A Novel Synthesis of Bioactive Catechols by Layer-by-Layer Immobilized Tyrosinase in an Organic Solvent Medium

Abstract

Commercial and extracted mushroom tyrosinases were supported on Eupergit C 250 L and protected by a coating of oppositely charged polyelectrolytes by means of the layer-by-layer technique. The kinetic parameters (K_m, V_max, and V_max/K_m) of these novel biocatalysts in both organic and aqueous media were evaluated, showing tyrosinase to be more reactive in organic solvent than in buffer solution. Heterogeneous tyrosinase systems were used for the oxidation of a large group of phenol derivatives to the corresponding catechols. Different enzyme reactivities were found depending on the substrate structure. The catalyst activity was retained for successive runs. Catechols are difficult to synthesize through traditional chemical methods under environmentally friendly conditions; the use of immobilized tyrosinase opens a novel synthetic alternative to this interesting biologically active family of substances.

Introduction

Tyrosinase (polyphenol oxidase, EC 1.14.18.1) is a widely diffused copper-containing monooxygenase with two distinct substrate-binding sites, one with high affinity for aromatic compounds, including phenolic substrates, and the other specific for metal-binding agents and dioxygen.1 This enzyme catalyzes both the hydroxylation of monophenols to catechols (cresolase or monophenolase activity) and the oxidation of catechols to ortho-benzoquinones (catecholase or diphenolase activity) by use of dioxygen as the primary oxidant.2 Catechols are characterized by several biological activities and are well recognized as antioxidant compounds.3 As catechols are often difficult to synthesize in an environmentally friendly way by means of chemical methods, the potential of tyrosinase to catalyze the oxidation of phenols has received great attention. Investigations have shown that a wide range of substrates can be transformed to the desired catechols in conventional aqueous-based systems.4 A limitation in the use of tyrosinase is, however, the facile polymerization of ortho-benzoquinones leading to inactivation of the enzyme.5 This drawback can be overcome by the addition of reducing agents such as ascorbic acid6 or by the use of organic solvents as opposed to conventional aqueous media.7 Developments in the field of non-aqueous enzymology have revealed numerous advantages: increased solubility of hydrophobic substrates, improved stability of tyrosinase in water-immiscible organic solvents, enhanced degree of selectivity, and limited ortho-benzoquinone polymerization.8 Moreover, in industrial applications, the immobilization of enzymes on inert supports favours the reusability, enhances stability of the enzyme, and facilitates purification of the products. Tyrosinase has been so far immobilized on various types of supports by different methods, such as physical adsorption, covalent crosslinking, and microcapsule entrapment.9 There are only a few examples, however, of the use of immobilized tyrosinase in organic solvents for the synthesis of catechols.5, 10

We have prepared novel biocatalysts through the immobilization of tyrosinase on Eupergit C 250 L, a commercially available and low-cost epoxy-activated acrylic resin that has been successfully used for the covalent binding of several oxidases such as laccase,11 glucose oxidase,12 and pyranose oxidase.13 To further increase the stability of the catalyst, the layer-by-layer (LbL) technique, first introduced by Decher,14 was applied. This method is based on the consecutive deposition of alternatively charged polyelectrolytes on the active species.15 LbL is an effective tool for the stabilization of enzymes because polyelectrolyte films can protect proteins from high-molecular-mass denaturing agents by modifying their permeability towards substrates that could enter the multilayer and react with the catalytic site.16 Both tyrosinases crosslinked on Eupergit C 250 L, and the biocatalysts prepared using the LbL technique were used for the oxidation of a large range of phenols in an organic mixed solvent, dichloromethane/buffer, to afford the corresponding catechols in good yield. A comparison of the efficiency and the selectivity of tyrosinase with and without the LbL treatment is also reported, as is the recyclability of the biocatalysts.

Results and Discussion

Evaluation of the enzyme activity and kinetic parameters of tyrosinase

In this study we used a commercially available mushroom tyrosinase (purchased from Sigma-Aldrich, Tyro^S) and a freshly purified tyrosinase from Agaricus bisporus (Tyro^E) obtained by a modification of a procedure previously reported in the literature.17

Tyrosinase activity was assayed by the dopachrome method using L-tyrosine (L-Tyr) as a substrate.18 All experiments were done in triplicate. One unit of enzyme activity (IU) was defined as the increase of 0.001 min⁻¹ in absorbance at 25 °C (3.0 mL reaction mixture) in sodium phosphate buffer (pH 7). The specific activity (U mg⁻¹) is defined as the ratio between the enzyme activity and the amount (mg) of enzyme. Both tyrosinases showed a significant value of enzyme activity, the commercial enzyme being more reactive than the extracted one (Table 1, entries 1 and 2). As tyrosinase preparations usually contain laccase contaminant, the activity of residual laccase was also evaluated by using conventional 2,2′-azino-bis(3-ethylbenzthiazoline-6-sulfonic acid (ABTS) assay.19 Values of the ratio of tyrosinase and laccase activity (Tyro/Lac) are reported in Table 1. The amount of laccase was found to be higher in the case of extracted enzyme (2300×10³ for Tyro^S and 23×10³ for Tyro^E).

Table 1. Specific activity and contaminant laccase activity of commercial (Tyro^S) and extracted (Tyro^E) tyrosinases.

Entry	Enzyme	Specific activity [U mg^−1]	Tyro/Lac activity [×10^3]
1	Tyro^S	13 881	2300
2	Tyro^E	2568	23

The kinetic parameters (K_m, V_max, and V_max/K_m) of Tyro^S and Tyro^E were determined by measuring the enzyme activity at different concentrations of L-Tyr (330–1000 μM) and plotting data to a Lineweaver–Burk plot (Table 2 and Figure 1).20 Tyro^E showed a K_m value higher than that of Tyro^S, which suggested a reduced affinity for the substrate. This pattern was confirmed by the value of the maximum reaction rate (V_max), which was lower for Tyro^E. Furthermore, the value of V_max/K_m for Tyro^S was higher than that for Tyro^E (Table 2, entry 1 vs. entry 2). Owing to the high cost of the commercial enzyme, the efficiency of both tyrosinases was compared and the possibility of using the mushroom-extracted option was evaluated.

Table 2. Kinetic parameters of commercial (Tyro^S) and extracted (Tyro^E) tyrosinases.

Entry	Enzyme	Km [μM]	Vmax×10^−3 [ΔAbs min^−1 μgenzyme^−1]	Vmax/Km×10^−6 [ΔAbs min^−1 μgenzyme^−1 μM^−1]^[a]
[a] Each experiment was conducted in triplicate; average errors in kinetic parameters were ±2–4 % for Km and ±1–3 % for Vmax.
1	Tyro^S	180	6.02	33.4
2	Tyro^E	257	4.00	16.1

Figure 1. Lineweaver–Burk plots of ▴ commercial (Tyro^S, R²=0.9951) and ▪ extracted (Tyro^E, R²=0.9988) tyrosinase activity determined at different concentrations of L-Tyr ([S]=330–1000 μM). The data are the mean values of three experiments with a standard deviation of less than 1 %.

Tyrosinase immobilization procedure

Initially, the immobilization of Tyro^S and Tyro^E was performed on the epoxy-activated acrylic beads of Eupergit C 250 L by using a modification of previously reported procedures.21 The appropriate enzyme (5.0 mg, 69 405 IU of Tyro^S and 12 840 IU of Tyro^E) was suspended briefly in a buffer (sodium phosphate buffer 0.1 M, pH 7) in the presence of Eupergit C 250 L (1.0 g) for 24 h at room temperature. The immobilized tyrosinases (Tyro^S/E and Tyro^E/E) were washed with water to remove excess of protein and treated with glycine to block residual epoxy groups. The effectiveness of the immobilization procedure was investigated in terms of immobilization yield by the analysis of the residual enzyme activity in wastewaters after the reaction with the support (Figure 2).

Figure 2. Schematic representation of a) Tyro/E and b) Tyro/E-LbL.

Under these experimental conditions, 31 362 IU of Tyro^S and 6197 IU of Tyro^E were immobilized. Tyro^S/E and Tyro^E/E retained approximately 37 % of their native activity (11 604  U g⁻¹ for Tyro^S/E and 2293 U g⁻¹ for Tyro^E/E). With the aim to further increase the stability of Tyro^S/E and Tyro^E/E, the LbL technique was applied by coating the particles through a sequential deposition of alternatively charged polyelectrolytes. Biocatalysts were suspended briefly in positively charged poly(allylamine hydrochloride) (PAH; 2 mg mL⁻¹ in 0.5 M NaCl), filtrated, and then treated with negatively charged polystyrene sulfonate (PSS; 2 mg mL⁻¹ in 0.5 M NaCl) (Figure 2). This procedure was repeated until the formation of three layers. The immobilized LbL enzymes retained approximately 87 % of the activity with respect to Tyro^S/E and Tyro^E/E (10 095 U g⁻¹ for Tyro^S/E-LbL and 1995 U g⁻¹ for Tyro^E/E-LbL).

Novel immobilized tyrosinases were characterized in terms of their kinetic properties by using L-Tyr (330–1000 μM) as a substrate (Table 3). Irrespective of procedures used for the immobilization, V_max decreased and K_m increased for supported tyrosinases, which led to a partial reduction in the catalytic efficiency with respect to free enzyme. Similar trends in K_m values were reported for tyrosinase immobilized on other carriers and are attributed to a possible mass transfer limitation.22 Tyro^S-based heterogeneous biocatalysts were more reactive than Tyro^E systems. Note that the difference in the reactivity of commercial tyrosinase and that of extracted tyrosinase is more pronounced in the case of immobilized enzymes. A set of scanning electron microscopy (SEM) photographs that show the morphology of the surface of particles is depicted in Figures 3, 3–5. Tyro^S/E shows particles with a very regular shape and an average value of diameter of the order of 150 μm (Figure 3 a).

Table 3. Kinetic parameters of free and immobilized commercial (Tyro^S) and extracted (Tyro^E) tyrosinases.

Entry	Enzyme		Km [μM]			Vmax×10^−3 [ΔAbs min^−1 μgenzyme^−1]			(Vmax/Km)×10^−6 [ΔAbs min^−1 μgenzyme^−1 μM^−1]^[a]
			Tyro^S	Tyro^E		Tyro^S	Tyro^E		Tyro^S	Tyro^E
[a] Each experiment was conducted in triplicate; average errors in kinetic parameters were ±2–4 % for Km and ±1–3 % for Vmax.
1	Free		180	257		6	4		33	16
2	Tyro/E		270	320		4	2		15	6
3	Tyro/E-LbL		300	385		3	1		10	3

Figure 3. SEM images of Tyro^S/E at magnifications of a)×200, scale=100 µm and b)×7500, scale=1 µm.

Figure 4. SEM images of Tyro^S/E after treatment in dichloromethane at magnifications of a)×200, scale=100 µm and b)×7500, scale=1 µm.

Figure 5. SEM image of TyroS/E-LbL at ×7500 magnification, scale=1 µm.

A low number of irregular fragments were observed, which are probably formed by a mechanical damage of particles during the preparation of the sample. At the largest magnification, the particles show an irregular surface characterized by grumes of different dimensions (Figure 3 b). A similar behaviour is observed for Tyro^S/E after treatment with dichloromethane, which suggests the stability of the resin in an organic medium (Figure 4). Figure 5 shows the SEM photograph of Tyro^S/E-LbL. The ultrathin coating layers cover the surface of the particles.

Optimization of water requirement for oxidation in an organic solvent

The fundamental role of water in the functioning of proteins and in their three-dimensional structure is well recognized and has been reviewed by several authors.23 In low-water-content media, the relationships between the solvation process and the enzyme kinetics have been reported,24 with the hydration of the active site of the enzyme playing a critical role in substrate recognition and transformation.25 Water molecules can interact with the protein surface and occupy internal cavities and deep clefts, which optimizes spatial configurations and minimizes energetic pathways.26 Because of the uncertainty of the effect of this process in organic solvents, the amount of added buffer has to be optimized for any specific catalytic system. For this reason, we evaluated the dependence of the reaction rate versus the buffer concentration.

para-Cresol 1 was selected as a representative phenol substrate. The amount of sodium phosphate buffer (10–70 μL) in the presence of dichloromethane (2.5 mL) was varied throughout the reactions. Data in Table 4 show that Tyro^S/E required more buffer per microgram of enzyme (9.3 μL μg_Tyro⁻¹) with respect to Tyro^E (2.9 μL μg_Tyro⁻¹) to reach the highest value of activity (7587 U mg⁻¹ for Tyro^S and 772 U mg⁻¹ for Tyro^E).

Table 4. Optimum buffer expressed as μL μg_Tyro⁻¹ for free enzyme and μL g_beads⁻¹ for immobilized enzyme.

Entry	Enzyme	Free Tyro [μL μgTyro^−1]	Tyro/E [μl gbeads^−1]	Tyro/E-LbL [μl gbeads^−1]
1	Tyro^E	2.9	1000	1000
2	Tyro^S	9.3	500	1000

In the case of immobilized Tyro^S, the optimal amount of buffer (defined as microliters per gram of support, μL g_beads⁻¹) was 500 μL g_beads⁻¹ for Tyro^S/E (2067 U g_beads⁻¹) and 1000 μL g_beads⁻¹ for Tyro^S/E-LbL (1798 U g_beads⁻¹), which showed a major requirement of buffer in the presence of polyelectrolyte layers. In the case of immobilized Tyro^E, both biocatalysts, Tyro^E/E and Tyro^E/E-LbL, needed the same amount of buffer (1000 μL g_beads⁻¹) to obtain the highest activity (420 U g_beads⁻¹ for Tyro^E/E and 365 U g_beads⁻¹ for Tyro^E/E-LbL). The kinetic parameters in dichloromethane/buffer were determined in the selected case of Tyro^S by measuring the tyrosinase activity at different concentrations of compound 1 (2–15 mM), using the amount of the buffer previously optimized. Measurements in buffer were also performed as references. As reported in Table 5, in the case of 1 the K_m of free and immobilized Tyro^S was higher in buffer than in dichloromethane, with Tyro^S/E showing the highest affinity for substrate in the organic solvent (Table 5, entry 2 vs. entries 1 and 3).27 With regards to catalytic efficiency (V_max/K_m), free and immobilized Tyro^S were more reactive in dichloromethane than in buffer, which showed a significant value of the acceleration factor (defined as the ratio between V_max/K_m in dichloromethane and in buffer) in the range of 1.30–4.96. Tyro^S/E was the most reactive immobilized system. These data are in accordance with the general effects shown by organic solvents on the magnitude of the acceleration factors for the oxidation of simple hydrophobic substrates.8b, c In fact, hydrophobic substrates are retained in the organic solvent with a higher efficiency than in the buffer; thus, less amount of the substrate is available for the active site, which gives rise to a higher catalytic efficiency and acceleration factor.28

Table 5. Kinetic parameters of free and immobilized Sigma Tyro (Tyro^S) using para-cresol as a substrate.

Entry	Enzyme	Vmax^[a]×10^−4 [ΔAbs min^−1 μgenzyme^−1]			Km (mM)			(Vmax/Km)×10^−4 [ΔAbs min^−1 μgenzyme^−1 μM^−1]			Acceleration factor^[a]
		CH2Cl2	Buffer		CH2Cl2	Buffer		CH2Cl2	Buffer
[a] Acceleration factor was defined as the ratio of Vmax/Km in CH2Cl2 to that in buffer; each experiment was conducted in triplicate; Average errors in kinetic parameters were ±2–4 % for Km and ±1–3 % for Vmax.
1	Free	350	410		1.5	8.6		233	47		4.96
2	Tyro^S/E	114	40		1.3	1.5		87	25		3.52
3	Tyro^S/E-LbL	447	60		3.4	6		13	10		1.30

Phenol oxidation

A range of phenols (Figure 6) was oxidized, including para-cresol 1, 4-ethyl phenol 2, 4-tert-butyl phenol 3, 4-sec-butyl phenol 4, 4-methoxy phenol 5, 4-chloro phenol 6, 4-chloro-2-methyl phenol 7, meta-cresol 8, and bis(4-hydroxyphenyl)methane 9. We initially studied the oxidation of 1 with both free and immobilized tyrosinases. The oxidation of 1 (0.05 mmol) with Tyro^S (263 IU) in dichloromethane/buffer (2.5 mL/176 μL; pH 7) at room temperature under O₂ atmosphere for 18 h afforded catechol 1 a as the main reaction product in 49 % yield and 97 % conversion of the substrate along with a low amount of pyrogallol derivative 1 b.

Figure 6. Phenols oxidized by free and immobilized tyrosinase.

Dimers 1 c–d, characterized by the formation of the CC crosslinkage between two phenol units, were also detected in appreciable amount (Scheme 1 and Table 6, entry 1). Pyrogallol 1 b is a product of further oxidation of 1 a. Pyrogallol derivatives are potent antioxidants characterized by several biological activities.29 In accordance with the data reported in the literature, the formation of dimers 1 c–d can be ascribed to reactive ortho-quinone intermediates through a nonenzymatic mechanism, even if the occurrence of a radical mechanism involving a phenoxy radical intermediate generated by the residual laccase activity cannot be completely ruled out.30 The catechol 1 a was recovered in lower yield when the same reaction was performed in reference buffer solution (Table 6, entry 2 vs. entry 1).

Scheme 1. Oxidation of phenols 1–4. Reagents and conditions: a) Tyro-based systems, O₂; b) CH₂Cl₂/buffer; c) CH₂Cl₂/buffer/CH₃CN; d) buffer.

Table 6. Oxidation of para-alkyl substituted phenols 1–4.^[a]

Entry	Substrate	Solvent	Biocatalysts	Products	Conversion [%]^[b]	Yield [%]^[b]
[a] Reaction conditions: substrate (0.05 mmol) and tyrosinase (263 IU) were taken in 2.5 mL of the appropriate solvent for 18–24 h in the presence of O2; [b] Conversion and yield were calculated by using GC–MS analysis using dodecane as the internal standard.
1	1	CH2Cl2/buffer	Tyro^S	1a (1b) [1c] {1d}	97	49 (4) [7] {25}
2	1	Buffer	Tyro^S	1a (1b) [1d]	95	22 (10) [26]
3	1	CH2Cl2/buffer	Tyro^S/E	1a (1b) [1c]	90	72 (7) [10]
4	1	CH2Cl2/buffer	Tyro^S/E-LbL	1a [1c]	85	78 [6]
5	1	CH2Cl2/buffer	Tyro^E	1a (1b) [1c] {1d}	89	60 (10) [12] {7}
6	1	Buffer	Tyro^E	1a (1b) [1c]	90	19 (39) [30]
7	1	CH2Cl2/buffer	Tyro^E/E	1a (1b) [1c]	91	53 (25) [12]
8	1	CH2Cl2/buffer	Tyro^E/E-LbL	1a (1b)	98	67 (29)
9	2	CH2Cl2/buffer	Tyro^E	2a (2b)	73	60 (10)
10	2	Buffer	Tyro^E	2a (2b) [2c] {2d}	92	15 (11) [30] {15}
11	2	CH2Cl2/buffer	Tyro^E/E	2a (2b)	91	67 (8)
12	2	Buffer	Tyro^E/E	2a (2b) [2c]	92	36 (42) [14]
13	2	CH2Cl2/buffer	Tyro^E/E-LbL	2a (2b)	98	84 (12)
14	2	Buffer	Tyro^E/E-LbL	2a (2b) [2c] {2d}	92	18 (16) [28] {16}
15	3	CH2Cl2/buffer/CH3CN	Tyro^E	3a	21	21
16	3	CH2Cl2/buffer/CH3CN	Tyro^E/E	3a	37	37
17	3	CH2Cl2/buffer/CH3CN	Tyro^E/E-LbL	3a	39	39
18	4	CH2Cl2/buffer/CH3CN	Tyro^E	4a	37	37
19	4	CH2Cl2/buffer/CH3CN	Tyro^E/E	4a	55	44
20	4	CH2Cl2/buffer/CH3CN	Tyro^E/E-LbL	4a	36	36

In this latter case, a high amount of 1 b was obtained, with compound 1 d being the only isolated dimer (Table 6, entry 2). The low mass balance of the reaction further suggested the formation of some overoxidized products, not recovered under our experimental conditions. Next, the efficiency of Tyro^S/E and Tyro^S/E-LbL was evaluated. The oxidation of 1 (0.05 mmol) with Tyro^S/E (263 IU) in dichloromethane/buffer (2.5 mL/13 μL; pH 7) at room temperature under O₂ atmosphere for 18 h afforded 1 a in 72 % yield and 90 % conversion of the substrate. Again, low amounts of 1 b and 1 c were detected (Table 6, entry 3). Regarding the effect of the LbL coating, compound 1 a was obtained again as the main reaction product by the treatment of 1 with Tyro^S/E-LbL in 78 % yield and 85 % conversion of the substrate (Scheme 1 and Table 6, entry 4). Note that in several of the cases studied, the reactivity and selectivity of tyrosinase were found to be increased after the immobilization, which suggested a beneficial effect of the support and of the LbL coating on the activity and selectivity of the catalyst (see Table 6, entry 3 vs. entry 1). The oxidation of 1 with Tyro^E was performed under similar experimental conditions by using a higher reaction time (24 h) to afford 1 a in 60 % yield and 89 % conversion of the substrate along with a low amount of 1 b and dimers 1 c–d (Scheme 1 and Table 6, entry 5). A lower selectivity was observed by performing the oxidation in buffer, in which case 1 b and 1 c became the main reaction products (Table 6, entry 7). Tyro^E/E-LbL showed a slightly higher reactivity than did Tyro^E, which afforded 1 a as the main reaction product in 67 % yield (Table 6, entries 7 and 8). In both cases, 1 b was obtained in significant amount, which suggested a high reactivity of the catalytic systems. As there are only a few chemical procedures available to synthesize pyrogallol derivatives, the use of immobilized tyrosinase opens a possible new synthetic alternative to this family of compounds.31 Tyro^S and Tyro^E showed similar selectivities under both homogeneous and heterogeneous conditions, therefore the successive oxidations were performed only with the low-price Tyro^E. The oxidation of 2 with Tyro^E showed a higher selectivity in dichloromethane/buffer than in buffer alone to afford catechol 2 a (60 % yield), which further confirms the benign role of the organic solvent to inhibit the dimerization processes that are operative in an aqueous medium (Scheme 1 and Table 6, entry 9 vs. entry 10). Pyrogallol 2 b was isolated in low yield. Similar results were obtained during the oxidation of 2 with Tyro^E/E (Table 6, entry 11 vs. entry 12).

Notably, the highest yield of catechol was obtained with Tyro^E/E-LbL, in which case 2 a was isolated in 84 % yield along with pyrogallol 2 b (Table 6, entry 13). On the contrary, a very low selectivity was observed with Tyro^E/E-LbL in buffer (Table 6, entry 14). The efficiency of Tyro^E-based systems in the oxidation of para-alkyl substituted phenols decreased upon increasing the steric hindrance of the substituent, as evaluated in the case of bulky substituted compounds 3 and 4. Irrespective of experimental conditions, a relatively low conversion of substrate was observed to yield the corresponding catechols 3 a and 4 a as the only recovered product. In these latter cases, oxidations were performed in the presence of a low amount of CH₃CN (100 μL) to increase the solubility of substrates. Again, Tyro^E/E and Tyro^E/E-LbL were the best catalysts (except for with phenol 4) (Scheme 1 and Table 6, entries 15–20). These data are in accordance with previously reported kinetic data on the low reactivity of bulky phenols with tyrosinase in organic medium, which owes to the hydrophobic solvent reducing conformational flexibility, which in turn restricts the access of substrate at the active site of the enzyme.28 This high selectivity was further confirmed in the oxidation of 4-methoxy phenol 5 bearing an electron-donating group in the para position of the aromatic ring. In this latter case, both Tyro^E/E and Tyro^E/E-LbL showed a higher selectivity than did Tyro^E to afford catechol 5 a as the main reaction product in 77 and 60 %, yield, respectively, along with minor amounts of pyrogallol 5 b and dimer 5 c (Scheme 2 and Table 7, entries 1–3).

Scheme 2. Oxidation of phenols 5–7. Reagents and conditions: a) Tyro-based systems, O₂; b) CH₂Cl₂/buffer; c) buffer.

Table 7. Oxidation of phenols 5–9.^[a]

Entry	Substrate	Biocatalysts	Products	Conversion [%]	Yield [%]
[a] Reaction conditions: substrate (0.05 mmol) and tyrosinase (263 IU) were taken in 2.5 mL of CH2Cl2/buffer for 24 h under O2; [b] Conversion and yield were calculated by using GC–MS analysis using dodecane as the internal standard; [c] Oxidation performed in CH2Cl2/buffer/CH3CN.
1	5	Tyro^E	5a (5b) [5c]	99	37 (12) [41]
2	5	Tyro^E/E	5a (5c)	99	77 (21)
3	5	Tyro^E/E-LbL	5a (5b)	99	60 (38)
4	6	Tyro^E	6a	73	73
5	6	Tyro^E/E	6a (6b)	75	63 (11)
6	6	Tyro^E/E-LbL	6a	93	75
7	7	Tyro^E	7a	>5	traces
8	8	Tyro^E	1a	94	62
9	8	Tyro^E/E	1a (1b)	98	84 (13)
10	8	Tyro^E/E-LbL	1a (1b)	98	86 (11)
11	9	Tyro^E	9a (9b)^[b]	82	78 (4)
12	9	Tyro^E/E	9a (9b)^[b]	82	78 (3)
13	9	Tyro^E/E-LbL	9a (6b)^[b]	77	74 (3)

In the oxidation of 4-chloro phenol 6, the dimer 6 b was isolated only in the case of Tyro^E/E (Scheme 2 and Table 7, entry 4 vs. entries 5 and 6). The highest yield of 6 a was obtained with Tyro^E/E-LbL (Table 7, entry 6). On the contrary, 4-chloro-2-methyl phenol 7 showed a low reactivity toward Tyro^E to afford 7 a in negligible yield and conversion of the substrate, owing probably to the steric encumbering of the ortho substituent7 (Scheme 2 and Table 7, entry 7). This inhibition effect was not observed in the case of 3-methyl phenol 8.In this latter case, the catechol 1 a was isolated in 62–86 % yield and quantitative conversion of the substrate, with Tyro^E/E-LbL being the best catalyst (Scheme 3 and Table 7, entries 8–10). Pyrogallol 1 b was obtained as a byproduct in low yield. Finally, we evaluated the oxidation of a diphenyl methane derivative, bis(4-hydroxyphenyl)methane 9. The reaction proceeded with high conversion of the substrate to afford selectively the mono-catechol derivative 9 a in the presence of traces of the bis-catechol 9 b (identified by use of GC–MS analysis; Scheme 3 and Table 7, entries 11–13; a low amount of CH₃CN was required to increase the solubility of 9). Tyro^E and immobilized tyrosinases showed a similar reactivity. This transformation is of synthetic interest because polyhydroxylated diphenylmethane derivatives are characterized by antiviral,32 antioxidant,33 and antimicrobial activities.34 With the aim to evaluate the reusability of immobilized tyrosinases, we selected para-cresol 1 as the representative phenol derivative. Compound 1 (0.05 mmol) was oxidized with immobilized tyrosinase systems (263 IU) (Tyro^S/E, Tyro^S/E-LbL, Tyro^E/E, and Tyro^E/E-LbL) in the dichloromethane/buffer medium under previously reported experimental conditions. The oxidations were followed spectrophotometrically at 389 nm. After 30 min, the immobilized biocatalyst was recovered, washed, and reused with a freshly added substrate. For successive runs, the enzyme activity measured in the first oxidation was used as the reference value. One unit activity (IU) was defined as the increase of 0.001 min⁻¹ in absorbance at 389 nm, 25 °C, in dichloromethane/sodium phosphate buffer (0.1 m, pH 7). Enzyme recycling in organic and aqueous media was compared. As shown in Table 8, the immobilized Tyro^S systems retained significant activity after 5 runs, which was higher in the dichloromethane/buffer medium than in the aqueous medium (Table 8, entries 1–5). The polyelectrolyte coating stabilized the enzyme further (Table 8, column 1 vs. column 2). Similar results were obtained with Tyro^E-based systems (Table 8, entries 1–5, columns 3 and 4). The general trend showed LbL-based biocatalysts on Eupergit C 250 L systems to be more stable and maintain higher activity. The general trend showed LbL-base biocatalyst on Eupergit C 250L system to be more stable and maintain higher activity than Eupergit alone. This behavior suggests that microcapsules create an internal microenvironment able to protect the enzyme from denaturing agents.35 Moreover, microcapsules may keep water inside, which ensures the hydration required for enzyme activity and stability.36

Scheme 3. Oxidation of phenols 8 and 9. Reagents and conditions: a) Tyro-based systems, O₂; b) CH₂Cl₂/buffer; c) buffer; d) CH₂Cl₂/buffer/CH₃CN; e) buffer/CH₃CN.

Table 8. Reusability of Tyro-immobilized systems.^[a]

Times reused	Tyro^S/E		Tyro^S/E-LbL		Tyro^E/E		Tyro^E/E-LbL
	CH2Cl2	Buffer	CH2Cl2	Buffer	CH2Cl2	Buffer	CH2Cl2	Buffer
[a] Reusability is expressed as the percentage of activity in each run with respect to that measured in the first reference oxidation.
1	84	72	86	74	88	68	90	70
2	70	38	79	54	79	52	81	60
3	50	12	59	23	55	20	62	27
4	39	8	47	10	41	9	45	10
5	32	4	37	7	32	3	38	5

Conclusions

Tyrosinase was found to be an efficient catalyst for the oxidation of phenols to the corresponding catechols under mild experimental conditions in an organic medium. The enzyme retained the catalytic activity and selectivity after immobilization on Eupergit C 250 L and successive coating by the LbL technique. Even if the free enzymes afford better kinetic parameters, in some of the cases studied, immobilized systems were more selective than the parent enzyme to yield catechols, which suggested a stabilization effect exerted by the support. This pattern was further strengthened by recycling experiments, which showed that the best stabilization effect was in the presence of the polyelectrolyte coating. Moreover, the immobilized systems were more stable in an organic medium than in water. It is well known that polyelectrolytes can influence the activity of oxidative enzymes, for example, by acting as surfactants or stabilizing the enzyme in a right conformation.37 These effects can be further modulated by the presence of an organic solvent. Although we have not studied in detail the mechanism by which polyelectrolytes poly(sodium 4-styrenesulfonate) (PSS) and poly(allylamine hydrochloride) (PAH) may affect the selectivity of tyrosinase, in agreement with data previously reported in the literature, it is reasonable to suggest that they may inhibit the dimerization of quinones formed by secondary processes.38 Regarding the qualitative structure–activity relationships, para- and meta-substituted phenols were efficiently oxidized whereas no reactivity was observed in the case of ortho-substituted derivatives. This inhibition owed probably to steric hindrance of the ortho substituent in the formation of the first complex intermediate with the copper atom at the active site of the enzyme. As catechols are biologically active compounds difficult to synthesize by traditional chemical methods under environmentally friendly conditions, the use of immobilized tyrosinases open a novel synthetic alternative for this interesting family of substances.

Experimental Section

Mushroom tyrosinase was either purchased from Sigma-Aldrich (Tyro^S) or extracted (Tyro^E) from Agaricus bisporus. Eupergit C 250 L, PSS (MW 70 000 g mol⁻¹), PAH (MW 56 000 g mol⁻¹), L-Tyr, ABTS, bovine serum albumin (BSA), ammonium sulfate [(NH₄)₂SO₄], dichloromethane (CH₂Cl₂), ethyl acetate (EtOAc), petroleum ether, acetonitrile (CH₃CN), sodium sulfite (Na₂SO₃), sodium sulfate anhydrous (Na₂SO₄), dodecane, pyridine, hexamethyldisilazane (HMDS), trimethylchlorosilane (TMCS), and phenols were purchased from Sigma-Aldrich. Water used during extraction was degassed and stored at 4 °C until use. All spectrophotometric measurements were made with a Varian Cary50 UV–Vis spectrophotometer equipped with a Peltier thermostatted single cell holder. Dichloromethane was dried on anhydrous sodium sulfate prior to use. All experiments were done in triplicate using free and immobilized Sigma-Aldrich tyrosinase (Tyro^S) and free and immobilized extracted tyrosinase (Tyro^E) in the dichloromethane/buffer system and in an aqueous medium. Buffer used was a sodium phosphate buffer (sodium phosphate, 0.1 m, pH 7).

Extraction of tyrosinase

The mushroom tyrosinase was partially purified from commercial A. bisporus.17 In a typical procedure, the sporocarps were cleaned to remove earthy residues and then washed with ascorbic acid (20 mM) maintained at 4 °C. After they had been dried, the sporocarps were sliced and frozen at −20 °C at least 1 day before extraction. The frozen sporocarps (1 kg) were homogenized twice in acetone (1.2 L) at −20 °C in a blender for 1 min. The obtained solid pulp was filtered through a Buchner funnel and homogenized with acetone (1.0 L, 30 % v/v) in water for 2–3 min. The mixture was centrifuged at 9000g for 20 min. To the supernatant, 1.5 volumes of acetone at −20 °C were added dropwise under vigorous stirring. The mixture was allowed to settle at 4 °C for 2–3 h; most of the supernatant fluid was decanted and discarded, the remainder was centrifuged, and the precipitate was dissolved in water and subjected to precipitation with calcium acetate 1 % saturation solution. The turbid mixture was frozen at −20 °C. Samples obtained from several days were stored frozen at this stage. They were then thawed, mixed, and centrifuged. Ammonium sulfate [(NH₄)₂SO₄] powder was added to the collected supernatant to make a 35 % saturated solution. The resulting solution was allowed to stand for 30 min at 4 °C and centrifuged at 9000g for 20 min. (NH₄)₂SO₄ was added to the supernatant to make a 70 % saturated solution. The solution was allowed to stand for 2 h at 4 °C and centrifuged. The precipitate was dissolved in a minimal volume of cold water and then dialyzed against water and concentrated by means of Vivaflow 50 equipped with a polyethersulfone membrane (10 000 molecular weight cut off). The resulting enzyme solution was lyophilized and stored at −20 °C.

Tyrosinase immobilization on Eupergit C 250 L

The enzyme immobilization was performed by a modification of literature procedures.21a, b Dry Eupergit C 250 L (1.0 g) was added to sodium phosphate buffer (0.1 M, pH 7, 8.0 mL) containing tyrosinase (5.0 mg, 69 405 IU for Tyro^S and 12 840 IU for Tyro^E). The mixture was incubated for 24 h at room temperature with orbital shaking. At the end of the coupling period, the beads were filtered and washed (5×8 mL) with sodium phosphate buffer (0.1 M, pH 7) until no activity was detected in the washing. The obtained beads were incubated with glycine (3.0 M) for 2 h to block residual epoxy groups39 and then washed with buffer and finally air dried and stored at 4 °C. The amount in milligrams and the units of coupled tyrosinase (Tyro^S/E and Tyro^E/E) were calculated by the difference between the amount/units loaded and that recovered in the washings by conventional Bradford and activity assay.

Tyrosinase immobilization on Eupergit C 250 L coated using the LbL method

Both Tyro^S/E and Tyro^E/E, synthesized as described above, were coated with the LbL method by a modification of literature procedures.40 PAH and PSS solutions (2 mg mL⁻¹ in 0.5 M NaCl) were added alternately and swiftly to Eupergit-supported tyrosinase: each polyelectrolyte layer was adsorbed for 20 min at room temperature with orbital shaking and then washed with NaCl (0.5 M). The deposition of polyelectrolytes was repeated three times (in the sequence PAH–PSS–PAH) and the obtained immobilized tyrosinases (Tyro^S/E-LbL and Tyro^E/E-LbL) were air dried and stored at 4 °C.

Determination of protein concentration

Protein concentration was determined spectrophotometrically at 595 nm according to the Bradford method using BSA as a standard.41

Activity assay

Tyrosinase assay was performed by using the dopachrome method as previously described.18 The L-Tyr solution (1 mL, 2.5 mM) in water was mixed briefly with sodium phosphate buffer (0.1 M, pH 7, 1.9 mL) and incubated at 25 °C for 10 min. Then, an appropriate amount of free or immobilized enzyme in sodium phosphate buffer (100 μL) was added to the mixture and the initial rate was measured immediately as a linear increase in the optical density at 475 nm owing to dopachrome production. One unit of enzyme activity was defined as the increase of 0.001 min⁻¹ in absorbance at pH 7, 25 °C, in a reaction mixture (3.0 mL) containing L-Tyr (1.0 ml, 2.5 mM) and sodium phosphate buffer (2.0 ml, 0.1 M, pH 7). As tyrosinase from A. bisporus contains a low amount of laccase, the laccase activity was determined spectrophotometrically by using ABTS as the substrate.19 The assay mixture contained ABTS (0.5 mM) and sodium phosphate buffer (0.1 M, pH 7), and the enzyme was incubated at 25 °C. The oxidation was followed by an absorbance increase at 415 nm for 1 min. One activity unit was defined as the amount of enzyme that oxidized 1 μmol_ABTS min⁻¹.

Optimization of water requirement

para-Cresol 1 was used as the standard phenolic substrate.10a Compound 1 (5 mg, 0.046 mmol), biocatalysts (2.69 μg Tyro^S, 16 μg Tyro^S/E or Tyro^S/E-LbL, 12 μg Tyro^E, and 7.5 μg Tyro^E/E or Tyro^E/E-LbL), and dichloromethane (2.5 mL) were placed in vials at 25 °C under O₂, to which sodium phosphate buffer (0.1 M, pH 7, 10–70 μL) was added. The reaction mixture was stirred vigorously for 30 min. Every 15 min, aliquots were removed and their absorbance at 389 nm, owing to ortho-quinone production, was measured; the aliquots were returned to the vials as rapidly as possible. One unit of enzyme activity (IU) was defined as an increase in absorbance of 0.001 min⁻¹ at 389 nm, 25 °C, sodium phosphate buffer (0.1 M, pH 7).

Kinetic assay

Kinetic parameters (K_m, V_max, and V_max/K_m) were determined by measuring enzyme activity at different concentrations of substrate and plotting data to a Lineweaver–Burk plot.20 To study the catalytic properties of enzymes in the organic solvent media, the same procedure was followed as for the optimization of hydration described above, using different concentrations of para-cresol 1 (2–15 mM) and optimum concentration of aqueous buffer. Absorbance was measured at 389 nm as described above. Reactions in the aqueous system were performed under similar experimental conditions.5 To compare the catalytic properties of commercial and extracted tyrosinase, the same procedure as for the activity assay was followed, using different concentrations of L-Tyr (330–1000 μM), 53 μg of Tyro^S and Tyro^E, and 88 μg of immobilized Tyro^S and Tyro^E. Absorbance was measured at 475 nm as described above.

Enzyme recycling

For recycling, para-cresol (5 mg), the immobilized Tyro enzyme (10 mg), optimal buffer, and dichloromethane (2.5 mL) were placed in vials at 25 °C under O₂. The reaction mixture was shaken vigorously for 30 min. Every 15 min, aliquots were removed to measure their absorbance at 389 nm and returned to the vials as quickly as possible. After 30 min, the biocatalyst was washed, recycled, and reused again. Reactions in the aqueous system were performed under similar experimental conditions. For each run, tyrosinase activity was expressed as a percentage activity relative to the value of the first run.

Phenol oxidation

A large range of phenols (Figure 6) has been oxidized, including para-monosubstituted phenols para-cresol 1, 4-ethyl phenol 2, 4-tert-butyl phenol 3, 4-sec-butyl phenol 4, 4-methoxy phenol 5, and 4-chloro phenol 6; ortho, para-disubstituted phenol 4-chloro-2-methyl phenol 7; meta-substituted phenol meta-cresol 8 and the bisphenol methane derivative bis(4-hydroxyphenyl)methane 9. The reactions were performed under both homogeneous and heterogeneous conditions in dichloromethane/buffer and in a reference aqueous system. As a general procedure, the appropriate phenol (0.05 mmol), tyrosinases (600–2400 IU), and the optimal amount of sodium phosphate buffer (0.1 M, pH 7) were placed in dichloromethane (2.5 mL) at 25 °C under O₂ and vigorous stirring. For insoluble aqueous phenols 3, 4, and 9, substrates were dissolved in CH₃CN (100 μL) and then added to dichloromethane/buffer solutions. Reactions in the aqueous system were performed under similar experimental conditions. The reaction was monitored by use of TLC. After the disappearance of the substrate, different workup procedures were used depending on the reaction conditions. The enzyme was recovered from an organic medium by decantation (free enzymes) or filtration (immobilized enzymes) and the organic layer was treated with an equal volume of sodium sulfite solution (1 % w/w) to reduce benzoquinones to catechols.5 The mixture was stirred for 5 min, and the phases were separated. The aqueous phase was acidified with HCl (1.0 n) and extracted twice with EtOAc. The organic extracts (CH₂Cl₂ and EtOAc) were treated with a saturated solution of NaCl and dried over anhydrous Na₂SO₄ before being filtered and concentrated under vacuum to yield a coloured crude product. If an aqueous medium was used, sodium sulfite was added to the reaction mixture and stirred for 5 min. The solution was then acidified with HCl (1.0 n), extracted twice with EtOAc, and the organic phase treated as described above. The obtained coloured residue was then analyzed by using GC–MS analysis. The residue was treated with anhydrous pyridine (kept over NaOH pellets) and HMDS/TMCS in a 2:1 v/v ratio under vigorous stirring at RT for 30 min and then allowed to stand for 5 min.42

Identification and characterization of oxidation products

All products were identified by using ¹H NMR, ¹³C NMR, and GC–MS. ¹H NMR and ¹³C NMR were recorded on a Bruker 200 MHz spectrometer with CDCl₃ as the solvent. GC–MS analysis was performed on a GC–MS QP5050 Shimadzu apparatus using an SPB column (25 m×0.25 mm and 0.25 mm film thickness) and an isothermal temperature profile of 100 °C for 2 min, followed by a 10 °C min⁻¹ temperature gradient to 280 °C for 25 min. The injector temperature was 280 °C. Chromatography-grade helium was used as the carrier gas with a flow of 2.7 mL min⁻¹. Mass spectra were recorded with an electron beam of 70 eV. Quantitative analyses were performed by using dodecane as the internal standard.

4-Methylcatechol (4-methyl-1,2-benzenediol), 1 a

Oil; ¹H NMR43 (200 MHz, CDCl₃): δ_H=2.24 (3 H, s, CH₃), 5.04 (1 H, br s, OH), 5.18 (1 H, br s, OH), 6.61–6.76 ppm (3 H, m, PhH); ¹³C NMR43 (50 MHz, CDCl₃): δ_C=20.8 (CH₃), 115.3 (CH), 116.2 (CH), 121.5 (CH), 131.1 (C), 141.0 (C), 143.3 ppm (C); MS: m/z 268 [M⁺], 253 [M−CH₃], 238 [M−(CH₃)₂], 223 [M−(CH₃)₃], 195 [M−Si(CH₃)₃], 179 [M−OSi(CH₃)₃], 164 [M−OSi(CH₃)₄], 149 [M−OSi(CH₃)₅], 134 [M−OSi(CH₃)₆], 106 [M−OSi₂(CH₃)₆], 90 [M−O₂Si₂(CH₃)₆].

5-Methylpyrogallol (5-methyl-1,2,3-benzenetriol), 1 b

Oil; ¹H NMR44 (200 MHz, CDCl₃): δ_H=2.20 (3 H, s, CH₃), 5.00 (1 H, s), 5.05 (1 H, s), 8.20 ppm (3 H, s, OH); ¹³C NMR (50 MHz, CDCl₃): δ_C=22.1 (CH₃), 109.0 (2×CH), 131.2 (C), 137.1(C), 146.1 ppm (2×C); MS: m/z 356 [M⁺], 341 [M−CH₃], 313 [M−(CH₃)₃], 283 [M−Si(CH₃)₃], 267 [M−OSi(CH₃)₃], 252 [M−OSi(CH₃)₄], 237 [M−OSi(CH₃)₅].

Dimer, 1 c

Oil; ¹H NMR (200 MHz, CDCl₃): δ_H=2.19 (3 H, s, CH₃), 2.42 (3 H, s, CH₃), 6.51 (3 H, bs, OH), 6.61–7.12 ppm (5 H, m, PhH); ¹³C NMR (50 MHz, CDCl₃): δ_C=16.0 (CH₃), 22.1 (CH₃), 116.6 (CH), 118.2 (CH), 122.0 (C), 125.6 (C), 127.0 (CH), 127.2 (C), 131.2 (CH), 133.1 (CH), 136.7 (C), 141.3 (C), 150.1 (C), 158.2 ppm (C); MS: m/z 446 [M⁺], 431 [M−CH₃], 329 [M−Si(CH₃)₃], 313 [M−OSi(CH₃)₃], 298 [M−OSi(CH₃)₄], 268 [M−OSi(CH₃)₆], 180 [M−O₂Si₂(CH₃)₆].

5,5′-Dimethyl-[1,1′-biphenyl]-2,2′,3,3′-tetrol, 1 d

Oil; ¹H NMR (200 MHz, CDCl₃): δ_H=2.41 (6 H, s, 2 CH₃), 6.62–6.91 ppm (4 H, m, PhH); ¹³C NMR (50 MHz, CDCl₃): δ_C=22.1 (2 CH₃), 117.2 (2 CH), 122.1 (2 C), 132.0 (2 CH), 138.1 (2 C), 141.3 (2 C), 150.3 ppm (2 C); MS: m/z 534 [M⁺], 519 [M−CH₃], 417 [M−Si(CH₃)₃], 401 [M−OSi(CH₃)₃], 386 [M−OSi(CH₃)₄], 371 [M−OSi(CH₃)₅], 268 [M−O₂Si₂(CH₃)₆].

4-Ethylcatechol (4-ethyl-1,2-benzenediol), 2 a

Oil; ¹H NMR45 (200 MHz, CDCl₃): δ_H=1.04 (3 H, m, CH₃), 2.36 (2 H, m, CH₂), 6.00–7.25 ppm (3 H, m, PhH); ¹³C NMR (50 MHz, CDCl₃): δ_C=15.2 (CH₃), 28.1 (CH₂), 116.5 (CH), 117.4 (CH), 124.2 (CH), 139.3 (C), 145.7 (C), 148.4 ppm (C); MS: m/z 282 [M⁺], 267 [M−CH₃], 252 [M−(CH₃)₂], 237 [M−(CH₃)₃], 209 [M−Si(CH₃)₃], 193 [M−OSi(CH₃)₃], 179 [M−OSi(CH₃)₄], 164 [M−OSi(CH₃)₅], 148 [M−OSi(CH₃)₆], 120 [M−OSi₂(CH₃)₆].

5-Ethyl-1,2,3-benzenetriol, 2 b

Oil; ¹H NMR (200 MHz, CDCl₃): δ_H=1.22 (3 H, m, CH₃), 2.62 (2 H, m, CH₂), 6.62 (2 H, m, PhH), 8.14 ppm (3 H, bs, OH); ¹³C NMR (50 MHz, CDCl₃): δ_C=15.2 (CH₃), 29.3 (CH₂), 111.1 (2 CH), 136.2 (C), 138.0 (C), 142.3 ppm (2 C); MS: m/z 370 [M⁺], 355 [M−CH₃], 282 [M−OSi(CH₃)₃], 267 [M−OSi(CH₃)₅], 251 [M−OSi(CH₃)₆], 209 [M−O₂Si₂(CH₃)₆], 194 [M−O₂Si₂(CH₃)₉].

Dimer 2 c

Oil; ¹H NMR (200 MHz, CDCl₃): δ_H=1.21 (3 H, m, CH₃), 1.32 (3 H, m, CH₃), 2.71 (2 H, m, CH₂), 2.82 (2 H, m, CH₂), 6.4–7.2 ppm (5 H, m, PhH); ¹³C NMR (50 MHz, CDCl₃): δ_C=15.2 (CH₃), 15.7 (CH₃), 27.2 (CH₂), 28.3 (CH₂), 106.4 (C), 107.2 (C), 112.3 (CH), 114.1 (CH), 128.3 (C), 130.1 (CH), 134.1 (CH), 134.6 (C), 135.1 (C), 136.2 (CH), 143.1 (C), 155.2 ppm (C); MS: m/z 474 [M⁺], 459 [M−CH₃], 429 [M−CH₃)₃], 341 [M−OSi(CH₃)₃], 326 [M−OSi(CH₃)₄], 311 [M−OSi(CH₃)₅].

5,5′-Diethyl-[1,1′-biphenyl]-2,2′,3,3′-tetrol, 2 d

Oil; ¹H NMR (200 MHz, CDCl₃): δ_H=1.22 (6 H, m, 2 CH₃), 2.73 (4 H, m, 2 CH₂), 6.4–7.2 ppm (4 H, m, PhH); ¹³C NMR (50 MHz, CDCl₃): δ_C=15.6 (2 CH3), 28.3 (2 CH₂), 102.2 (2 C), 114.0 (2 CH), 134.3 (2 C), 136.1 (2 C), 138.3 (2 CH), 143.2 ppm (2 C); MS: m/z 562 [M⁺], 517 [M−CH₃)₃], 489 [M−Si(CH₃)₃], 474 [M−OSi(CH₃)₃], 459 [M−OSi(CH₃)₄], 444 [M−OSi(CH₃)₅], 430 [M−OSi(CH₃)₆], 385 [M−O₂Si₂(CH₃)₆].

4-tert-Butylcatechol (4-tert-butylbenzene-1,2-diol), 3 a

Oil; ¹H NMR (200 MHz, CDCl₃): δ_H=1.33 (9 H, s, CH₃), 6.63–7.11 ppm (3 H, m, PhH); ¹³C NMR (50 MHz, CDCl₃): δ_C=31.2 (3 CH₃), 34.5 (C), 116.5 (CH), 116.9 (CH), 122 (CH), 144.3 (C), 146.2 (C), 147.1 ppm (C); MS: m/z 310 [M⁺], 295 [M−CH₃], 280 [M−(CH₃)₂], 265 [M−(CH₃)₃], 237 [M−Si(CH₃)₃], 222 [M−OSi(CH₃)₃], 207 [M−OSi(CH₃)₄], 192 [M−OSi(CH₃)₅], 176 [M−OSi(CH₃)₆], 148 [M−OSi₂(CH₃)₆].

4-sec-Butylcatechol (4-(1-methylpropyl)-1,2-benzenediol), 4 a

Oil; ¹H NMR (200 MHz, CDCl₃): δ_H=1.10 (3 H, m, CH₃), 1.22 (3 H, m, CH₃), 1.53 (2 H, m, CH₂), 3.23 (1 H, m, CH), 6.52–6.84 ppm (3 H, m, PhH); ¹³C NMR (50 MHz, CDCl₃): δ_C=11.2 (CH₃), 22.3 (CH₃), 31.2 (CH₂), 43.1 (CH), 113.3 (CH), 114.1 (CH), 124.4 (CH), 136.2 (C), 145.1 (C), 147.0 ppm (C); MS: m/z 310 [M⁺], 295 [M−CH₃], 280 [M−(CH₃)₂], 237 [M−Si(CH₃)₃], 222 [M−OSi(CH₃)₃], 207 [M−OSi(CH₃)₄], 192 [M−OSi(CH₃)₅], 149 [M−OSi₂(CH₃)₆], 133 [M−O₂Si₂(CH₃)₆].

4-Methoxycatechol (4-methoxy-1,2-benzenediol), 5 a

Oil; ¹H NMR (200 MHz, CDCl₃): δ_H=3.72 (3 H, s, CH₃), 6.41–6.73 (3 H, m, PhH), 6.94 ppm (2 H, bs, OH); ¹³C NMR (50 MHz, CDCl₃): δ_C=56.3 (CH₃), 101.4 (CH), 108.3 (CH), 115.6 (CH), 140.4 (C), 146.0 (C), 154.4 ppm (C); MS: m/z 284 [M⁺], 269 [M−CH₃], 254 [M−(CH₃)₂], 239 [M−(CH₃)₃], 196 [M−OSi(CH₃)₃], 181 [M−OSi(CH₃)₄], 166 [M−OSi(CH₃)₅], 150 [M−OSi(CH₃)₆].

5-Methoxy-1,2,3-benzenetriol, 5 b

Oil; ¹H NMR (200 MHz, CDCl₃): δ_H=3.71 (3 H, s, CH₃), 6.10 (2 H, s, PhH), 9.11 ppm (3 H, bs, OH); ¹³C NMR (50 MHz, CDCl₃): δ_C=57.3 (CH₃), 95.2 (2 CH), 131.5 (C), 147.3 (2 C), 151.4 ppm (C); MS: m/z 372 [M⁺], 357 [M−CH₃] 342 [M−(CH₃)₂], 255 [M−Si(CH₃)₃], 224 [M−OSi(CH₃)₄], 209 [M−OSi(CH₃)₅], 194 [M−OSi(CH₃)₆].

Dimer 5 c

Oil; ¹H NMR (200 MHz, CDCl₃): δ_H=3.32 (3 H, s, CH₃), 3.74 (3 H, s, CH₃), 6.23–6.72 (3 H, m, PhH), 6.94 (3 H, s, OH), 7.10–7.32 ppm (2 H, m, PhH); ¹³C NMR (50 MHz CDCl₃): δ_C=55.3 (2 CH₃), 103.4 (CH), 111.2 (CH), 114.5 (CH), 117.0 (CH), 117.5 (C), 120.3 (CH), 121.2 (C), 136.4 (C), 150.2 (C), 154.3 (C), 156.0 (C), 157.9 ppm (C); MS: m/z 478 [M⁺], 432 [M−(OCH₃)₂], 401 [M−(O₂(CH₃)₃], 386 [M−(O₂(CH₃)₄], 371 [M−(O₂(CH₃)₅], 343 [M−(O₂Si(CH₃)₅], 328 [M−(O₃Si(CH₃)₅], 313 [M−(O₃Si(CH₃)₆], 270 [M−(O₃Si₂(CH₃)₇].

4-Chlorocatechol (4-chloro-1,2-benzenediol), 6 a

Oil; ¹H NMR (200 MHz, CDCl₃): δ_H=6.72–6.83 (3 H, m, PhH), 8.10 ppm (2 H, s, OH); ¹³C NMR (50 MHz, CDCl₃): δ_C=113.0 (CH), 117.3 (CH), 125.4 (CH), 127.2 (C), 145.0 (C), 152.3 ppm (C); MS: m/z 288 [M⁺], 273 [M−CH₃], 258 [M−(CH₃)₂], 243 [M−(CH₃)₃], 215 [M−Si(CH₃)₃], 199 [M−OSi(CH₃)₃], 184 [M−OSi(CH₃)₄], 169 [M−OSi(CH₃)₅], 126 [M−OSi₂(CH₃)₆].

5,5′-Dichloro-(1,1′-Biphenyl)-2,2′,3-triol, 6 b

Oil; ¹H NMR (200 MHz, CDCl₃): δ_H=6.71–7.23 (5 H, m, PhH), 7.62 ppm (3 H, s, OH); ¹³C NMR (50 MHz, CDCl₃): δ_C=111.2 (C), 114.2 (C), 114.5 (CH), 117.3 (CH), 126.5 (C), 127.4 (C), 129.3 (CH), 130.2 (CH), 135.1 (CH), 137.2 (C), 152.3 (C), 155.0 ppm (C); MS: m/z 486 [M⁺], 471 [M−CH₃], 353 [M−OSi(CH₃)₃], 338 [M−OSi(CH₃)₄], 236 [M−OSi₂(CH₃)₆].

5-Chloro-3-methylcatechol (5-chloro-3-methyl-1,2-benzenediol), 7 a

Oil; ¹H NMR (200 MHz, CDCl₃): δ_H=2.10 (3 H, s, CH₃), 6.53–6.74 (2 H, s, PhH), 7.52 ppm (2 H, s, OH); ¹³C NMR (50 MHz CDCl₃): δ_C=17.2 (CH₃), 116.3 (CH), 123.2 (C), 125.4 (C), 126.3 (CH), 140.2 (C), 151.0 ppm (C); MS: m/z 302 [M⁺], 287 [M−CH₃], 272 [M−(CH₃)₂], 229 [M−Si(CH₃)₃], 213 [M−OSi(CH₃)₃], 198 [M−OSi(CH₃)₄], 168 [M−OSi(CH₃)₆].

4-(para-Hydroxybenzyl)-pyrocatechol (4-[(4-hydroxyphenyl)methyl]-1,2-benzenediol), 9 a

Oil; ¹H NMR (200 MHz, CDCl₃): δ_H=4.10 (2 H, m, CH₂), 6.51–7.22 ppm (7 H, m, PhH); ¹³C NMR (50 MHz, CDCl₃): δ_C=42.2 (CH), 115.5 (CH), 116.1 (2 CH), 116.5 (CH), 121.3 (CH), 130.2 (2 C), 133.5 (C), 134.5 (C), 144.2 (C), 145.3 (C), 156.1 ppm (C); MS: m/z 432 [M⁺], 417 [M−CH₃], 402 [M−(CH₃)₂], 359 [M−Si(CH₃)₃], 343 [M−OSi(CH₃)₃], 329 [M−OSi(CH₃)₄], 314 [M−OSi(CH₃)₅], 298 [M−OSi(CH₃)₆].

4,4′-Methylenedipyrocatechol (4,4′-methylenebis-1,2-benzenediol), 9 b

MS: m/z 520 [M⁺], 505 [M−CH₃], 447 [M−Si(CH₃)₃], 431 [M−OSi(CH₃)₃], 417 [M−OSi(CH₃)₄], 343 [M−O₂Si₂(CH₃)₆], 329 [M−O₂Si₂(CH₃)₇].

Acknowledgements

Italian Miur PRIN 2008 is acknowledged.