One-Step Synthesis of Cellulose from Cellobiose via Protic Acid

Aug 7, 2012 - ABSTRACT: Direct and efficient enzymatic synthesis of long-chain cellulose from cellobiose in its original form was successfully achieve...
0 downloads 0 Views 2MB Size
Download PDF
Article pubs.acs.org/Biomac

One-Step Synthesis of Cellulose from Cellobiose via Protic AcidAssisted Enzymatic Dehydration in Aprotic Organic Media Shizuka Egusa,† Masahiro Goto,‡ and Takuya Kitaoka*,† †

Department of Agro-Environmental Sciences, Graduate School of Bioresource and Bioenvironmental Sciences, and Biotron Application Center, Kyushu University, 6-10-1 Hakozaki, Higashi-ku, Fukuoka, 812-8581 Japan ‡ Department of Applied Chemistry, Graduate School of Engineering, Kyushu University, 744 Motooka, Nishi-ku, Fukuoka, 819-0395 Japan S Supporting Information *

ABSTRACT: Direct and efficient enzymatic synthesis of long-chain cellulose from cellobiose in its original form was successfully achieved via the combination of a surfactant-enveloped enzyme (SEE) and a protic acid in an aprotic organic solvent, lithium chloride/N,N-dimethylacetamide system. The SEE biocatalyst was prepared by protecting the surface of cellulase with the nonionic surfactant dioleyl-N-D-glucona-L-glutamate for keeping its enzymatic activity in nonaqueous media. Fourier transform infrared and nuclear magnetic resonance analyses elucidated the successful synthesis of cellulose, β-1,4-linked D-glucopyranose polymer, through the reverse hydrolysis of cellobiose. By using protic acid cocatalysts, a degree of polymerization of assynthesized cellulose reached more than 120, in a ca. 26% conversion, which was 5 times higher than that obtained in an acid-free SEE system. A novel-concept biocatalysis, i.e., a protic acid-assisted SEE-mediated reaction, enables a facile, one-step chain elongation of carbohydrates without any activation via multistep organic chemistry, and can provide potential applications in the functional design of glycomaterials.

■

INTRODUCTION Facile and efficient synthesis of carbohydrate chains has recently attracted considerable attention from biological, medical, and biomaterial-engineering standpoints since a variety of carbohydrates play essential roles in all living systems.1 Many researchers have devoted much effort to synthesize and design glyco-materials. Conventional chemical glycosynthesis methods generally require complicated procedures,2 thus, enzyme has become a potential option for efficient glycomaterial synthesis due to its substrate-specific ability.3 However, in most cases, the chemical activation of starting sugar via organic synthesis still remains indispensable for enzymatic synthesis, known as chemo−enzymatic synthesis.4−6 Direct, nonprotective synthesis methods of carbohydrate chains and their conjugates have long been eagerly desired in glycomaterial-engineering fields. In our previous studies, we have proposed a new-conceptual enzymatic glycosynthesis, using a surfactant-enveloped enzyme (SEE). The direct in vitro synthesis of cellulose from cellobiose, with no anomeric carbon activation, was first achieved in © 2012 American Chemical Society

dimethylacetamide (DMAc) containing lithium chloride (LiCl).7 The LiCl/DMAc solvent can dissolve various carbohydrates, but deactivate enzyme. However, the SEE biocatalysts can preserve their initial enzymatic activity to a certain degree in nonaqueous solvents8 that are too severe for enzymes.4 This novel strategy has allowed us to achieve the synthesis of a long-chain cellulose with a degree of polymerization (DP) greater than 100,7 the one-step glycosylation of hydrophobic alcohols,9 and the direct surface glyco-modification of cellulose matrix.10 However, the enzymatic dehydration proceeded with a relatively low efficiency, even in nonaqueous media usually suitable for dehydration; thus the average conversion from cellobiose to water-insoluble long-chain cellulose remained insufficient, e.g., 2−5%.7,8 Received: April 29, 2012 Revised: July 24, 2012 Published: August 7, 2012 2716

dx.doi.org/10.1021/bm3006775 | Biomacromolecules 2012, 13, 2716−2722

Biomacromolecules

Article

Scheme 1. The Protic Acid-Assisted Enzymatic Synthesis of Cellulose from Cellobiose via SEE-Mediated Biocatalysis in Organic Mediaa

a

Route 1: the proposed reverse hydrolysis to promote the chain-elongation reaction of cellulose in nonaqueous/quasi-nonaqueous media with a protic acid cocatalyst; l = 1 and m = 2 in the initial stages. Route 2: the general acid hydrolysis of cellulose in an aqueous system.

■

The key to solving this problem is proposed to be a protonmediated event in the enzymatic dehydration reaction. A proton (H+) is also the simplest and most versatile catalyst in organic synthesis.11 Protic acid catalysts, such as hydrochloric and sulfuric acids, can efficiently hydrolyze polysaccharides in aqueous media,12 and the carboxylates present in glycohydrolases can catalyze the same reaction.13 This is similar to the way protons coordinate to a glycosidic oxygen to initiate a hydrolytic reaction. Recently, Jacobsen et al. have reported a combined sulfinamido urea/o-nitrobenzenesulfonic acid (oNBSA) system for highly efficient catalysis.14 A number of synthetic reactions proceed via the noncovalent interaction of a primary catalyst with a reactive intermediate produced in situ by an acid cocatalyst. Additionally, a variety of cascade reaction systems via chemo−bio interactions have rapidly advanced as one of the important future directions in facile and efficient catalysis.15 These strategies provide us a hint to improve our nonaqueous biocatalysis for the direct synthesis of cellulose from cellobiose via the reverse reaction of acid-catalyzed polysaccharide hydrolysis. Herein, we describe a protic acid-assisted SEE-mediated polymerization of cellobiose to synthesize cellulose in an aprotic LiCl/DMAc solvent system, as shown in Scheme 1. The general hydrolysis of glycosidic linkage is triggered by the coordination of a proton, derived from an acid catalyst, to the oxygen atom of the linkage (route 2 in Scheme 1). In contrast, our strategy is to promote the “reverse” of this reaction in an aprotic organic medium, through the addition of a protic acid cocatalyst. This may in situ produce a cellobiose intermediate with an activated C1 carbon at a reducing end (route 1 in Scheme 1). Hence, SEE biocatalysts that are active even in aprotic media7−10 could more efficiently mediate the catalytic dehydration of the activated cellobiose to form a new glycosidic linkage in a one-pot system. As-synthesized products were characterized by Fourier transform infrared (FTIR) and nuclear magnetic resonance (NMR) analyses. The effects of synthetic conditions on conversion ratios, DP values and catalytic efficiency were investigated. Possible reaction mechanism and kinetics are also discussed.

EXPERIMENTAL SECTION

Materials. Crude cellulase, consisting of 1,4-β-D-glucan 4glucanohydrolase (EC 3.2.1.4) from Trichoderma viride, was purchased from Wako Pure Chemical Industries Co., Ltd. The nonionic surfactant dioleyl-N-D-glucona-L-glutamate (2C18Δ9GE) was synthesized according to Goto’s method.16 Lithium chloride (LiCl, Wako Pure Chemical Industries Co., Ltd.), nitrobenzenesulfonic acids (oNBSA, mNBSA, pNBSA, Tokyo Chemical Industry Co. Ltd.), ptoluenesulfonic acid (pTSA, Sigma-Aldrich Co., LLC), and trifluoromethanesulfonic acid (TFMSA, Tokyo Chemical Industry Co., Ltd.) were dried under reduced pressure at 50 °C for 12 h before use. Cellobiose (Sigma-Aldrich Co., LLC) was also dried under reduced pressure at 50 °C for 12 h before use. DMAc (Wako Pure Chemical Industries Co., Ltd.) was desiccated for at least 3 days in the presence of 4 Å molecular sieves (Sigma-Aldrich Co., LLC), which were predried at 250 °C for 30 h. The water used in this study was purified using a Milli-Q system (Millipore Co. Ltd.). Other chemicals were of reagent grade and used without further purification. Preparation of SEE. Cellulase (30 mg) in sodium carbonate buffer (1.5 mL, 50 mM, pH 9.1) and 2C18Δ9GE (15 mg) in toluene (3 mL) were mixed and vigorously homogenized at 15 000 rpm for 90 s. The cloudy water-in-oil emulsion was immediately frozen with liquid nitrogen, and then freeze-dried for at least 24 h. The SEE biocatalyst was obtained in a powdery form.7−10 The cellulase solution was stirred at ca. 100 °C for 10 min to deactivate the enzyme, and the inactive SEE as a control was prepared from the deactivated cellulase in a similar manner. Cellulose Synthesis. The powder-state SEE was poured into 0.5 mL DMAc-containing cellobiose with 1.0 wt % LiCl and an acid catalyst, such as concentrated sulfuric acid (minimum concentration: 97%), oNBSA, mNBSA, pNBSA, hydrochloric acid (HCl), pTSA, or TFMSA. As controls, the reaction mixtures containing either the SEE or the acid catalyst were also prepared. The condensation reaction proceeded at 40 °C for 2−24 h, and the conversion ratios were obtained by the following equation:

Conversion(%) = [consumed cellobiose]/[initial cellobiose] × 100

(1)

where the [consumed cellobiose] is obtained by subtracting the detected [residual cellobiose] from the supplied [initial cellobiose]. The products were precipitated with methanol, thoroughly washed with methanol and chloroform to remove the LiCl, DMAc, and 2C18Δ9GE, and freeze-dried. The water-soluble oligomer fractions, which contained unreacted cellobiose, were separated from the 2717

dx.doi.org/10.1021/bm3006775 | Biomacromolecules 2012, 13, 2716−2722

Biomacromolecules

Article

Figure 1. Optical images of the reaction media of the SEE-mediated biocatalysis after 24 h incubation with (a) and without (b) sulfuric acid. (c) SEE-free system with sulfuric acid. (d) The final solution treated with the SEE and sulfuric acid exhibited a significant viscosity. containing 20 μL of a sodium acetate buffer solution (1.0 M, pH 4.8). Then, 20 μL of a 10 mM p-nitrophenyl lactoside solution was added to start the hydrolytic reaction. After an incubation period of 10 min at 30 °C, the reaction was quenched by adding 20 μL of a sodium carbonate solution (2.0 M). The levels of p-nitrophenol liberated were detected at 405 nm, using an iMark microplate absorbance reader (Bio-Rad Laboratories Inc.), and compared to crude cellulase and Cel7A. The enzymatic activity assay was repeated at least three times in independent experiments. The averaged absorbance at 405 nm was 0.487 and 0.348 for Cel7A and crude cellulase, respectively. As a result, the enzymatic activity of the 0.6% crude cellulase solution corresponded to that of a ca. 7.1 μM Cel7A solution. In this work, the TOF values were approximately calculated by using the estimated molar content of cellulase equivalent to that of pure Cel7A. Calculation of TOF Value. The TOF values were calculated from the molar decreases in concentration for the residual (unreacted) cellobiose. Free glucose as a hydrolyte was not detected in all cases. Thus, the TOF values are theoretically lower than the real ones. This is because SEE biocatalysts can catalyze reactions between the oligomers to form long-chain cellulose, while a constant cellobiose concentration is maintained. The TOF values were obtained according to the following equation:

synthesized cellulose via dissolution in water. The water-insoluble portions were then accurately measured by weight to obtain the yields after freeze-drying, and subjected to analytical characterization. Acid Hydrolysis of Synthetic Cellulose. Acid hydrolysis of water-insoluble synthetic products was performed to quantitatively measure the component sugars according to the previous report.17 The samples were dispersed in 1 M HCl and incubated at 105 °C for 6 h. The clear solutions obtained were neutralized by adding 2 M sodium carbonate solution, and the supernatants were analyzed by high-performance liquid chromatography (HPLC). Acetylation of Synthetic Cellulose. Acetylation of synthetic cellulose was carried out for structural characterization according to the previous report.18 Pyridine (0.1 mL) was added to a solution (1 mL) of synthetic cellulose (5 mg), LiCl (80 mg), and DMAc (1 mL). Acetic anhydride (0.14 mL) was added dropwise to the mixture, followed by stirring at 60 °C for 13 h. Analytical Characterization. HPLC was carried out to quantitatively determine the amounts of residual cellobiose at a given time, free glucose in reaction media, and glucose obtained by acid hydrolysis of products. A TOSOH HPLC instrument equipped with NH2P column (Shodex Co., Ltd.) was used; mobile phase, 72% acetonitrile/ultrapure water; flow rate, 1.0 mL min−1; detector, refractive index. FTIR spectra of samples mixed with potassium bromide powder were recorded using a JASCO FTIR-620 spectrometer under transmission mode from 800 to 4000 cm−1 with a 2 cm−1 resolution. 1 H NMR analysis was performed on a JEOL JMN-AL400 FT-NMR spectrometer (400 MHz). CDCl3 was used as a solvent. Viscometric analysis was carried out to measure the viscosityaverage degree of polymerization (DPv) of the water-insoluble products without any derivatization.7 The samples were completely dissolved in 0.5 M cupriethylenediamine solution. The intrinsic viscosities [η] (mL g−1) of the cellulose solution were measured by using an Ostwald-type viscometer in a water bath set at 27 °C, and the DPv values were calculated according to the following modified Mark−Houwink−Sakurada equation:19

[η](mL g −1) = KM a = 0.571 × DPv1.0

TOF (h−1) = consumed cellobiose (μmol) /cellulase in SEE (μmol)/reaction time (h)

(3)

where the consumed cellobiose (μmol) is obtained by subtracting the detected residual cellobiose (μmol) from the supplied initial cellobiose (μmol), and the cellulase in SEE is the estimated molar quantity of cellulase (μmol) obtained by cellulase activity assay.

■

RESULTS AND DISCUSSION Cellulose Synthesis via Protic Acid-Assisted SEEMediated Biocatalysis. The enzymatic synthesis of cellulose from cellobiose, in its original form, was carried out using the combination of an SEE biocatalyst and concentrated sulfuric acid (30 mM) in the LiCl/DMAc system. Then, the reaction media shifted from being nonaqueous to quasi-nonaqueous (microaqueous, water content: less than 1 mM). After 4 h incubation at 40 °C, the reaction solution was significantly viscous and became a gel-like state (Figure 1a). Conversely, in the absence of either the SEE (Figure 1b) or sulfuric acid (Figure 1c), there was nearly no viscoelastic change observed. After 24 h incubation (Figure 1d), the products were precipitated for FTIR spectroscopy and DPv measurement. Subsequently, the precipitates were acetylated for 1H NMR analysis and DPw determination by GPC. As-synthesized samples showed the characteristic FTIR spectrum for cellulose (Figure 2). A regenerated cellulose sample as a control was prepared by mercerization using 20% sodium hydroxide aqueous solution according to the conventional protocol.21 The broad absorption bands at 3100−3700 cm−1 correspond to the stretching vibration of OH groups. The bands at 2800− 3000 cm−1 are attributed to the vibration of CH2 groups.22 The

(2)

Gel-permeation chromatography (GPC) analysis was performed to determine the weight-average DP (DPw) of water-insoluble synthetic cellulose after acetylation.18 A TOSOH HPLC instrument equipped with a TSKgel α-M column (TOSOH Co., Ltd.) was used; mobile phase, tetrahydrofuran; flow rate, 1.0 mL min−1; detector, refractive index. Cellulase Activity Assay. Crude cellulase from Trichoderma viride contains various types of cellulases such as cellobiohydrolase (CBH; Cel7A, a major component) and endoglucanase (EG; Cel7B, minor one); the actual component ratios depend on fungal origins and culture conditions. Thus, the molar content of cellulase in the asdesigned SEE biocatalyst was roughly determined by comparison with the enzymatic activity for pure Cel7A to approximately calculate the turnover frequency (TOF) values. The Cel7A stock solution concentration was determined from the absorbance at 280 nm, using a U-3000 spectrophotometer (HITACHI Co., Ltd.). A 10 μM Cel7A solution was prepared (molar absorbance coefficient: 84650 M−1).20 Either 20 μL of a 0.6% crude cellulase solution or 20 μL of a 10 μM Cel7A solution was poured into 140 μL of deionized water 2718

dx.doi.org/10.1021/bm3006775 | Biomacromolecules 2012, 13, 2716−2722

Biomacromolecules

Article

acid catalysts on the viscoelastic features and conversions of synthetic cellulose were investigated. At first, it was confirmed that neither viscoelastic change nor cellobiose consumption was found with regard to all the protic acids in the SEE-free system. In the case of using the SEE biocatalyst prepared from previously deactivated cellulase, no consumption of cellobiose was detected by HPLC; thus the conversion ratios calculated from the residual cellobiose were kept ca. 0% after 24 h incubation. Therefore, it was elucidated that enzyme-mediated catalysis is essential for the synthesis and chain elongation of cellulose, while a nonenzymatic reaction, including acid hydrolysis to produce free glucose, did not occur at a low concentration of acid cocatalysts. The oNBSA, mNBSA, and pNBSA could act as cocatalysts to provide the required viscosity. Among them, the oNBSA exhibited the strongest viscosity, despite all three NBSA regioisomers having almost the same pKa value.25 By contrast, poor viscosities were obtained with either HCl, pTSA, or TFMSA, similar to that in the acid-free scenario. Acid concentrations ranging from 15−60 mM were investigated, with the greatest viscosities being obtained with 30 mM sulfuric acid or 15 mM NBSA. However, the sulfuric acid is a much stronger acid than NBSA. Therefore, this system probably involves rather complicated processes, not depending simply on the acid strength, and it is assumed that both proton availability and the nature of conjugate base can affect the SEE-mediated biocatalysis efficiency. Figure 4 compares the time-lapse conversion profiles for the sulfuric acid-assisted, oNBSA-assisted, and acid-free systems;

Figure 2. FTIR spectra of synthetic product (a) and regenerated cellulose sample (b).

characteristic peaks at 1056 cm−1 indicate the C−O bond stretching vibration of C−O−C group in the anhydroglucose ring.23 In a 1H NMR spectrum, the characteristic chemical shift of cellulose acetate was confirmed for acetylated samples (Figure 3),2 strongly suggesting the formation of cellulose. The

Figure 3. 1H NMR spectrum of acetylated product.

degree of acetylation was determined to be ca. 2.89 from the 1H NMR data. In addition, on the acid hydrolysis of the synthetic products, the glucose content reached up to ca. 94% in each case, and the rest might be partially hydrolyzed cellooligomers. It was reported that the aqueous hydrolysis with HCl could not thoroughly decompose long-chain cellulose to glucose monomers.17 As-synthesized insoluble fractions presumably occupy most of the dry weight of the chemical components of synthetic cellulose, and may be regarded as the practical yield. Therefore, the synthesized product was presumably identified to be cellulose. Additionally, high DPv and DPw values were obtained for synthetic cellulose (Table S1). The average molecular weight reached a DPv of ca. 120, being greater than previous record.7 The significant polymerization was proved in the SEE system. These results strongly suggest the successful synthesis of long-chain cellulose via a protic acid-assisted chain elongation catalysis in a one-step reaction system. SEE-Mediated Catalytic Behavior with Various Protic Acid Cocatalysts. Catalytic behavior of the cooperative catalysis depends both on the properties of each catalyst and their compatibility with one another.24 The influences of protic

Figure 4. Time-lapse conversion profiles for cellobiose via the SEEmediated biocatalysis, for the sulfuric acid-assisted (●), oNBSAassisted (■), acid-free (▲), and SEE-free (×) systems. The conversion ratios were quantitatively determined by HPLC from the amounts of residual cellobiose at a given time.

the conversion behavior was remarkably improved upon the addition of acid cocatalysts. For SEE in the absence of acid, the consumption levels of cellobiose were ca. 14% (entry 33 in Table S1, Supporting Information) after 24 h incubation. On the other hand, upon the addition of sulfuric acid, the conversion reached ca. 26% (entry 11 in Table S1). More noteworthy, the acid-free SEE system (entry 33 in Table S1) led to a large amount of water-soluble oligomers, and only ca. 5% fraction for long-chain cellulose was obtained, while watersoluble fractions were negligible for acid-assisted systems. Thus, the substantial conversion ratios increased by 5-fold through the assistance of a protic acid cocatalyst. In the case of oNBSA, ca. 18% (entry 20 in Table S1) conversion was observed after 24 h incubation. These values are significantly higher than those 2719

dx.doi.org/10.1021/bm3006775 | Biomacromolecules 2012, 13, 2716−2722

Biomacromolecules

Article

enveloped with surfactants and buffer salts, then suspended in organic media. In our previous studies, we have reported that the pH of the water phase of the water-in-oil emulsion during SEE preparation significantly influenced the catalytic efficiency.7,8,10 Zaks and Klibanov have reported that enzymes in organic media possessed a memorized ionogenic state corresponding to the pH in the final aqueous solution.27 In the present study, carbonate buffer at pH 10.5 was used for SEE preparation, and thus both carboxyl groups at the catalytic center of cellulase must be deprotonated at least at an initial stage. In a nonaqueous system, any oxygen atoms in the hydroxyl groups, hemiacetal groups, and glycosidic bonds within the cellobiose may be coordinated to protons. When the hemiacetal hydroxyl oxygen is coordinated to a proton, the cellobiose becomes a favorable activated intermediate for enzymatic condensation reactions, having an analogous form as produced by general acid hydrolysis (Scheme 1). Conversely, it is presumed that the proton coordinated to the C4 hydroxyl group at the nonreducing end of cellobiose inhibits the dehydration condensation reaction. Thus, it is reasonable that there is an optimum level of acid required with regard to a more efficient glycosynthesis. Such a proposed catalytic mechanism, however, has the following problems to be solved: (1) the proton coordination to DMAc and (2) the effect of a protic acid on the SEE biocatalyst. The DMAc is an aprotic organic solvent and acts as a proton acceptor to form a conjugate acid with first priority in this system. On the other hand, it was reported that as-formed [DMAc−Li]+ complexes strongly bind to the hydroxyl groups of cellulose for complete dissolution.28 Thus, such combination of protic and aprotic species possibly provided some positive contribution to promote the protonation and activation of cellobiose conjugated to the [DMAc−Li]+ complex, being effective for the following dehydration reaction. Protic acids also have a great influence on the enzymatic activity, which is, in general, sensitive to pH in aqueous media. However, in this nonaqueous system, the cellulase with buffer salts is significantly protected by nonionic surfactants to acquire the high solvent tolerance, and eventually kept active even in the nonaqueous LiCl/DMAc solvent,8 which is too severe for enzymes. In a similar manner, acid cocatalysts may exert an influence on the enzymatic activity of SEEs; however, some acid tolerance is also expected because an acid-assisted SEE-mediated reaction was more effective to achieve higher conversion ratios, higher TOF values, and longer chain elongation, as compare to an acid-free SEE system. Thus, it was presumed that acid cocatalysts containing DMAc−proton conjugate acids had little influence on the SEE biocatalyst. Besides, oNBSA made a great contribution to an SEEmediated biocatalysis, although mNBSA and pNBSA exhibited less efficiency. In cellulase-mediated hydrolytic reactions, two carboxyl groups in the active center of cellulase play an important role on the formation of transition states of enzyme− substrate complex. At an initial stage, both carboxyl groups in SEEs are deprotonated, and then activated cellobiose possibly interacted with the deprotonated carboxyl group, as shown in Scheme 1. Eventually, the activation efficiency of cellobiose and/or the proton supply to the carboxyl groups in SEEs might partly depend on the steric structures of NBSAs. Some issues still remain to be solved, and will be elucidated in our future work. Consideration on Enzymatic Kinetics. A clear correlation between the TOF values and the concentrations of reaction

with SEE alone. In the absence of the SEE biocatalyst, no reaction occurred, even in the presence of sulfuric acid. Thus, a cooperative reaction between the SEE and the protic acid allows us to promote the chain-elongation reaction. TOF Behavior. The TOF values for the SEE biocatalysis, with and without an acid cocatalyst, were investigated. In this study, the apparent TOF values were obtained by quantitatively measuring the consumption of cellobiose, on the assumption that the apparent enzymatic activity of crude cellulase in SEE is mainly derived from Cel7A, which plays a dominant role in the dehydration condensation reaction at the initial stage. In our previous study,8 the SEE containing Cel7A produced cellulose with average DP ca. 20 in ca. 15% yield. By contrast, the SEECel7B provided longer-chain cellulose with DP ca. 60 in very low yield of ca. 3%. Thus, cellobiose was preferentially consumed by Cel7A, and subsequent chain elongation reaction between as-synthesized oligomers was promoted by Cel7B. Thus, the TOF values were calculated by dividing the consumption of cellobiose by supplied cellulase, which corresponds to Cel7A at a rough estimate in an enzymatic activity. As shown in Table S1, the TOF value for the sulfuric acid-assisted SEE biocatalysis reached ca. 19 h−1 (entry 9) under the optimum reaction conditions; 30 mM sulfuric acid, 150 mM cellobiose and 30 mg cellulase in SEE with a 2 h incubation. The TOF values were very sensitive to the type and amount of acid cocatalyst added, with unique optimal conditions for each case. Entries 9 and 11 in Table S1 provided high conversion ratios, although no products were obtained for entries 3 and 4. Thus, the molar ratio of cellobiose and acid has a striking effect on reaction efficiency. For further details, an increase in cellobiose concentration leads to an increase in TOF, with and without the addition of an acid cocatalyst. The as-designed 1.0 wt % LiCl/DMAc solvent can dissolve up to ca. 150 mM cellobiose; the maximum TOF values are shown under each condition at this concentration (entries 9, 18, and 31 in Table S1). Ogata et al. have reported that a higher substrate concentration can favor the reverse reaction even in aqueous media, i.e., dehydration reaction for glycosynthesis.26 In other words, the direction of this reaction depended on a substrate concentration due to an equilibrium law. In our reaction system, nonaqueous media is advantageous for a dehydration reaction; additionally, higher cellobiose concentration led to higher conversion ratios. Therefore, it was suggested that nonaqueous biocatalysis possibly proceeded according to an equilibrium law. The DPv and DPw values do not match each other as absolute ones; there is a clear difference found due to analytical methods. This is because partial decomposition must occur during acetylation to achieve the high degree of acetylation (ca. 2.89) as reported in our previous paper.7 Proposed Reaction Mechanism. In our reaction, the protonation of cellobiose at the C1 hydroxyl group of a reducing end would be a key step to initiate the “reverse hydrolysis” reaction (route 1 in Scheme 1). The catalytic mechanisms of cellulases in normal aqueous systems have been well characterized and comprehensively reviewed.13 Cellulase has two carboxyl groups at the active center: one is deprotonated, and the other is protonated at optimal, neutral pH regions. In the presence of strong acids, the protonation of both carboxyl groups must occur; however, in this nonaqueous aprotic media, it remains unknown whether the direct protonation of carboxyl groups of enzymes inside SEEs may occur via strong acids, or not. In this study, cellulase is 2720

dx.doi.org/10.1021/bm3006775 | Biomacromolecules 2012, 13, 2716−2722

Biomacromolecules

Article

lases, activated sugar derivatives are beforehand synthesized, and then the reactions are kinetically controlled in watercontaining media; that is, transglycosylation of the activated donor to an acceptor sugar proceeds prior to inherent hydrolysis reaction of polysaccharides. By contrast, in this study, the protonation to hydroxyls groups of cellobiose for in situ activation would occur at first; however, a small amount of water added completely stopped the reverse hydrolysis reaction, as described above. Therefore, based on the clear influences of water and cellobiose contents on the reaction efficiency, it is presumed that the SEE-mediated reverse hydrolysis reaction may be thermodynamically controlled, without taking into consideration the gradual deactivation of SEE biocatalysts. In general, enzymatic reaction rates are very low in glycosynthesis using glycohydrolases, resulting in much difficulty in determining the accurate kinetic parameters. Nevertheless, to the best of our knowledge, we have successfully calculated the rough TOF values by quantitatively monitoring the decrease in a concentration of cellobiose, for the first time in glycosynthesis using glycohydrolases. Enzymatic Features. The crude cellulase from Trichoderma viride contains CBHs and EGs, processive and nonprocessive type, respectively.30 For sulfuric acid, the reaction solution exhibited negligible viscosity for the initial 2 h, but gradually became more viscous after a 4 h incubation period. A sequential oligomerization proceeded primarily via CBHs in the SEE for the initial 2 h. Once the oligomers were produced, the EGs in the SEE began to connect the oligomers to generate a long chain, resulting in high viscosity. In our previous study, the SEE from pure Cel7A (CBH) provided short-chain cellulose within a DP of ca. 20. Longer-chain cellulose with a DP of ca. 60 was obtained using SEE from pure Cel7B (EG).8 However, the apparent yields were only 2−3%. We also reported a synergetic effect between the two types of cellulases: CBHs and EGs may promote the synthesis of long-chain cellulose with a DPv of ca. 100 in 5% yield.7 In the present work, commercial crude cellulase from Trichoderma viride was used to prepare the SEE biocatalysts. Therefore, the novel proton-assisted nonaqueous/ microaqueous biocatalysis using the SEE biocatalysts in aprotic organic media can provide good yields (ca. 26%) of long-chain cellulose from cellobiose with no need for chemical preactivation, with a high DP up to ca. 120 in a direct, highly regioselective, one-pot reaction.

components was not observed (Table S1), due to the complicated situation in the SEE system. At first, in our previous study, it was reported that the enzymatic activity of SEE biocatalyst was kept more than 65% after 3 h incubation; however, the activity gradually decreased with time.8 Therefore, a simple kinetic model is difficult to directly apply for the SEEmediated enzymatic reaction. At this stage, the reusability of SEE biocatalyst is also limited. However, in this system, an increase in the cellobiose concentration led to an increase in apparent TOF values, with or without the addition of an acid cocatalyst (Table S1). In general, glycohydrolases have relatively high Km values for hydrolyzed components, such as cellobiose for cellulase, than that for original polysaccharides.29 Therefore, the reaction rates in reverse hydrolysis, i.e., dehydration condensation, would greatly depend on the concentration of component sugars. As mentioned above, a higher substrate concentration can favor the dehydration reaction for glycosynthesis,26 and a similar trend was observed in our system. In an SEE-mediated biocatalysis, efficient dehydration reaction proceeded in water-free media; thus it was assumed that the excess water must act as an inhibitor. In fact, a small amount of water added to the reaction mixture had a great negative influence on the conversion ratios of the products. In the case of adding 0.1 v/v% (ca. 56 mM) of water in LiCl/ DMAc, the conversion ratios drastically decreased to less than 10%, and 0.2 v/v% (ca. 111 mM) water completely stopped the reverse hydrolysis reaction. After 2 and 24 h incubations with sulfuric acid, the conversion ratios reached ca. 18% and ca. 26%, possibly implying ca. 27 mM and ca. 39 mM water produced in theory, respectively. Therefore, it is reasonable that the TOF values decrease with time, and thus initial TOF values should be evaluated at the earlier stage as much as possible, while keeping quantitative reliability. The water content is a striking factor for an SEE-mediated biocatalysis to promote the sugarchain elongation reaction, and the removal of undesirable water from the reaction systems would be effective for more efficient reverse hydrolysis reactions. In this study, the TOF values are theoretically lower than the real ones because SEE biocatalysts can catalyze reactions between the oligomers to form long-chain cellulose, while a constant cellobiose concentration is maintained. Then, the DP values of synthetic cellulose probably increase with time due to polymer−polymer condensation reaction, even at a constant concentration of cellobiose. However, water produced during reactions must inhibit the chain elongation reaction, and thus keeping a constant water content is required to estimate the time-dependent DPs. Thus, at this stage it cannot be expected to obtain clear correlations among conversion ratios, TOF values, and DPs. Hiraishi et al. have reported the synthesis of cellooligomers from glucose and α-D-glucose 1-phosphate by using cellodextrin phosphorylase.5 The kinetic parameters in the glycosynthesis direction could not be determined, although the detailed kinetic data for phosphorolysis was obtained. In the chain elongation direction, it was quite difficult to conclude the kinetic parameters since as-synthesized oligosaccharides became other acceptors for glycosylation, in addition to the fact that the enzymatic activity for transglycosylation is, in general, very low. On the other hand, in this study, protic acids were used as a cocatalyst; optimal acid concentrations to achieve higher TOF values differed much, depending on the types of protic acids. In conventional chemo−enzymatic reactions using glycohydro-

■

CONCLUSIONS We have achieved the efficient enzymatic condensation reaction of cellobiose in its original form to synthesize long-chain cellulose via a protic acid-assisted SEE-mediated biocatalysis in aprotic nonaqueous media. The conversions of synthetic cellulose polymer with more than 120 of DP reached ca. 26%, which is much higher than those previously reported (ca. 5%). The simple, in situ activation of cellobiose using an acid cocatalyst would make great impact on the improvement of biocatalytic efficiency. A virtually infinite number of combinations can be envisaged in terms of enzymes, substrate sugars, acid cocatalysts, and reaction media choice to find potential applications in glycomaterial-engineering fields.

■

ASSOCIATED CONTENT

S Supporting Information *

Table S1 for conversion ratios, TOF, and DP values of synthetic cellulose from cellobiose in an SEE-mediated 2721

dx.doi.org/10.1021/bm3006775 | Biomacromolecules 2012, 13, 2716−2722

Biomacromolecules

Article

(24) Knowles, R.; Jacobsen, E. Proc. Natl. Acad. Sci. U.S.A. 2010, 107, 20618−20619. (25) French, D.; Crumrine, D. J. Org. Chem. 1990, 55, 5494−5496. (26) Ogata, M.; Murata, T.; Murakami, K.; Suzuki, T.; Hidari, K.; Suzuki, Y.; Usui, T. Bioorg. Med. Chem. 2007, 15, 1383−1393. (27) Zaks, A.; Klibanov, A. M. Science 1984, 224, 1249−1251. (28) McCormick, C.; Callais, P.; Hutchinson, B. Macromolecules 1985, 18, 2394−2401. (29) Nidetzky, B.; Zachariae, W.; Gercken, G.; Hayn, M.; Steiner, W. Enzyme Microb. Technol. 1994, 16, 43−52. Igarashi, K.; Samejima, M.; Eriksson, K.-E. Eur. J. Biochem. 1998, 253, 101−106. (30) Rouvinen, J.; Bergfors, T.; Teeri, T.; Knowles, J.; Jones, T. Science 1990, 249, 380−386.

glycosynthesis with or without a protic acid is provided. This material is available free of charge via the Internet at http:// pubs.acs.org.

■

AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]; Tel and fax: +81 92 642 2993. Notes

The authors declare no competing financial interest.

■

ACKNOWLEDGMENTS This research was supported by a Grant-in-Aid for Young Scientists (S: 21678002) from the Ministry of Education, Culture, Sports, Science and Technology, Japan (T.K.).

■

REFERENCES

(1) Varki, A. Cell 2006, 126, 841−845. (2) Nakatsubo, F.; Kamitakahara, H.; Hori, M. J. Am. Chem. Soc. 1996, 118, 1677−1681. (3) Kadokawa, J. Chem. Rev. 2011, 111, 4308−4345. (4) Kobayashi, S.; Kashiwa, K.; Kawasaki, T.; Shod, S. J. Am. Chem. Soc. 1991, 113, 3079−3084. (5) Hiraishi, M.; Igarashi, K.; Kimura, S.; Wada, M.; Kitaoka, M.; Samejima, M. Carbohydr. Res. 2009, 344, 2468−2473. (6) Kaneko, Y.; Matsuda, S.; Kadokawa, J. Biomacromolecules 2007, 8, 3959−3964. (7) Egusa, S.; Kitaoka, T.; Goto, M.; Wariishi, H. Angew. Chem., Int. Ed. 2007, 46, 2063−2065. (8) Egusa, S.; Kitaoka, T.; Igarashi, K.; Samejima, M.; Goto, M.; Wariishi, H. J. Mol. Catal. B: Enzym. 2010, 67, 225−230. (9) Okutani, Y.; Egusa, S.; Ogawa, Y.; Kitaoka, T.; Goto, M.; Wariishi, H. ChemCatChem 2010, 2, 950−952. (10) Esaki, K.; Yokota, S.; Egusa, S.; Okutani, Y.; Ogawa, Y.; Kitaoka, T.; Goto, M.; Wariishi, H. Biomacromolecules 2009, 10, 1265−1269. Egusa, S.; Yokota, S.; Tanaka, K.; Esaki, K.; Okutani, Y.; Ogawa, Y.; Kitaoka, T.; Goto, M.; Wariishi, H. J. Mater. Chem. 2009, 19, 1836− 1842. (11) Eigen, M. Angew. Chem., Int. Ed. Engl. 1964, 3, 1−19. (12) Cabiac, A.; Guillon, E.; Chambon, F.; Pinel, C.; Rataboul, F.; Essayem, N. Appl. Catal. A: Gen. 2011, 402, 1−10. (13) Koshland, D. Biol. Rev. Cambridge Philos. Soc. 1953, 28, 416− 436. Davies, G. J.; Henrissat, B. Structure 1995, 3, 853−859. (14) Xu, H.; Zuend, S.; Woll, M.; Tao, Y.; Jacobsen, E. Science 2010, 327, 986−990. (15) Bruggink, A.; Schoevaart, R.; Kieboom, T. Org. Process Res. Dev. 2003, 7, 622−640. Kraußer, M.; Winkle, T.; Richter, N.; Dommer, S.; Fingerhut, A.; Hummel, W.; Gröger, H. ChemCatChem 2011, 3, 293− 296. (16) Goto, M.; Matsumoto, M.; Kondo, K.; Nakashio, F. J. Chem. Eng. Jpn. 1987, 20, 157−164. (17) Isogai, T.; Yanagisawa, M.; Isogai, A. Cellulose 2008, 15, 815− 823. (18) Tosh, B.; Saikia, C.; Dass, N. Carbohydr. Res. 2000, 327, 345− 352. (19) Smith, D. K.; Bampton, R. F.; Alexander, W. J. Ind. Eng. Chem., Process Des. Develop. 1963, 2, 57−62. Isogai, A.; Mutoh, N.; Onabe, F.; Usuda, M. Sen-i Gakkaishi 1989, 45, 299−306. (20) Samejima, M.; Sugiyama, J.; Igarashi, K.; Eriksson, K.-E. Carbohydr. Res. 1998, 305, 281−288. (21) Richter, G. A.; Glidden, K. E. Ind. Eng. Chem. 1940, 32, 1122− 1228. (22) Lateef, H.; Grimes, S.; Kewcharoenwong, P.; Feinberg, B. J. Chem. Technol. Biotechnol. 2009, 84, 1818−1827. (23) Murakami, M.; Kaneko, Y.; Kadokawa, J. Carbohydr. Polym. 2007, 69, 378−381. 2722

dx.doi.org/10.1021/bm3006775 | Biomacromolecules 2012, 13, 2716−2722