Polymer-Assisted Biocatalysis: Effects of Macromolecular Architectures on the Stability and Catalytic Activity of Immobilized Enzymes toward Water-Soluble and Water-Insoluble Substrates.

The aim of this study is to develop efficient enzyme immobilization media that will enable the reuse of the biocatalysts over multiple cycles, increase their thermal stability, and attenuate their activity toward hydrophobic substrates for "green" transformations in aqueous media. For this purpose, amphiphilic AB and ABA block copolymers were synthesized and tested with laccase (a multicopper oxidase). In all cases, the hydrophilic B block consisted of poly(ethylene glycol), PEG, with molecular masses of 3, 5, 13, 20, or 13 kDa poly(ethylene oxide). The hydrophobic A blocks were made of linear poly(styrene), PS; hyperbranched poly(p-chloromethyl styrene); or dendritic poly(benzyl ether)s of generations 2, 3, and 4 (G2, G3, and G4) with molecular masses ranging from 1 to 24 kDa. A total of 23 different copolymers (self-assembling into micelles or physical networks) were evaluated. Notable activity enhancements were achieved with both micelles (up to 253%) and hydrogels (up to 408%). The highest enzymatic activity and thermal stability were observed with laccase immobilized in hydrogels consisting of the linear ABA block copolymer PS2.7k-PEG3k-PS2.7k (13 290 μkat/L, 65 °C, ABTS test). This represents a 1245% improvement over native laccase at the same conditions. At 25 °C, the same complex showed a 1236% higher activity than the enzyme. The highest polymerization yield for a water-insoluble monomer was achieved with laccase immobilized in hydrogels composed of linear-dendritic ABA copolymer G3-PEG5k-G3 (85.5%, 45 °C, tyrosine monomer). The broad substrate specificity and reusability of the immobilized laccase were also demonstrated by the successful discoloration of bromophenol blue, methyl orange, and rhodamine B over eight repetitive cycles.

Introduction

The implementation of enzymes in synthetic chemistry is often viewed as very lucrative because of their high catalytic activity, substrate specificity, stereo- and regiospecificity, and lack of byproducts.[1,2] Among enzymes currently attracting significant interest is laccase, a benzenediol:oxygen oxidoreductase containing a multicopper core.[3,4] Having a broad substrate specificity, laccase plays a multifaceted role in biological synthesis including pigment formation, lignin degradation, and cellular repair.[5] Synthetically, laccases are well-known for their ability to polymerize phenol and phenol derivatives via oxidizing the phenol substrate with the concomitant reduction of oxygen to water.[3,6] This polymerization occurs through the production of phenoxy radicals which can then undergo C–C or C–O coupling at the ortho- and/or para-position. This ability to easily oxidize phenols has allowed laccase to find a possible niche for environmental cleanup from wastewaters of phenolic substances, such as phenol and catechol, and synthetic dyes.[7−9] Laccases have also found use as catalysts for oxidative transformations of nonphenolic small molecules, especially in the presence of radical mediators. Such reactions include oxidation of alcohols to aldehydes and ketones, coupling of phenols and steroids, and carbon–nitrogen bond formation.[1−3]

Regardless of the intended use, reactions involving laccase as a catalyst are continually inhibited by several common factors, namely, the relatively high cost of laccase, activity limited to substrates soluble in water, and low enzyme activity, resulting in prolonged reaction times and reduced yields.[10] To circumvent these problems, modifications to the reaction conditions are often made such as the addition of an organic cosolvent to increase the solubility of the substrate and the immobilization of the enzyme to afford reusability after each reaction cycle.[4,11] However, organic additives often lead to a decreased enzymatic activity over time periods typical of an enzymatic reaction due to denaturation.[12] Enzyme modifications such as immobilization have also habitually resulted in a decreased enzymatic activity because of severe restrictions in enzyme mobility in addition to the issues involving active site orientation.[4]

Previously published work from our group[13] showed that laccase was able to form complexes with amphiphilic linear–dendritic and dendritic–linear–dendritic copolymers composed of benzyl ether dendritic moieties and linear poly(ethylene glycol), PEG. Complexation of laccase with both types of copolymers via π–H interactions resulted in an increased enzymatic activity toward water-soluble and water-insoluble substrates. This enabled the unprecedented transformation of C60 in water at very mild reaction conditions.[14] This concept was further developed to produce an unnatural poly(amino acid) directly from tyrosine.[15] Although linear–dendritic and dendritic–linear–dendritic copolymer architectures have proven to increase the laccase activity and reaction yields, their expense and time-intensive synthesis will most likely limit their use in industrial applications. It is therefore the aim of this study to find alternative complexing agents which use similar mechanisms of complexation and which can be produced at significantly reduced costs in a two-step synthesis.

Complexing agents of interest include linear diblock poly(styrene) (PS)–PEG and triblock PS–PEG–PS copolymers composed of PEG and PS and linear–hyperbranched diblock and hyperbranched–linear–hyperbranched triblock copolymers composed of the same linear PEG and hyperbranched poly(p-chloromethylstyrene) (PPCMS). Copolymers capable of forming hydrogels and micelles (Figure ) will be synthesized because both groups offer practical advantages: hydrogel complexes with the encapsulated enzyme could be recycled, whereas micelles enable a greater substrate diffusivity. Because of the ease at which molecular mass (MM) can be controlled, atom transfer radical polymerization (ATRP)[16−18] will be employed to synthesize these new complexing agents. MM control via ATRP will allow for the synthesis of amphiphilic copolymers capable of self-assembling into either micelles or hydrogels by controlling the hydrophilic/hydrophobic (PEG/PS and PEG/PPCMS) weight ratios.

Figure 1

Schematic illustration of enzyme complexation using micellar complexes (A) and hydrogel complexes (B–D) with dendritic (red), hyperbranched (blue), and linear (green) copolymers.

Experimental Section

Materials

dl-Tyrosine (99+%), 2,2′-azino-bis(3-ethylbenzo-thiazoline-6-sulfonic acid) (ABTS, 98+%), 1-hydroxybenzotriazole (HOBT, 95+%), bromophenol blue (95%), rhodamine B base (97%), methyl orange (95+%), and p-nitrophenyl palmitate (all from Sigma-Aldrich, St. Louis MO) were used as received. Chlorobenzene (99+%, Acros) was degassed with argon and stored over 4 Å molecular sieves. Laccase was produced and used as previously described.[19] All linear–dendritic and dendritic–linear–dendritic copolymers were synthesized by the coupling of preformed reactive fragments[20] or via living ring-opening anionic polymerization of ethylene oxide initiated by third-generation benzyl ether dendrons.[21] All linear–linear and linear–hyperbranched copolymers were produced by controlled radical polymerization (ATRP). Details are provided in the Supporting Information.

Methods

NMR Spectroscopy

1H NMR and 13C NMR spectra were recorded using CDCl3 as a solvent at 22 °C with a Bruker AVANCE 600 MHz instrument.

Dynamic Light Scattering

All dynamic light scattering (DLS) measurements were performed on a Malvern Zetasizer ZS instrument. The instrument used a 633 nm laser source with a fixed backscattering detector at 173°. Size calculations were performed using a CONTIN procedure.

UV–Vis Spectroscopy

All UV–vis spectroscopic measurements for the enzymatic activity and decolorization experiments were conducted on an Agilent 8453 UV–vis spectrophotometer at room temperature in deionized (DI) water.

Immobilization of Enzymes via Solvent Exchange

Enzyme complexation with all copolymers was conducted by first dissolving 20 ± 0.5 mg of the desired copolymer in 200 μL of tetrahydrofuran (THF). This solution was then added dropwise to a vigorously stirring solution of laccase, previously diluted to 77 μkat/L. The complexed laccase solution was then allowed to equilibrate for 4 h. All complexed laccase solutions were prepared immediately before use.

Scanning Electron Microscopy

Scanning electron microscopy (SEM) images were obtained by using a JEOL-5800-LV SEM instrument with an acceleration voltage of 15 kV. The swollen hydrogel samples were immersed in water for 48 h and then quickly frozen with liquid nitrogen. The frozen hydrogels were subjected to cryofracturing followed by lyophilization. The dry samples were sputter-coated with gold for SEM analysis.

Enzyme Activity Assay

Activity assays of native laccase and laccase complexes were conducted using ABTS as a substrate at room temperature. ABTS (5 mM) in DI water was freshly prepared before each assay. After blanking the UV–vis spectrophotometer with 1880 μL of DI water and 100 μL of 5 mM ABTS solution, 20 μL of enzyme solution was added to the spectrometer cell. The solution was mixed by inversion, and the absorbance at 414 nm (ε414 = 36 000 M–1 cm–1)[22] was recorded immediately every 7 s for 2 min or until a plateau was observed. Enzyme activities were expressed in microkatals (μkat), the amount of enzyme that catalyzes the conversion of 1 μmol of substrate/s at room temperature. For activity studies conducted at temperatures other than room temperature, native laccase or complexed laccase solutions were incubated in a water bath for 20 min and were then immediately analyzed. Laccases with particularly high activities were diluted by a factor of 100 prior to the assay. All activity tests were completed in triplicates.

Polymerization of Tyrosine

Polymerization of tyrosine was conducted as previously described.[15] Briefly, in a typical polymerization, 60 mg of dl-tyrosine is added to a vigorously stirred laccase (activity 77 μkat/L) or complexed laccase solution. Polymerization of tyrosine was conducted at 45 °C for 72 h. The unreacted monomer was removed via centrifugation. The supernatant pH was then adjusted to 2.6–2.7 with HCl and NaOH to precipitate poly(tyrosine). Further purification included dissolving the obtained poly(tyrosine) in neutral DI water and precipitating into THF to obtain a dark brown solid product. All polymerizations were completed in triplicates.

Decolorization of Synthetic Dyes

Decolorization of the synthetic dyes bromophenol blue, methyl orange, and rhodamine B base was conducted by first preparing a native laccase or laccase complex solution with an enzyme concentration of 4 mg/mL in an acetate buffer of pH 4.5. This solution was incubated at 45 °C for 4 h after which 0.77 mg of HOBT dissolved in 50 μL of THF was added. The resulting solution was allowed to incubate for an additional 2 h. To this solution, 1 mL of a stock dye solution was added for a final dye concentration of 0.096 mg/mL. The absorbance of the solution was recorded at predetermined time intervals at the wavelengths of 592, 465, and 544 nm for bromophenol blue, methyl orange, and rhodamine B base, respectively.

For recyclability studies, experiments were set up as described above for the initial cycle (cycle 1). After complete decolorization, the solution was centrifuged for the removal of the supernatant. Fresh dye and buffer were then added for a second decolorization (cycle 2). After decolorization, this solution was also centrifuged to remove the supernatant. Fresh laccase in an amount equivalent to the first cycle, dye, and HOBT, unless otherwise specified, were then added for the third cycle. This procedure was repeated over eight cycles.

Results and Discussion

Copolymer Synthesis and Characterization

The linear–linear and linear–hyperbranched copolymers were synthesized by ATRP using PEG functionalized with α-chlorophenylacetyl chloride and either styrene or PCMS.[16−18] The formation of the copolymers was verified via 1H NMR and size-exclusion chromatography, SEC (see description and Figures S1–S4 in the Supporting Information). MMs of the produced polymers (Table S1) were determined by comparing the aromatic signals from the 1H NMR spectra to the ethylene signals of PEG. The SEC traces of the produced linear(A)–linear(B) and linear(B)–linear(A)–linear(B) copolymers showed monomodal distributions, indicating an efficient initiation of the macroinitiator and confirming that a homogeneous product was obtained (Figure S3 in the Supporting Information).

Copolymer Self-Assembly in Aqueous Media

Thirteen of the amphiphilic copolymers self-assemble into micelles (Table ). With two exceptions (PS750–PEG3k–PS750, 51% and PPCMS1.5k–PEG3k–PPCMS1.5k, 72%), their hydrophobic content is below 51%. All of these complexing agents were found to form micelles with a peak hydrodynamic diameter between 21 and 400 nm (Figures –4). Overall, the MMs of the copolymer building blocks affected the hydrodynamic diameters of the micelles formed. With the same PEG chain length, macromolecules with larger hydrophobic domains formed micelles with significantly higher hydrodynamic diameters (Figure ).

Table 1

Enzyme Activity at 45 °C and Polymerization Yields Depending on the Catalyst Type

copolymer (form of self-assembly)	activity ABTS (μkat/L)a	poly(tyrosine) yield (%)b	hydrophobic blocks (%)
native	1999	58.8	N/A
PS2.7k–PEG3k–PS2.7k (H)a	8161	66.9	64
PS10k–PEG13k–PS10k (H)	2027	51.1	61
PS12.5k–PEG20k–PS12.5k (H)	2598	71.4	56
PS4.2k–PEG20k–PS4.2k (H)	3518	79.0	30
PPCMS6.5k–PEG3k–PPCMS6.5k (H)	2026	67.0	81
PPCMS9k–PEG13k–PPCMS9k (H)	2513	52.1	74
PPCMS13k–PEG13k–PPCMS13k (H)	2505	78.4	67
PPCMS24k–PEG20k–PPCMS24k (H)	3284	57.6	58
G3–PEG5k–G3 (H)	433	85.5	39
G3–PEO13k–G4 (H)	4222	72.3	26
PS750–PEG3k–PS750 (M)b	3671	62.1	51
PEG5k–PS5.2k (M)	2292	75.5	40
PEG5k–PS1k (M)	4664	69.0	33
PS4.2k–PEG13k–PS4.2k (M)	3384	56.9	17
PS750–PEG13k–PS750 (M)	1972	84.3	10
PS1k–PEG20k–PS1k (M)	2878	72.3	9
PPCMS1.5k–PEG3k–PPCMS1.5k (M)	4441	74.6	72
PEG5k–PPCMS13k (M)	3038	69.8	50
PEG5k–PPCMS1.7k (M)	3250	73.7	26
PPCMS2.3k–PEG13k–PPCMS2.3k (M)	4024	85.0	25
PPCMS2.5k–PEG20k–PPCMS2.5k (M)	2345	60.3	20
G3–PEO13k–G2 (M)	5067	65.7	15
G3–PEO13k (M)	2554	70.5	11

H—hydrogel.

M—micelle.

Figure 2

Hydrodynamic diameters, Dh, of PEG3kDa ABA copolymers in aqueous solution at 25 °C (1 mg/mL): PS750–PEG3k–PS750 (--- black) and PPCMS1.5k–PEG3k–PPCMS1.5k (— red).

Figure 4

Hydrodynamic diameters, Dh, of PEG20kDa ABA copolymers at 25 °C (water, 1 mg/mL): PPCMS2.5k–PEG20k–PPCMS2.5k (--- black) and PS1k–PEG20k–PS1k (— red).

Similarly, it was observed that copolymers having hydrophobic blocks of similar MM but longer PEG chains formed larger micelles compared to their shorter analogues. This trend is evident when comparing PS–PEG–PS complexing agents PS750–PEG3k–PS750 (hydrodynamic diameter of 18.12 nm), PS750–PEG13k–PS750 (hydrodynamic diameter of 327.4 nm), and PS1k–PEG20k–PS1k (hydrodynamic diameter of 351.8 nm) (Figure ).

Figure 3

Hydrodynamic diameters, Dh, of PS750–PEG3k–PS750 (— red), PS750–PEG13k–PS750 (--- black), and PS1k–PEG20k–PS1k (··· blue) at 25 °C (water, 1 mg/mL).

Macromolecular architecture also seems to play a role in the dimensions of the micelles formed. With the hyperbranched blocks being more compact, their supramolecular assemblies tend to have smaller hydrodynamic diameters compared with their linear analogues (Figure ).

By comparing the hydrodynamic diameters of native laccase, an “empty” copolymer micelle and the enzyme/copolymer complex formed by solvent exchange (Figure ) indicate a noticeable increase in the hydrodynamic diameter. This increase in diameter between 20 and 50 nm was observed often and is evidence of the incorporation of enzyme into the micelles formed by the synthesized copolymers (see also Figures S8–S10 in the Supporting Information).

Figure 5

Hydrodynamic diameters, Dh, of native laccase (··· blue), empty PS750–PEG13k–PS750 micelle (— red), and PS750–PEG13k–PS750 laccase complex (--- black) at 25 °C (water, 1 mg/mL).

Amphiphilic copolymers of the hydrophilic/hydrophobic weight ratio above 50% were capable of forming physical networks in aqueous solution with two notable exceptions (linear–dendritic copolymers with high-generation hydrophobic blocks: G3–PEG5k–G3 and G3–poly(ethylene oxide) PEO13k–G4) (Table ). SEM analyses showed that the hydrogels have a well-developed pore-in-pore structure, which did not change upon binding of the enzyme. An example is shown in Figure .

Figure 6

SEM of physical networks formed in water from the amphiphilic linear–dendritic copolymer G3–PEG5k–G3: (A) self-assembled with no laccase added and (B) self-assembled in the presence of laccase.

Activity of Complexed Laccase toward ABTS

With the exception of PS750–PEG13k–PS750, all micellar complexes experienced a significant increase in activity (Table ) at 45 °C. This increase can be attributed to the favorable interactions between the glycosidic regions of the enzyme and the hydrophobic portions of the copolymers which facilitate the transportation of the substrate to the enzyme active site. When comparing the linear–dendritic and dendritic–linear–dendritic copolymers of equivalent PEG chain lengths (PEG13kDa), it can be seen that increasing the dendron/PEG weight ratio results in a beneficial effect on the enzyme activity (Table ). It was found that for dendritic complexing agents, the size of the hydrophilic PEG chain affects the form of self-assembly (micelle vs hydrogel) and heavily influences the activity (Table , Figure ), with G3–PEG5k–G3 (hydrogel) experiencing one of the lowest activities. This is presumably due to the high density of the network (see Figure ) with relatively short interlink distances and a high hydrophobic content, resulting in a low swelling and limited mobility of the enzyme because of the shortened PEG chain length. These factors do not facilitate the partitioning of the water-soluble ABTS substrate, its diffusion into the hydrogel, and its interaction with the enzyme active site.

Figure 7

Temperature dependence on the activities of native laccase (black circle), G3–PEG5k–G3/laccase (red square), G3–PEO13k/laccase (green triangle), G3–PEO13k–G2/laccase (yellow circle), and G4–PEO13k–G3/laccase (blue diamond).

With lower molecular weight PEG-based complexing agents (PEG3kDa and PEG monomethyl ether 5 kDa), longer linear hydrophobic regions were found to induce higher activities (Figures and 9). The contrary was observed with PPCMS-containing copolymers, where the smaller hydrophobic regions led to a higher activity (Figure ). It should be noted that the PS2.7k–PEG3k–PS2.7k/laccase complex had a remarkably high activity, approximately 3.5× that of the native laccase at 60 °C and 10× that of the native laccase at 20 °C (Figure ).

Figure 8

Temperature dependence on the activities of native laccase (black circle) and PEG3kDa copolymers PPCMS1.5k–PEG3k–PPCMS1.5k/laccase (red square), PS750–PEG3k–PS750/laccase (green triangle), PS2.7k–PEG3k–PS2.7k/laccase (yellow circle), and PPCMS6.5k–PEG3k–PPCMS6.5k/laccase (blue diamond).

Figure 9

Temperature dependence on the activities of native laccase (black circle) and PEG monomethyl ether 5kDa copolymers PEG5k–PPCMS1.7k/laccase (red square), PEG5k–PPCMS13k/laccase (green triangle), PEG5k–PS1k/laccase (yellow circle), and PEG5k–PS5.2k/laccase (blue diamond).

For both PEG13kDa and PEG20kDa-based copolymers, it was observed that linear(B)–linear(A)–linear(B) and hyperbranched–linear–hyperbranched copolymers are both capable of increasing the enzyme activity when the hydrophobic regions are of sufficient size (Figures and 11). Examples of this trend include PPCMS2.3k–PEG13k–PPCMS2.3k and PPCMS24k–PEG20k–PPCMS24k of hyperbranched–linear–hyperbranched morphology and PS4.2k–PEG20k–PS4.2k and PS4.2k–PEG13k–PS4.2k of linear(B)–linear(A)–linear(B) morphology which all contain hydrophobic/hydrophilic weight ratios of at least 0.35 and show increased activities when compared with the native enzyme. This is in contrast to the linear(B)–linear(A)–linear(B) complexing agents PS750–PEG13k–PS750 and PS1k–PEG20k–PS1k and the hyperbranched–linear–hyperbranched complexing agent PPCMS2.5k–PEG20k–PPCMS2.5k which show activities similar to that of the native laccase and have a noticeably lower hydrophobic/hydrophilic weight ratio, most likely resulting in weak interactions between the complexing agent and the enzyme.

Figure 10

Temperature dependence on the activities of native laccase (black circle) and PEG13kDa copolymers PS4.2k–PEG13k–PS4.2k/laccase (red square), PPCMS13k–PEG13k–PPCMS13k/laccase (green triangle), PS10k–PEG13k–PS10k/laccase (yellow circle), PPCMS9k–PEG13k–PPCMS9k/laccase (blue diamond), PPCMS2.3k–PEG13k–PPCMS2.3k/laccase (purple square), and PS750–PEG13k–PS750/laccase (gray triangle).

Figure 11

Temperature dependence on the activities of native laccase (black circle) and PEG20kDa copolymers PS12.5k–PEG20k–PS12.5k/laccase (red square), PPCMS2.5k–PEG20k–PPCMS2.5k/laccase (green triangle), PS1k–PEG20k–PS1k/laccase (yellow circle), PS4.2k–PEG20k–PS4.2k/laccase (blue diamond), and PPCMS24k–PEG20k–PPCMS24k/laccase (purple square).

Both native and complexed laccase experienced an increase in the enzymatic activity with increasing temperature to a maximum between 60 and 65 °C (Figures –11). At 70 °C, all laccase solutions showed diminished activities, with many showing none at all. This significant decrease in activity is most likely due to the denaturation of the enzyme at elevated temperature in addition to the reduction in PEG solubility. As the hydrophilic part of all complexing agents is composed of PEG, which experiences a reverse solubility effect between 63 and 65 °C, a decrease in activity is not unexpected because of the PEG collapse and the resulting restriction of enzyme mobility and limited interaction with the substrates. When comparing the linear–dendritic and dendritic–linear–dendritic copolymers of PEG chain lengths of 13 kDa, it was observed that increasing the dendron/PEG weight ratio resulted in beneficial effects on the enzyme stability at elevated temperatures (Figure , G4–PEO13k–G3/laccase). This increased enzyme stability is most likely due to more efficient dendron docking onto the glycoside domains and a better shielding of the enzyme from the surrounding environment.

PS2.7k–PEG3k–PS2.7k could be singled out as the most efficient complexing agent with activities increasing from 8000 μkat/L to over 13 000 μkat/L in a temperature range of 45–65 °C (Figure ). This is in contrast to PPCMS6.5k–PEG3k–PPCMS6.5k/laccase (a hyperbranched–linear–hyperbranched hydrogel with the same PEG3kDa), which shows a little difference in activity from the native enzyme. The same was seen with PS750–PEG3k–PS750/laccase and PPCMS1.5k–PEG3k–PPCMS1.5k/laccase, which both show moderate increases in activity. It was observed that laccase bound to copolymers of PEG13kDa and PEG20kDa with large linear hydrophobic blocks experienced limited benefits in regard to activity and stability. This might be due to the inefficient adhesion of the linear hydrophobic fragments onto the glycoside regions (with an increased screening of the active site) and the loose enzyme association within the large network pores as well.

Application of Immobilized Laccase for Tyrosine Polymerizations

To determine the application of complexed laccase for the transformation of substrates with low water solubility, tyrosine polymerizations were conducted using a previously described method.[15]

It was found that G3–PEG5k–G3 had the highest yield (85.5%), whereas PS10k–PEG13k–PS10k had the lowest yield (51.1%) (Table ). Overall, it was observed that a high activity (as determined with ABTS) does not necessarily result in a high poly(tyrosine) yield. This is most likely due to the differences between ABTS and tyrosine in their partitioning coefficients and diffusion through the complexing agents. Additionally, weakly complexed enzymes, which still experience the beneficial activity effects of copolymer association, would have larger pore sizes than those of more tightly complexed enzymes (G3–PEG5k–G3/laccase) which bind tyrosine closer in proximity to the enzyme active site. With few exceptions, it was found that linear-based complexes had a noticeably reduced yield when compared with either linear–dendritic, dendritic–linear–dendritic, linear–hyperbranched, or hyperbranched–linear–hyperbranched complexing agents. This is most likely the result of linear-based complexes forming micelles or hydrogels with the hydrophobic regions of increased density. This increased density leads to a lower tyrosine binding capacity and likely hampers the permeation of tyrosine to the enzyme active site. It was found that micelles in general had a higher yield (70.75%) when compared with the hydrogels (68.13%), most probably a result of greater substrate diffusion efficacy in micelles.

In regard to poly(tyrosine) microstructure (C–C vs C–O coupling), the presence of any complexing agent had no apparent effect, with all polymers synthesized containing a C–O coupling of approximately 76.4%.

Decolorization of Synthetic Dyes

To determine the applicability of the synthesized immobilizing agents for bioremediation purposes, the decolorization of bromophenol blue, methyl orange, and rhodamine B base (all of them being common synthetic dyes) was investigated. The hydrogel complexes showing the most suitable activity/easiness of production ratio were chosen for the trials (PS2.7k–PEG3k–PS2.7k and PPCMS24k–PEG20k–PPCMS24k). Compared to previously reported literature values[23] for bromophenol blue, the observed reaction rate was markedly higher. HOBT, a radical mediator, was required for the treatment of methyl orange and rhodamine; it also significantly enhanced the decolorization rate of bromophenol blue. The hydrogel complexes degraded the dyes at a slower rate than the native laccase (Figures and 13), presumably because of the slower substrate diffusion through the hydrogel to the enzyme active site. On the other side, it should be noted that methyl orange is repeatedly (!) transformed at the same rate as with the pure enzyme. This clearly indicates that partition coefficient of the substrate (dye in this case) plays a major role in defining (enhancing or slowing) the rate of diffusion and ultimately the rate of the reaction. However, the physical entrapment of the biocatalyst within these networks did allow for the reusability of both the laccase and immobilizing agent. The enzymatic hydrogels were recycled by centrifuging and the removal of the supernatant. After the introduction of a fresh dye and buffer, a reduced speed of decolorization was observed (second cycle and every even cycle). Several factors might have contributed for this lower rate. Model experiments, performed at the same conditions (copolymer, centrifugation speed, temperature, and enzyme concentration), showed that during centrifugation, a part of the enzyme was “squeezed” out of the hydrogels because of their intrinsic softness (Figure S12 in the Supporting Information). During centrifugation, gravitational forces might have also caused the collapse of the pores, thus reducing the diffusivity of the soft networks (see Figure ). Addition of fresh laccase and HOBT to the swollen hydrogels was found to restore the activity to levels close to those of the original solution (third cycle and every odd cycle thereafter). In that way, an economy of 50% enzyme and mediator could be achieved while using the same initial carrier hydrogel (100% network recyclability).

Figure 12

Recyclability of immobilized laccase in PS2.7k–PEG3k–PS2.7k hydrogel for decoloring bromophenol blue with HOBT additive (A), bromophenol blue without HOBT additive (B), methyl orange with HOBT additive (C), and rhodamine B base with HOBT additive (D). Fresh HOBT and/or laccase was added every odd cycle (full symbols).

Figure 13

Recyclability of immobilized laccase in PPCMS24k–PEG20k–PPCMS24k hydrogel for decoloring bromophenol blue with HOBT additive (A), bromophenol blue without HOBT additive (B), methyl orange with HOBT additive (C), and rhodamine B base with HOBT additive (D). Fresh HOBT and/or laccase was added every odd cycle (full symbols).

The proposed laccase/HOBT-catalyzed degradation mechanism of bromophenol blue (Scheme ) centers on the disruption of the quinone–phenol resonance structure of bromophenol blue. Under the reaction conditions used (pH 4.5 buffer), this structure is favored, allowing for a hydroxyl radical attack. Further degradation occurs with the radical-facilitated removal of bromine from the molecule, resulting in a colorless degradation product. Matrix-assisted laser desorption ionization time-of-flight mass spectrometry (MALDI-TOF MS) of the degradation mixture confirms the loss of four bromine atoms and the addition of a hydroxyl group (see product with 365 m/z in Figure S13B). The NMR monitoring of bromophenol blue degradation shows the appearance of four additional equivalent aromatic hydrogens at 7.96 ppm (H6, Figure ), confirming the loss and replacement of bromine with hydrogen. Further evidence for the proposed mechanism is provided by the coincidence of NMR data (integral intensity ratio H6/H1, Figure ) with the colorimetric assay results (Figure ).

Scheme 1

Proposed Laccase/HOBT-Catalyzed Degradation of Bromophenol Blue

Figure 14

Time evolution of NMR spectra showing the progression of bromophenol blue degradation.

Figure 15

Bromophenol blue degradation by native laccase/HOBT monitored by NMR (blue circle) and colorimetric assay (orange diamond). Ar-OH H/ArSO3 H is the ratio of the integral intensity for aromatic hydrogen at 7.96 ppm (Ar-OH H, H6 in Figure ) to constant aromatic hydrogen at 8.06 ppm (ArSO3 H, H1 in Figure ).

Conclusions and Outlook

Amphiphilic linear(A)–linear(B), linear(B)–linear(A)–linear(B), linear–hyperbranched, and hyperbranched–linear–hyperbranched copolymers composed of PEG of various chain lengths and either PS or PPCMS were synthesized via ATRP for use as enzyme complexing agents. DLS studies confirmed that micellar complexing agents were capable of binding to the enzyme, resulting in the desired complex. PS2.7k–PEG3k–PS2.7k, a linear(B)–linear(A)–linear(B) PEG3kDa-based copolymer, provided the greatest benefit to the enzyme activity and stability at elevated temperatures (up to +1245% vs the native enzyme), while producing only a modest increase in the tyrosine polymerization yield (+13.7% vs the native enzyme). Although G3–PEG5k–G3 was found to substantially reduce the enzyme activity (433 vs 1999 μkat/L), it provided the largest tyrosine polymerization yield at 85.5% because of its strong binding of the substrate. Thus, the macromolecular architecture of the complexing agents has a strong influence on polymerization yields. Overall, the laccase complexes of linear–dendritic copolymers produced poly(tyrosine) with the highest yield followed by the linear–hyperbranched analogues, probably because of the nanoporosity of their hydrophobic blocks, which facilitates efficient sequestration and binding of the hydrophobic substrate and its migration to the active site. In contrast, the dense packing of the PS blocks during the self-assembly of the linear–linear copolymers prevents the efficient binding and transport of tyrosine to the active site of laccase.

An increased enzymatic activity resulting from enzyme complexing reported in this work is an encouraging improvement over the traditional immobilization methods, which often result in significant activity decreases either from the decreased enzyme mobility, restriction of the substrate access to the enzyme active site, or limited diffusion of the substrate to the enzyme.[24] Current examples which exemplify these shortcomings include the immobilization of laccase via entrapment in alginate–gelatin gels, which resulted in an activity decrease of 15%,[25] and the covalent immobilization of laccase to titania nanoparticles, which incurred an activity decrease between 21 and 85%.[26] In an earlier report by Mateescu et al., the final activity of laccase, immobilized on p-benzoquinone-activated agarose, was only 27% compared to the free laccase because of an insufficient laccase binding of only 18%.[27] Thus, the laccase complexation method reported in this work offers distinct advantages to many previously reported immobilization methods regarding retained activity.

Successful decolorizations of the synthetic dyes bromophenol blue, methyl orange, and rhodamine B base display the versatility of immobilized laccase to decolor a wide array of dyes (phenolic, diazo, and heterocyclic). The broad substrate specificity and reusability of the immobilized laccase make the described complexing method very desirable for bioremediation and other catalytic roles.