1 mM MnSO₄, 0.1 mM H₂O₂, and 0.023 µM enzyme and a pH range from 3.5 to 10.0 was used. In ABTS oxidation, the assay mixtures were 1.6 mM ABTS, 0.1 mM H₂O₂, and 0.006 µM enzyme with a pH range of 3.0 to 5.5.

Similarly, the pH optima for all rPr-MnP3 mutants were also determined for Mn (II) and ABTS as substrates. In the Mn (II) oxidation, the assay mixtures were enzyme concentration of 0.4 µM E40H/E44H, 0.34 µM E44H, 0.3 µM E40H, 0.3 µM D186H and 0.21 µM D186N, along with 0.1 mM hydrogen peroxide, 25 mM MnSO₄, and the pH ranged from 3.5 to 12.0. For ABTS oxidation by rPr-MnP3 mutants, the assay mixtures were 0.0022 µM E40H/E44H, 0.0094 µM E44H, 0.0015 µM E40H, 0.0096 µM D186H and 0.0086 µM D186N, in addition to 1.6 mM ABTS, 0.1 mM H₂O₂ and the pH ranged from 2.5 to 5.5. The total volume of all assay mixtures was 1 ml and assays were performed at 25˚C. The reaction was initiated by the addition of hydrogen peroxide and the increase in absorbance at 238 nm or 414 nm, for Mn (II) and ABTS respectively, was measured using a UV/Vis spectrophotometer (Shimadzu UV-2401 PC). All measurements were in triplicate. To determine a pKa for Mn (II) oxidation, the mean rates for the various pHs were calculated; from the data log K (observed) was calculated and plotted against pH. The pKawas then extrapolated from each plot.

Data analysis, plotting and manipulations were carried out using the solver tools of Microsoft Excel and the statistical analysis software, SigmaPlot for Windows V4.01 (SPSS UK Ltd, Woking, UK). The Km (Michaelis-Menten constant) and Kcat obtained were used to calculate the specificity constants (kcat/Km). In addition, an activity assay was further conducted at pH 8.0 using the above steady-state conditions to see how rPr-MnP3 enzymes will behave catalytically at alkaline pH.

3. Results

3.1. pH Optimum for Mn (II) Oxidation by Wild-Type and Mutant rPr-MnP3 Enzymes

Figure 1 and Table 1 show the pH optima data for Mn (II) substrate oxidation by rPr-MnP3 enzymes. A sharp acidic pH optimum was observed in the wild-type rPr-MnP3 activity. As shown above, all mutants retained significant residual activity at pH values > 5.5 (up to 35% of wild-type).

Table 1

pH optima and apparent pKa’s for Mn (II) oxidation by wild-type and mutants rPr. radiata MnP3 enzymes. The assays were conducted at 25˚C

Enzyme	pH optima	Apparent pKa
Wild-type	~5.0/9.0	~4.5; 5.5; 6.5; ~9.0
E40H	6.0 - 8.0	~5.5
E44H	6.0 - 8.0	~5.5
E40H/E44H	7.5 - 10.0	~7.0
D186H	6.5 - 8.0	~6.5
D186N	5.5 - 12	~5.5

In Figure 2(a) the wild-type rPr-MnP3 pKa reflected at four points labeled 1, 2, 3, and 4. As presented in Table 1, the wild-type enzyme had its pKa at ~4.5, 5.5, ~6.5 and ~9.0. The variants E40H, E44H and D186N had their respective pKas at ~5.5 while that of the D186H mutant and E40H/E44H double variant were 6.5 and ~7.0 respectively.

3.2. pH Optimum for ABTS Oxidation by Wild-Type and Mutant rPr-MnP3 Enzymes

As shown in the Figure 3 below, both the wild-type and the rPr-MnP3 mutant enzymes, E40H, E44H, E40H/E44H, D186H and D186N had their optimum pH at pH 3.0.

3.3. Mn (II) Oxidation by rPr-MnP3 Wild-Type and Mutant Variants at pH 5.0 and 8.0

The results of Mn (II) oxidation by rPr-MnP3 wild-type and mutant variants at pH 5.0 and 8.0 are shown in Figure 4(a) and Figure 4(b) for Mn (II) oxidation by rPr-MnP3 mutants. The kinetic parameters for Mn (II) oxidation by these enzymes are presented in Table 2.

4. Discussion

4.1. Determination of pH Optimum and Apparent pKa for Mn (II) Oxidation by Wild-Type and Mutant Recombinant Phlebiaradiata MnP3 Enzymes

The optimal conditions for the catalysis by different peroxidases are not identical [42]. Thus, the optimum pH for the oxidation of Mn (II) and ABTS by recombinant P.radiata MnP3 enzyme was determined by assaying enzyme activity in 100 mM tartrate buffers of varying pH ranges (from 3.5 to 12.0). Figure 1 and Table 1 showed the pH optima data for Mn (II) substrate oxidation by rPr-MnP3. pH is a determining factor in the expression of enzymatic activity as it changes the ionization states of amino acid side chains or ionization of the substrate [43]. The active site of enzymes is frequently composed of ionized groups and that must be in the proper ionic form in order to maintain the conformation of the active site. It is worthy of note that the behaviour of the enzyme

activity at different pH values gives information on the identities of the phototropic groups at the active site [44].

A sharp acidic pH optimum for wild-type rPr-MnP3 activity was observed in this study and the highest wild-type rPr-MnP3 turnover number was found to be at pH 5.0, while the enzyme was very active at the pH range of 4.5 to 5.5. Though the enzyme has it pH optimum in the acidic region, it was observed to be moderately active at alkaline pH, showing similar activity of approximately 18% of the maximum in the pH range of 6.5 to 8.5 with a sharp decrease towards no activity at pH 10.0. Wild-typerPr-radiata MnP3 with a pH optimum at 5.0 is consistent with the literature [45] [46]. The optimum pH for Mn (II) oxidation by MnP from P.chrysosporium is reported to be 4.5 [47] while those of other MnPs, P.sordid [20], D.squaleus [48], and L.edodes [49] are also in the acidic range. All mutant variants had substantially decreased activity at low pH, consistent with the loss reported to be 4.5 of one or more Mn ligands at the interaction site, but all retained significant residual activity at pH values > 5.5 (up to 35% of wild-type). The variants broadly fell into three groups (Figure 1 and Table 1): those that were maximally active at pH 6.0 (E40H, E44H, E40H/E44H), D186H (pH values ≥ 6.5) and D186N (pH values > 7.5). It is important to note that several variants actually have a higher activity than wild-type at pH ~10.0 and approximately 40% of the wild-type activity in the mid alkaline range.

A plot of log k (observed) against pH (Figure 1) was used to determine the apparent pKa for Mn (II) dependent oxidation. Figure 1 shows that the wild-type enzyme behaviour is complex and may involve as many as four critical

ionizations, labelled 1,2,3,4 in Figure 2(a). These presumably reflect deprotonation of one or more carboxylate ligands at the Mn site, starting at pKa ~4.5, and followed at higher pH by pKa’s at ~5.5, ~6.5 and ~9.0 respectively. The origins of these are unclear at present, although the complex dissociation and ionization equilibrium of the Mn tartrate complexes relevant as substrates are being considered. An acidic pKa at ~5.5 is conserved in the E40H, E44H and D186N mutants, but is shifted to 6.5 in the D186H mutant, implying that for this variant deprotonation of the new H186 residue is essential to allow Mn oxidation at

Table 2

A table showing kinetic parameters during the pH dependence of Mn (II) oxidation by wild-type and mutants recombinant Phlebiaradiata manganese peroxidase 3 at pH 5.0 and 8.0

Enzymes	pH	Km (mM)	Kcat (s−1)	Kcat/Km (s−1·mM−1)	pH	Km (mM)	Kcat (s−1)	Kcat/Km (s−1·mM−1)
MnP III (WT)	5.0	0.17 ± 10	175 ± 3.0	1029.41	8.0	0.011 ± 0.0001	34.2 ± 0.8	3109.1
E40H	5.0	11.1 ± 0.4	12.0 ± 0.2	1.01	8.0	0.222 ± 0.03	20.5 ± 0.7	92.3
E44H	5.0	8.5 ± 0.5	12.1 ± 0.3	1.49	8.0	0.124 ± 0.01	30.2 ± 0.4	243.6
E40H/E44H	5.0	19.9 ± 2.5	1. 7 ± 0.1	0.08	8.0	1.086 ± 0.10	31.7 ± 1.5	29.2
D189H	5.0	23.1 ± 3.9	0.84 ± 0.1	0.04	8.0	0.42 ± 0.05	53.0 ± 1.2	128.0
D189N	5.0	30.3 ± 3.7	8.0 ± 0.5	0.30	8.0	0.33 ± 0.02	5.0 ± 0.1	15.2

A data point represents the mean of three independent determinations with standard errors indicated. Assays were carried out using standard rPr-MnP3 protocol as described in Section 2.2. Data were fitted to the Michaelis-Menten equation using SigmaPlot8.0.

higher pH. Similarly the apparent pKa for the E40H/E44H double variant is shifted to ~7.0 perhaps because both His residues must be deprotonated to allow activity. Mn oxidation in mid-alkaline range with up to two of the classical carboxylate ligands replaced is still quite significant (~40% of wild-type). Moreover, the results for D186N and D186H variants imply that histidine is better tolerated in this respect than asparagine. The loss of carboxylate ligands clearly impacts the low pH end of the activity profile more than the alkaline end. It would appear that at higher pH Mn oxidation is less dependent on the precise ionization of the Mn binding site, particularly if histidine is the substituting residue.

This discovery that P.radiata wild-type recombinant Pr-MnP3 enzyme is able to oxidize Mn (II) under both acidic (faster) and alkaline (slower) conditions, and the relatively good performance of the E40H/E44H and D186H variants at alkaline pH values, is particularly interesting in the light of potential industrial applications. Pulping and pulp bleaching are routinely performed under alkaline conditions and thus any enzyme used in the pulp and paper industry is required to react under alkaline conditions. Most MnP’s from wood-colonizing white-rot fungi have been found to have highly acidic pH optima [50], so in previous studies on lignin degrading enzymes [47] [51] [52], the enzymes tried were active only under acidic pH conditions. Wild-type recombinant P.radiata MnP3 enzyme being moderately active at alkaline pH range therefore makes it unique in comparison with other MnP’s of white-rot fungi studied to date. The ability of recombinant MnP3 enzymes from P.radiata to be active under alkaline conditions might make them useful in the pulp and paper industry for treatment of alkaline black liquor and bleaching effluent and/or to bleach the pulp previously cooked under alkaline conditions.

4.2. Determination of pH Optimum ABTS Oxidation by Wild-Type and Mutant Recombinant Phlebiaradiata MnP3 Enzymes

Figure 4 illustrates the pH dependence of ABTS turnover for the wild-type and mutants (E40H, E44H, E40H/E44H, D186H and D186N) rPr-MnP3 enzymes. As observed, all the six enzymes showed a strong acidic pH preference. The activity for ABTS oxidation by wild-type rPr-MnP3 was highest at pH 3.0 and decreased to zero at pH 5.5. All the rPr-MnP3 mutant enzymes exhibited a similar profile with pH optimum at pH 3.0. A non-specific interaction site for ABTS in the general region of the Mn (II) binding site cannot be excluded since there are significant effects for all variants. It implies that protonation events are required for ABTS oxidation. The general effect is probably related to charge neutralization of sulphonate groups on ABTS.

4.3. pH Dependence of Mn (II) Oxidation Michaelis-Menten Kinetic Parameters

Given earlier results (Figure 1) showing significant Mn (II) oxidation activity at both high and low pH for wild-type rPr-MnP3, Mn (II) oxidation activity was measured at pH 5.0 and pH 8.0 in the presence of 100 mM tartrate. The pH variations of Mn²⁺-MnP3 reactions may provide uniquely useful information to assess relative importance of ionisable functional groups on the enzyme as well as the substrates.

The pH dependence of Mn (II) oxidation activity was measured to determine whether mutation of the Mn-binding site has affected the pH dependence kinetic parameters of the wild-type rPr-MnP3 and the mutant variants. The results as presented in Figure 4(a) for wild-type Mn (II) oxidation at pH 5.0 and 8.0, and in Figure 4(b) for Mn (II) oxidation by mutant enzymes at pH 8.0 illustrate the pH dependence of Mn (II) steady-state kinetic parameters, particularly, Km, Kcat and catalytic efficiency (Kcat/Km) for wild-type recombinant Phlebiaradiata MnP3 and variants in which the anionic charge at position 40, 44 and 186 has been removed (E40H, E44H, E40H/E44H, D186H, and D186N). The complete results for these reactions are presented in Table 2. The catalytic activity of most peroxidase enzymes is strongly dependent on pH. Manganese peroxidase is known to have a low pH optimum, achieving the highest at pH 4.5 (4). However, in this study the pH optimum for Mn (II) by wild-typePhlebia radiata MnP3 is at pH 5.0 and the mutants showed greater effectiveness at different pH values ranging from about neutral to alkaline.

In wild-type and mutants rPr-MnP3 enzymes, the apparentK_m value for Mn (II) oxidation decreased as the pH was increased. Activity assay at pH 5.0 yielded Km of 0.17 ± 10 mM for the wild-type rPr-MnP3. This Km values was observed to dramatically drop by approximately 94% (0.011 ± 0.0001 mM) when the reaction was conducted at pH 8.0. Likewise, the MnP3 mutant variants show a similar trend in Km at pH 5.0 and 8.0 as the Km for E40H, E44H, E40H/E44H, D179H, and D179N mutant variants at pH 8.0 decreased by 98%, 98.5%, 94.5%, 98.2% and 98.9% respectively relative to the value at pH 5.0. In this case, alkaline pH (8.0) extremely enhanced recombinant Phlebiaradiata MnP3 enzymes affinity for Mn²⁺. The wild-type distinctly shows acidic pH dependence with the Kcat value of 175 ± 3.0 s⁻¹ at the enzyme’s pH optimum (5.0) but at pH 8.0, the Kcat drops to 34.2 ± 0.8 s⁻¹ (81% decrease). Despite the huge drop in Kcat, the catalytic efficiency at pH 8.0 was approximately 3-times higher relative to the value at pH 5.0 It therefore implies that at pH 8.0 all rP.radiata MnP3 enzymes have enhanced affinity for Mn (II), showing that deprotonation of the Mn site results in progressively higher affinity.

The optimum pH of almost all ligninolytic enzymes, including MnP, reported to date lies in the acidic range. However, some industry activities, such as pulping and bleaching are mainly performed under highly alkaline conditions and the waste generated is also alkaline, this is an exciting finding. Ligninolytic enzymes having activity only in the acidic conditions cannot be used under alkaline conditions [53]. The ability of rPr-MnP3 to have a high catalytic efficiency over a wide pH range makes this enzyme unique. There is dearth of comparative data for P.chrysosporium MnP and other lignolytic peroxidases at high pH. Therefore, whether this property has simply been overlooked or not is unclear. In contrast to the wild-type enzyme, the k_cat values at pH 8.0 for four of the rPr-MnP3 mutant enzymes, E40H, E44H, E40H/E44H and D186H, were significantly higher by approximately 42%, 60%, 95% and 98% respectively than the corresponding values at pH 5.0 (Table 2). This, along with the already mentioned dramatic decreases inK_m leads to an increase effectiveness ratio for all variants. However, none of the mutant enzymes reach the efficiency of the wild-type enzyme, even at pH 8.0. This implies that the mutation of the Mn²⁺-binding site of MnP3 has generated enzyme with reasonable effectiveness at alkaline pH region. With the significant reduction in the Km for rPr-MnP3 enzymes at pH 8.0 and the enzyme’s corresponding increase in affinity for Mn²⁺, it is strongly believed that binding of Mn (II) to MnP3 will be highly favoured under alkaline conditions. Interestingly, like in MnP, the catalytic activity of wild-type Phlebiaradiata MnP3 increases with decreasing pH. This trend may reflect the increasing redox potential of reaction intermediates with decreasing pH observed for numerous peroxidases [54] [55] [56].

Mutation of the Mn²⁺ binding ligands had profound effects on the catalytic properties of the enzyme. The removal of a carboxylate ligand at the Mn (II)-binding site was found to severely depress the catalytic efficiency of E40H (1.0 mM⁻¹·s⁻¹), E44H (1.5 mM⁻¹·s⁻¹), E40H/E44H (0.08 mM⁻¹·s⁻¹¹), D186 (0.04 mM⁻¹·s⁻¹) and D186N (0.30 mM⁻¹·s⁻¹) for Mn (II) compared to wild-type (1029 mM⁻¹·s⁻¹) at lower pH. Mutation specific preferences were also created with the rPr-MnP3 DI86N variant being the kinetically poorer than D186H at higher pH. Clearly His is a more acceptable alternative for catalysis than Asn. These changes in k_cat and the specificity constants for Mn (II) oxidation by rPr-MnP3 mutants were to some extent expected from the nature of the mutants involved and prior work on the Phanerochaetechrysosporium MnP enzymes [57].

Excitingly, it was observed that histidine substitutions at the Mn (II) binding site lead to a mutant with a similar effectiveness ratio to the wild-type enzyme at higher pH, when the His ligands were in a deprotonated state. Activity towards Mn (II) at lower pH was greatly decreased, presumably due to the presence of the positively charged histidine. In contrast to expectation, the enzyme can therefore function at higher pH with alternative ligands in the Mn²⁺ site and does not have an absolutely obligate requirement for an all carboxylate ligand set. The results obtained for wild-type rPr-MnP3 for Mn (II) oxidation at pH 5.0 and 8.0 also showed enhanced affinity for Mn (II) at alkaline pH. Progressive deprotonation of Mn ligands results in a progressively higher affinity for Mn, which is evident in a relatively good catalytic efficiency at higher pH, even for the mutants, E40H (92 mM⁻¹·s⁻¹), E44H (244 mM⁻¹·s⁻¹), E40H/E44H (29 mM⁻¹·s⁻¹), D186H (128 mM⁻¹·s⁻¹) and D186N (15 mM⁻¹·s⁻¹) But none of the catalytic efficiencies of the mutants was in any way near the of the wild-type (3109 mM⁻¹·s⁻¹) at alkaline pH. The rPr-MnP3 may be used as an alternative catalytic agent in some industrial applications since it was heat stable up to ~60°C [58] and in this study its activity was retained over a wide range of pH values (5.0 - 9.0).

5. Conclusion

The wild-type recombinant Phlebia radiata MnP3 enzyme showed optimal activity with Mn (II) at pH 5.0 and remained moderately active (approximately 40%) in the pH range 6.0 - 8.5; the rPr-MnP3 mutants maximum activity ranged between the pH of 5.5 and 8.0. The activity of manganese peroxidase at higher pH is not common and a unique characteristic that has not been reported for any MnP enzyme before and therefore would have considerable interest for commercial uses. Wild-type rPr-MnP3 seemed to have a substantially higher pH optimum than the more classical Phanerochaete MnP enzyme. The rPr-MnP3 mutants exhibited a similar pH profile to wild-type, with an optimum at pH 3.0, for ABTS oxidation. This strongly suggests that ABTS is oxidised at the Mn (II) binding site, as charge neutralisation mutations decreased K_m for ABTS and improved effectiveness. Our findings offer a new insight into MnPs with novel properties that will find applications in the paper and pulp industries. The results offered new insight into MnPs with novel properties for industrial applications.

Acknowledgements

The authors would like to acknowledge the financial support this project has received from Akwa Ibom State University, Nigeria. The authors sincerely thank Prof. Andrew Smith of the University of Sussex for providing the laboratory space and technical support for research activity. Dr. Wendy Doyle is also acknowledged for her immense technical support during the experiments.

Conflicts of Interest

The authors declare no conflicts of interest regarding the publication of this paper.

References

1. Shin, K.S., Kim, Y.H. and Lim, J.S. (2005) Purification and Characterization of Manganese Peroxidase of the White-Rot Fungus Irpex lacteus. Journal of Microbiology, 43, 503-509.

2. Xu, H., Guo, M., Gao, Y., Bai, X. and Zhou, X. (2017) Expression and Characteristics of Manganese Peroxidase from Ganoderma lucidum in Pichia pastoris and Its Application in the Degradation of Four Dyes and Phenol. BMC Biotechnology, 17, Article No. 19. https://doi.org/10.1186/s12896-017-0338-5

3. Paszczynski, A., Huynh, V.B. and Crawford, R. (1985) Enzymatic Activities of an Extracellular, Manganese-Dependent Peroxidase from Phanerochaete chrysosporium. FEMS Microbiology Letters, 29, 37-41. https://doi.org/10.1111/j.1574-6968.1985.tb00831.x

4. Glenn, J.K. and Gold, M.H. (1985) Purification and Characterization of an Extracellular Mn(II)-Dependent Peroxidase from the Lignin-Degrading Basidiomycete, Phanerochaete chrysosporium. Archives of Biochemistry and Biophysics, 242, 329-334. https://doi.org/10.1016/0003-9861(85)90217-6

5. Lundell, T.K., Makela, M.R. and Hildén, K. (2010) Lignin-Modifying Enzymes in Filamentous Basidiomycetes-Ecological, Functional and Phylogenetic Review. Journal of Basic Microbiology, 50, 5-20. https://doi.org/10.1002/jobm.200900338

6. Tello, M., Corsini, G., Larrondo, L.F., Salas, L., Lobos, S. and Vicuna, R. (2000) Characterization of Three New Manganese Peroxidase Genes from the Ligninolytic Basidiomycete Ceriporiopsis subvermispora. Biochimica et Biophysica Acta, 1490, 137-144. https://doi.org/10.1016/S0167-4781(99)00227-4

7. Maeda, Y., Kajiwara, S. and Ohtaguchi, K. (2001) Manganese Peroxidase Gene of the Perennial Mushroom Elfvingia applanata: Cloning and Evaluation of Its Relationship with Lignin Degradation, Biotechnology Letters, 23, 103-109. https://doi.org/10.1023/A:1010341232388

8. Forrester, I.T., Grabski, A.C., Mishra, C., Kelly, B.D., Striekl, W.N., Leatham, G.E. and Burgess, R.R. (1990) Characteristics and N-Terminal Amino Acid Sequence of a Manganese Peroxidase Purified from Lentinula edodes Culture Grown on a Commercial Wood Substrate. Applied Microbiology and Biotechnology, 33, 359-365. https://doi.org/10.1007/BF00164536

9. Hatakka, A.I. (1994) Lignin-Modifying Enzymes from Selected White-Rot Fungi-Production and Role in Lignin Degradation. FEMS Microbiology Reviews, 13, 125-135. https://doi.org/10.1111/j.1574-6976.1994.tb00039.x

10. Perez, J. and Jeffries, T.W. (1992) Roles of Manganese and Organic Acid Chelators in Regulating Lignin Degradation and Biosynthesis of Phanerochaete chrysosporium. Applied and Environmental Microbiology, 58, 2402-2409. https://doi.org/10.1128/aem.58.8.2402-2409.1992

11. Sarkar, S., Martinez, A.T. and Martinez, M.J. (1997) Biochemical and Molecular Characterization of a Manganese Peroxidases Isoenzyme from Pleurotus ostreatus. Biochimica et Biophysica Acta, 1339, 23-30. https://doi.org/10.1016/S0167-4838(96)00201-4

12. Kanayama, N., Suzuki, T. and Kawai, K. (2002) Purification and Characterization of an Alkaline Manganese Peroxidase from Aspergillus terreus LD-I. Journal of Bioscience and Bioengineering, 93, 405-410. https://doi.org/10.1016/S1389-1723(02)80075-5

13. Purcell, W.L. and Erman, J.E. (1976) Cytochrome c Peroxidase Catalyzed Oxidations of Substitution Inert Iron II Complexes. Journal of the American Chemical Society, 98, 7033-7037. https://doi.org/10.1021/ja00438a049

14. Hayashi, Y. and Yamazaki, I. (1979) The Oxidation-Reduction Potentials of Compound I, Compound II and Compound II Ferric Couples of Horseradish Peroxidases A2 and C. Journal of Biological Chemistry, 254, 101-106. https://doi.org/10.1016/S0021-9258(19)86816-3

15. Ivan, A., Antonio, L., De Lacey, F., Cacada, J., Francisco, J., Ruiz-Due, C. and Martnez, A.T. (2019) Increase of Redox Potential during the Evolution of Enzymes Degrading Recalcitrant Lignin. Journal of the American Chemical Society, 25, 2708-2712. https://doi.org/10.1002/chem.201805679

16. Kishi, K., Kustersvan Someren, M., Mayfield, M.B., Sun, J., Loehr, T.M. and Gold, M.H. (1996) Characterization of Manganese(II) Binding Site Mutants of Manganese Peroxidase. Biochemistry, 35, 8986-8994. https://doi.org/10.1021/bi960679c

17. Ufot, U.F. and Akpanabiatu, M.I. (2014) Influence of CaCl2 and EDTA on Reversible Thermal Inactivation of Recombinant Wild-Type and Mutant (E40H/E44H) Phlebia radiata Manganese Peroxidase 3(rPr-MnP3). African Journal of Biochemistry Research, 8, 118-126. https://doi.org/10.5897/AJBR2014.0788

18. Perie, F.H., Sheng, D. and Gold, M.H. (1996) Purification and Characterization of Two Manganese Peroxidase Isozymes from the White-Rot Basidomycete Dichomitus squalens. Biochimica et Biophysica Acta, 1297, 139-148. https://doi.org/10.1016/S0167-4838(96)00096-9

19. Johnson, C.R., Cullen, D. and Lamar, R.T. (1994) Manganese Peroxidases of the White Rot Fungus Phanerochaete sordida. Applied and Environmental Microbiology, 60, 599-605. https://doi.org/10.1128/aem.60.2.599-605.1994

20. Hinder, A.N.P., Ruiz, J.H., Lopez, J.N.R. Canovas, F.G., Brisset, N.C., Smith, A.T., Arnao, M.B. and Acosta, M. (2002) Reactions of the Class II Peroxidase, Lignin Peroxidase and Arthro ramosus Peroxidase with Hydrogen Peroxidase. Journal of Biological Chemistry, 277, 26879-26885. https://doi.org/10.1074/jbc.M200002200

21. Bonnen, A.M., Anton, L.H. and Orth, A.B. (1994) Lignin Degrading Enzymes of the Commercial Button Mushroom, Agaricus bisporus. Applied and Environmental Microbiology, 60, 960-965. https://doi.org/10.1128/aem.60.3.960-965.1994

22. Whitaker, J.R. (1994) Effect of pH on Enzyme Catalysed Reactions. In: Principles of Enzymology for the Food Science, 2nd Edition, Marcel Dekker, New York, 271-300. https://doi.org/10.1201/9780203742136-10

23. Dogan, S., Dogan, M. and Arslan, O. (2003) Characterization of Polyphenol Oxidase from Thymus (Thymus longicaulis var subisophyllus). Advances in Food Sciences, 25, 6-64.

24. Abelskov, A.K., Smith, A.T., Rasmussen, C.B., Dunford, H.B. and Welinder, K.G. (1995) pH-Dependence and Structural Interpretation of the Reactions of Coprinus cinereus Peroxidase with Hydrogen Peroxide, Ferulic Acid, and 2,2’-Azinobis(3-ethylbenzthiazoline-6-sulfonic acid). Biochemistry, 34, 4022-4029.

25. Ufot, U.F. and Akpanabiatu, M.I. (2012) An Engineered Phlebia radiata Manganese Peroxidase: Expression, Refolding, Purification and Preliminary Characterization. American Journal of Molecular Biology, 2, 359-370. https://doi.org/10.4236/ajmb.2012.24037

26. Campos, A.M. and Lissi, E.A. (1997) Kinetics of the Reaction between 2,2-Azinobis (3-ethylbenzothiazoline-6-sulfonic acid) (ABTS) Derived Radical Cations and Phenols. International Journal of Chemical Kinetics, 29, 219-224. https://doi.org/10.1002/(SICI)1097-4601(1997)29:3<219::AID-KIN9>3.0.CO;2-X

27. Aliaga, C. and Lissi, E.A. (1998) Reaction of 2,2-Azinobis(3-ethylbenzothiazoline-6-sulfonic Acid) (ABTS) Derived Radicals with Hydroperoxides. Kinetics and Mechanism. International Journal of Chemical Kinetics, 30, 565-570. https://doi.org/10.1002/(SICI)1097-4601(1998)30:8<565::AID-KIN5>3.0.CO;2-Q

28. Hofrichter, M., Vares, K., Scheibner, K., Galkin, S., Sipila, J. and Hatakka, A. (1999) Mineralization and Solubilization of Synthetic Lignin by Manganese Peroxidases from Nematoloma frowardii and Phlebia radiata. Journal of Biotechnology, 67, 217-228. https://doi.org/10.1016/S0168-1656(98)00180-1

29. Martinez, M.J., Ruiz-Duenas, F.J., Guillén, F. and Martínez, A.T. (1996) Purification and Catalytic Properties of Two Manganese Peroxidase Isoenzymes from Pleurotus eryngii. European Journal of Biochemistry, 237, 424-432. https://doi.org/10.1111/j.1432-1033.1996.0424k.x

30. Paice, M.G., et al. (1993) Manganese Peroxidase, Produced by Trametes versicolor during Pulp Bleaching Demethylates and Delignifies Kraft Pulp. Applied and Environmental Microbiology, 59, 260-265. https://doi.org/10.1128/aem.59.1.260-265.1993

31. Childs, R.E. and Bardsley, W.G. (1975) The Steady-State Kinetics of Peroxidase with 2,2’-Azino-di-(3-ethyl-benzthiazoline-6-sulphonic acid) as Chromogen. Biochemical Journal, 145, 93-103. https://doi.org/10.1042/bj1450093

32. Perez, J. and Jeffries, T.W. (1990) Mineralization of 14C-Ring-Labeled Synthetic Lignin Correlates with the Production of Lignin Peroxidase, Not Mn-Peroxidase or Laccase. Applied and Environmental Microbiology, 56, 1806-1812. https://doi.org/10.1128/aem.56.6.1806-1812.1990

33. Brown, J., Glenn, J.K. and Gold, M.H. (1990) Manganese Regulates Expression of Manganese Peroxidase by Phanerochaete chrysosporium. Journal of Bacteriology, 172, 3125-3130. https://doi.org/10.1128/jb.172.6.3125-3130.1990

34. Bonnarme, P. and Jeffries, T.W. (1990) Mn(II) Regulation of Lignin Peroxidase and Manganese-Dependent Peroxidase from Ligni-Degrading White-Rot Fungi. Applied and Environmental Microbiology, 56, 210-217. https://doi.org/10.1128/aem.56.1.210-217.1990

35. Wariishi, H., Valli, K. and Gold, M.H. (1991) In Vitro Depolymerization of Lignin by Manganese Peroxidase of Phanerochaete chrysosporium. Biochemical and Biophysical Research Communications, 176, 269-275. https://doi.org/10.1016/0006-291X(91)90919-X

36. Wariishi, H., Akileswaran, L. and Gold, M.H. (1988) Manganese Peroxidase from the Basidiomycete Phanerochaete chrysosporium: Spectral Characterization of the Oxidized States and the Catalytic Cycle. Biochemistry, 27, 5365-5370. https://doi.org/10.1021/bi00414a061

37. Pankaj, C., Gargi, S., Garima, R., Luiz, F., Romanholo, F. and Ram, N.B. (2019) Microbial Manganese Peroxidase: A Ligninolytic Enzyme and Its Ample Opportunities in Research. SN Applied Sciences, 1, 45. https://doi.org/10.1007/s42452-018-0046-3

38. Glenn, J.K., Akileswaran, L. and Gold, M.H. (1986) Mn (II) Oxidation Is the Principal Function of Extracellular Mn-Peroxidase from Phanerochaete chrysosporium. Archives of Biochemistry and Biophysics, 251, 688-696. https://doi.org/10.1016/0003-9861(86)90378-4

39. Garcia, J.S., Magalhnes, C.S.d. and Arruda, M.A.Z. (2006) Trends in Metal-Binding and Metalloprotein Analysis. Talanta, 69, 1-15. https://doi.org/10.1016/j.talanta.2005.08.041

40. Maneiro, M., Ruettinger, W.F., Bourles, E., McLendon, G.L. and Dismukes, G.C. (2003) Kinetics of Proton-Coupled Electron-Transfer Reactions to the Manganese-Oxo “Cubane” Complexes Containing the MnO4 and MnO4 Core Types. Proceedings of the National Academy of Sciences, 100, 3707-3712. https://doi.org/10.1073/pnas.0637229100

41. Wariishi, H., Valli, K. and Gold, M.H. (1992) Manganese (II) Oxidation by Manganese Peroxidase from the Basidiomycete Phanerochaete chrysosporium. Kinetic Mechanism and Role of Chelators. Journal of Biological Chemistry, 267, 23688-23695. https://doi.org/10.1016/S0021-9258(18)35893-9

42. Kennedy, M.L. and Gibney, B.R. (2001) Metalloprotein and Redox Protein Design. Current Opinion in Structural Biology, 11, 485-490. https://doi.org/10.1016/S0959-440X(00)00237-2

43. Gregory, D.S., Martin, A.C.R., Cheetham, J.C. and Rees, A.R. (1993) The Prediction and Characterization of Metal Binding Sites in Proteins. Protein Engineering, 6, 29-35. https://doi.org/10.1093/protein/6.1.29

44. Wariishi, H., Dunford, H.B., MacDonald, I.D. and Gold, M.H. (1989) Manganese Peroxidase from the Lignin-Degrading Basidiomycete Phanerochaete chrysosporium. Transient State Kinetics and Reaction Mechanism. Journal of Biological Chemistry, 264, 3335-3340. https://doi.org/10.1016/S0021-9258(18)94070-6

45. Paszczyński, A., Huynh, V.-B. and Crawford, R. (1986) Comparison of Ligninase-I and Peroxidase-M2 from the White-Rot Fungus Phanerochaete chrysosporium. Archives of Biochemistry and Biophysics, 244, 750-765. https://doi.org/10.1016/0003-9861(86)90644-2

46. Kishi, K., Wariishi, H., Marquez, L., Dunford, H.B. and Gold, M.H. (1994) Mechanism of Manganese Peroxidase Compound II Reduction. Effect of Organic Acid Chelators and pH. Biochemistry, 33, 8694-8701. https://doi.org/10.1021/bi00195a010

47. Kuan, I.C., Johnson, K.A. and Tien, M. (1993) Kinetic Analysis of Manganese Peroxidase. The Reaction with Manganese Complexes. Journal of Biological Chemistry, 268, 20064-20070. https://doi.org/10.1016/S0021-9258(20)80694-2

48. Praveen, K., Usha, K.Y., Viswanath, B. and Reddy, B.R. (2012) Kinetic Properties of Manganese Peroxidase from the Mushroom Stereum ostrea and Its Ability to Decolorize Dyes. Journal of Microbiology and Biotechnology, 22, 1540-1548. https://doi.org/10.4014/jmb.1112.12011

49. Susla, M., Novotny, C., Erbanová, P. and Svobodová, K. (2008) Implication of Dichomitus squalens Manganese-Dependent Peroxidase in Dye Decolourization and Cooperation of the Enzyme with Laccase. Folia Microbiologica, 3, 479-485. https://doi.org/10.1007/s12223-008-0075-1

50. Champagne, P.P. and Ramsay, J.A. (2005) Contribution of Manganese Peroxidase and Laccase to Dye Decolouration by Trametes versicolor. Applied Microbiology and Biotechnology, 69, 276-285. https://doi.org/10.1007/s00253-005-1964-8

51. Boer, C.G., Obici, L., de Souza, C.G. and Peralta, R.M. (2004) Decolourization of Synthetic Dyes by Solid State Cultures of Lentinula (Lentinus) Edodes Producing Manganese Peroxidase as the Main Ligninolytic Enzyme. Bioresource Technology, 94, 107-112. https://doi.org/10.1016/j.biortech.2003.12.015

52. Rodríguez, C.S., Domínguez, A. and Sanromán, A. (2002) Production of Manganese-Dependent Peroxidase in a New Solid-State Bioreactor by Phanerochaete chrysosporium Grown on Wood Shavings. Application to the Decolorization of Synthetic Dyes. Folia Microbiologicaogica, 47, 417-421. https://doi.org/10.1007/BF02818701

53. Qin, X., Zhang, J., Zhang, X. and Yang, Y. (2014) Induction, Purification and Characterization of a Novel Manganese Eroxidase from Irpex lacteus CD2 and Its Application in the Decolorization of Different Types of Dye. PLoS ONE, 9, e113282. https://doi.org/10.1371/journal.pone.0113282

54. Steffen, K.T., Hofrichter, M. and Hatakka, A. (2003) Purification and Characterization of Manganese Peroxidases from the Litter-Decomposing Basidiomycetes Agrocybe praecox and Stropharia coronilla. Enzyme and Microbial Technology, 30, 550-555. https://doi.org/10.1016/S0141-0229(01)00525-7

55. Dong, Y.C., Dai, Y.N., Xu, T.Y., Cai, J. and Chen, Q.H. (2014) Biodegradation of Chestnut Shell and Lignin-Modifying Enzymes Production by the White-Rot Fungi Dichomitus squalens, Phlebia radiata. Bioprocess and Biosystems Engineering, 37, 755-764. https://doi.org/10.1007/s00449-013-1045-9

56. Sakamoto, Y., Nakade, K., Nagai, M., Uchimiya, H. and Sato, T. (2009) Cloning of Lentinula edodes lemnp2, a Manganese Peroxidase That Is Secreted Abundantly in Sawdust Medium. Mycoscience, 50, 116-122. https://doi.org/10.1007/S10267-008-0463-Z

57. Lankinen, P., Hildén, K., Aro, N., Salkinoja-Salonen, M. and Hatakka, A. (2005) Manganese Peroxidase of Agaricus bisporus: Grain Bran-Promoted Production and Gene Characterization. Applied Microbiology and Biotechnology, 66, 401-407. https://doi.org/10.1007/s00253-004-1731-2

58. Johansson, T., Nyman, P.O. and Cullen, D. (2002) Differential Regulation of mnp2, a New Manganese Peroxidase-Encoding Gene from the Ligninolytic Fungus Trametes versicolor PRL 572. Applied Microbiology and Biotechnology, 68, 2077-2080. https://doi.org/10.1128/AEM.68.4.2077-2080.2002