Solubilization Mechanism and Characterization of the Structural

Apr 11, 2012 - *E-mail: [email protected]. .... Solid-, Solution-, and Gas-state NMR Monitoring of 13C-Cellulose Degradation in an Anaerobic Microb...
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Solubilization Mechanism and Characterization of the Structural Change of Bacterial Cellulose in Regenerated States through Ionic Liquid Treatment Keiko Okushita,†,§ Eisuke Chikayama,†,§ and Jun Kikuchi*,†,‡,§,∥ †

RIKEN Plant Science Center and ‡Biomass Engineering Program, RIKEN Research Cluster for Innovation, 1-7-22 Suehiro-cho, Tsurumi-ku, Yokohama 235-0045, Japan § Graduate School of Nanobioscience, Yokohama City University, 1-7-29 Suehirocho, Tsurumi-ku, Yokohama 230-0045, Japan ∥ Graduate School of Bioagricultural Sciences and School of Agricultural Sciences, Nagoya University, 1 Furo-cho, Chikusa-ku, Nagoya-shi 464-8601, Japan S Supporting Information *

ABSTRACT: A statistical approach was used to characterize the heterogeneous structures of bacterial cellulose samples pretreated with four kinds of ionic liquids (ILs). The structural heterogeneity of these samples was measured by Fourier transform infrared spectroscopy as well as solid-state NMR methods such as cross-polarization magic-angle spinning and dipolar-assisted rotational resonance. The obtained data matrices were then evaluated by principal components analysis. The measured 1-D data clearly revealed the modification of crystalline cellulose; in addition, the statistical approach revealed subtle structural changes that occurred upon pretreatment with different kinds of ILs. To investigate whether such regenerated structural changes occurred because of solubilization, we examined the intermolecular nuclear Overhauser effect between cellulose and an IL. Our results clarify how the nucleophilic imidazole is attacked and suggest that the cation of the IL is associated with the collapse of hydrogen bonds in cellulose.

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INTRODUCTION Cellulose is the most abundant biopolymer on earth. Given that the technologically important physical properties of cellulose fibers and their behavior in chemical reactions are related to the supermolecular structure of the fiber polymers, molecular ordering has been a focus of research for many years.1,2 Natural cellulose is a partially crystalline polymer of 1−4 linked β-Dglucose residues. In native cellulose I type fibers, chains of cellulose aggregate and form fibrils that are deposited in the cell wall during biosynthesis.3,4 Crystalline cellulose has an amphipathic structure; that is, the cellulose chains include a hydrophobic surface consisting of pyranose-ring hydrogens, and the hydrophilic surface corresponds to the hydroxyl groups directed toward both sides of the ring. In addition, ordered © 2012 American Chemical Society

regions are generally assumed to alternate with less-ordered regions in these fibrils.5 This cellulose polymorph, whose structure is derived from its hydrogen bonding pattern, is extremely complex. The usefulness of cellulose would be enhanced if this structural packing by hydrogen bonds could be controlled. One way to modify these hydrogen bonds is to treat cellulose with ionic liquids (ILs). Room-temperature ILs, defined as a class of low-melting-point organic salts, are considered to be “green” and recyclable alternatives to many traditional volatile Received: January 5, 2012 Revised: April 5, 2012 Published: April 11, 2012 1323

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cellulose), α-cellulose, and cellopentaose were purchased from Fluka Biochemika, Nacalai Tesque, and Seikagaku, respectively. In addition, 1-butyl-3-methylimidazolium acetate (BmimAc) and 1-ethyl-3-methylimidazolium (EmimAc) acetate were obtained from SIGMA-Aldrich, and 1-ethyl-3-methylimidazolium chloride (EmimCl) and 1-ethyl-3methylimidazolium diethylphosphate (EmimDEP) were purchased from Kanto Chemical. Sample Preparation. Solid samples of pretreated IL were prepared as follows. Pieces of BC pellicles (10 mg of 13C or 12C) were dissolved in an IL (1 g) at 120 °C. After 30 min, the cellulose was regenerated by adding 2 mL of pure water to the mixture while agitating with a vortex mixer. The regenerated cellulose mixture was filtered and washed with pure water 10 times and then dried in vacuum for 6 h at room temperature.25−27 The BC samples easily dissolved or swelled in each IL at 120 °C to form a heterogeneous brown liquid. The viscosities of BmimAc and EmimDEP were quite high at room temperature but decreased at 120 °C, allowing the BC samples to dissolve easily. EmimCl was solid at room temperature but was a clear liquid with low viscosity at temperatures above 80 °C. When the liquid was heated for 30 min, the cellulose appeared as a clear brown precipitate. When water was added to the mixture of BC and IL, a fiber-like material appeared (regeneration of solid BC). The solid isolated by filtration was initially brown or orange but became almost white after washing several times with Milli-Q water. A sample for solution-state NMR analysis was prepared by heating the mixture of 13C BC and EmimAc/DMSO-d6 in an NMR tube as follows.28 After mixing EmimAc (8.5 wt %) and DMSO-d6 (1.5 wt %) to produce a mixed solvent, 4.5 mg of d6-sodium 2,2-dimethyl-2silapentane-5-sulfonate (d6-DSS) was dissolved in 450 μL of the mixed solvent (EmimAc/DMSO-d6) as the chemical shift reference. This solution (450 μL) and 13C BC (75 mg) were added to the NMR tube in small steps. The final concentration was calculated to be 13.90 wt % because of sampling loss (13C BC: 71.9 mg, solvent: 445.5 mg). The NMR tube was then heated in an incubator to 120 °C for 2 h. Infrared Spectroscopy. Regenerated solid samples were compressed between anvils with a hand press. Attenuated total reflectance (ATR) Fourier transfer infrared (FTIR) spectra were obtained by collecting 512 scans (resolution: 4 cm−1) in the mid-infrared region (ATR-MIR) and 128 scans (resolution: 16 cm−1) in the near-infrared (NIR) region with a Nicolet 6700 FTIR spectrometer (Fisher Scientific, Yokohama, Japan). The absorbances of individual ATRMIR spectral bands were normalized by the square root of the sum of the squared absorbances over 3560−950 cm−1 (MIR), whereas those of NIR spectral bands were normalized by the absorbance summed over 7200−6100 cm−1 (NIR). Their observed spectra were smoothed by using OMIC software for spectral noise reduction before the normalization. Second-derivative spectra were calculated by the Savitzky-Golay method for MIR29,30 with OMIC software and normalized by their z-scores. Solid-State NMR Spectroscopy. Solid-state NMR experiments were performed with an Avance 800 standard-bore spectrometer (800 MHz Bruker-BioSpin, Billerica, MA) with a Bruker 4-mm magic-angle spinning (MAS) triple resonance probe. The rotor was filled with about 100 mg of 13C BC or 10 mg of IL-pretreated BC by using PTFE thread seal tape (AS ONE) as a spacer. The MAS frequency was fixed at 12 kHz. The contact time for CP/MAS31 and the recycle delay were set at 1.2 ms and 3 s, respectively. The 90° 13C pulse length for decoupled MAS (DD/MAS)32 was 4.5 μs, and the recycle delay was 20 s. Dipolar-assisted rotational resonance (DARR) spectra were measured with a mixing time of 1.25 ms and a recycle delay of 3 s.33−35 DARR recoupling was accomplished by applying 1H continuous wave irradiation with a 1H radio frequency intensity equal to twice the spinning frequency, that is, 24 kHz. Quadrature detection was achieved with the time-proportional phase increment method. A total of 256 t1 increments were collected for an f1 spectral width of 300 ppm. Two-dimensional DARR spectra were processed with NMRPipe software. A variable-amplitude CP sequence was used for 2D experiments. The contact time was 1.2 ms. Glycine (specifically,

organic solvents. They have unique physicochemical properties such as negligible vapor pressure, high thermal stability, a wide liquid range, and tunable solvation properties.6,7 Recently, it has been reported that certain ILs can readily dissolve cellulose.8−10 However, the mechanism of cellulose dissolution in ILs is not understood. In particular, conclusions regarding the role of IL cations in the cellulose dissolution mechanism remain inconsistent. Some 13C and 35/37Cl nuclear magnetic resonance (NMR) relaxation studies have suggested that the naturally high chloride content and activity in [C4mim]Cl are the main reasons for the ability of this IL to dissolve cellulose.11,12 The nonhydrated and strongly hydrogen-bonded chloride ions are thought to disrupt the hydrogen-bonding network present in the cellulose polymer, thereby facilitating its dissolution in the IL. There are no specific interactions between IL cations and sugar solutes. Molecular simulation studies have also suggested that the dominant contribution to the sugar−IL interaction energy comes from favorable hydrogen-bonding interactions between the hydroxyl and chloride groups. However, they additionally indicate that cations form weak hydrogen-bonding and van der Waals interactions with glucose.13 When chemicals including natural metabolites such as amino acids, sugars, and phenylpropanoids are polymerized, the macromolecules can form tertiary structures with characteristic physicochemical properties. Spectroscopic analysis methods such as NMR and infrared (IR) spectroscopy have been used extensively to characterize the structure of macromolecules. Together with previous studies, spectral assignments determined with a spectral database enable qualitative and quantitative analyses. By using the appropriate pulse technique together with the 13C labeling technique,14−16 NMR allows one to probe the local structure of a macromolecule. In particular, in complex systems such as plant cell walls, which comprise the representative biomass, cross-polarization (CP) spectral editing in solid-state NMR measurement has been performed by using relaxation time differences that reflect the local structure and mobility.17−21 To date, the main approach to spectroscopic analysis has been to analyze a single signal. Many IR analyses of cellulose have focused only on the intensities of individual signals, for example, that of the 1370, 710, or 670 cm−1 band.22 NMR spectral analyses of cellulose have revealed its structure on the basis of C4 chemical shifts.23 Recently, analytical approaches with a “bird’s-eye-view”adequate to obtain an overview of the entire spectral regionhave been augmented to extract additional information. Life sciences researchers, who frequently deal with information on heterogeneous molecular mixtures, have successfully used statistical approaches such as principal component analysis (PCA). In the present work, we used NMR and IR data to characterize the structural changes in bacterial cellulose (BC) samples modified by ILs by using PCA. Four kinds of lowviscosity ILs were chosen as model systems. To consider the mechanism of the structural modification, we investigated molecular interactions between BC and an IL in the solubilized state via the intermolecular nuclear Overhauser effect (NOE), which is related to the distance between molecules.

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MATERIALS AND METHODS

Materials. 13C and 12C samples of BC were prepared by stationary cultivation of Acetobacter xylinum in Hestrin-Schramm (HS) medium24 supplemented with 13C6-glucose (13C > 99%). The pellicles were cut into pieces with a food cutter. Avicel PH-101 (microcrystalline 1324

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Figure 1. (a) PCA results for ATR-MIR spectra (top left: score plot, bottom left: loading plot). The samples are color-coded as follows. BC is yellow; BC pretreated by EmimAc, BmimAc, EmimCl, or EmimDEP is pink, green, purple, or light blue, respectively; Avicel is white; and α-cellulose is gray. Each sample was measured three times (n = 3). The second-derivative spectra of the OH region (2700 to 3500 cm−1) for the three kinds of sample (1: α-cellulose, 2: BC, 3: BC pretreated by EmimAc) are shown in panel b. The OH region had a remarkable pattern in the loading plot. Cellopentaose (4) is shown as the reference in panel b. the 13C chemical shift of its carbonyl carbon at 176.03 ppm) was used as the external reference. Solution-State NMR Spectroscopy. Solution-state NMR experiments were performed on a DRX-500 standard-bore spectrometer (500 MHz, Bruker Biospin, Billerica, MA) with a 5 mm BBI probe. Two-dimensional 1H−13C HSQC (the hsqcetgp pulse program from the Bruker library) and HSQC-NOESY36 spectra were measured at 120 °C with the recycle delays set at 2.5 s. Quadrature detection was achieved with the echo-antiecho method. A total of 384 t1 increments were collected for an f1 spectral width of 180 ppm. The mixing times of HSQC-NOESY were 50, 125, 250, 500, and 1000 ms. Their chemical shifts were measured in relation to the d6-DSS peak at 0 ppm, which was used as an inert reference. Adding 1.70 ppm to the DSSreferenced carbon chemical shift referenced the chemical shift to the tetramethylsilane (TMS) peak. Statistical Calculations. The R package was used to perform PCA. The basic strategy and method for the statistical calculation of spectroscopic data have been reported elsewhere.14,37−39 Before performing PCA, 1D solid-state NMR spectra were first normalized by the summed areas from 55−108 ppm (722 points). 2D DARR spectra were processed for PCA analysis by the NMRPipe software,40 and binned matrices were created for f1 and f2 over 55−115 ppm by using the FT2DB webtool (https://database.riken.jp/ecomics/) to acquire their intensities. The intensities were normalized by the summed intensities of all peaks in this region. (Binned areas and schemes are shown in Figure S1 of the Supporting Information.)

Information) to obtain structural information of BC pretreated with various ILs. In the score plots (Figure 1a, top) calculated from each spectrum, the pre- (Figure 1a, top, BC, Avicel, αcellulose) and post-IL-pretreated samples (Figure 1a, top, Emim and Bmim series) were clearly distinguished along the PC1 direction. In the loading plot (Figure 1a, bottom), markedly changing regions along PC1 appeared in the ranges of 980−1160 and 3200−3600 cm−1 corresponding to the C−O− C and OH stretching regions, respectively.29,30 IR has been widely used as a method for the solid-state analysis of polymers such as cellulosic materials. The ATR-MIR spectrum of cellulose has OH stretching vibrations in the 3200−3600 cm−1 range, indicative of hydrogen bonding. For this reason, it has been used to investigate higher-order structural changes.29,30 The second-derivative spectrum of the OH stretching band region is shown in Figure 1b. The second-derivative spectra of both α-cellulose (Figure 1b, 1) and BC (Figure 1b, 2) showed peaks from 3200 to 3400 cm−1 that indicate crystalline form I. The OH band of the IL-pretreated BC samples (Figure 1b, 3) showed two peaks (3442 and 3490 cm−1) characteristic of the crystalline form II.41−43 The second-derivative spectra of the MIR spectra (Figure 1b) revealed that the bands assigned to type II crystalline cellulose41−43 only appeared after IL pretreatment. These peaks are similar to those for cellopentaose (3444 and 3492 cm−1, Figure 1b, 4). This indicates that the hydrogen-bonding state of IL-pretreated BC samples is similar to that of cellopentaose. PCA was also carried out on the NIR spectra (Figure S3 of the Supporting Information). BC with high crystallinity appeared on the negative side of PC1 in the score plot. The 6450 cm−1 band assigned to the type I crystalline form44 had a negative value of PC1 in the loading plot. IL-pretreated BC samples assigned as amorphous material44 had positive values of PC1 around 7000 cm−1 in the loading plot. With regard to PC2, the band assigned to crystalline regions (6287 cm−1) and

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RESULTS AND DISCUSSION Statistical Approach for Structural Characterization. We employed a statistical approach to characterize the structural changes in BC by using multiple spectroscopic instruments. The MIR, NIR, and NMR results revealed that ILs broke the hydrogen bonds of highly crystalline BC. During the regeneration step when water was added, BC was predominantly transformed into an amorphous material with some type II crystalline cellulose. Statistical Analysis of FTIR Spectra. PCA was carried out on the ATR-MIR spectra (Figure 1 a, Figure S2 of the Supporting 1325

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that assigned to semicrystalline regions (6718 cm−1) appeared with negative and positive values, respectively, in the loading plot. The entire spectral region was simultaneously analyzed to characterize the structural modifications from type I crystalline to type II crystalline and amorphous forms. The findings are consistent with previously published IR band assignments,29,30,41−43 and verify the applicability of the statistical approach to characterizing the structural change of cellulose in biomass polymers. Assuming that a more complete use of NMR data would enable us to extract more detailed information, we then investigated chemical shifts as indicators of structural change. Statistical Analysis of 1D CP/MAS and DD/MAS Spectra. Figure 2 shows the solid-state 13C CP/MAS (a) and DD/MAS

of BC (Figure 2, yellow spectra) from those of other samples (Figure 2, red, green, blue, and purple spectra) or the spectra of Ac-series samples (Figure 2, red and green spectra) from those of other IL-pretreated samples (Figure 2, blue and purple spectra) more clearly than the CP/MAS (Figure 2a) and DD/ MAS spectra (Figure 2b) of each sample. This tendency was also visualized clearly in the score plot (Figure 3a, left), in which PC1 separated 13C BC (Figure 3a, left, yellow circles) from the other samples, and the difference between ILs (Figure 3a, left, green, pink, blue, and purple circles) used for pretreatment was reflected in PC2. In the spectra of BC (Figure 2, yellow spectra), sharp peaks of mainly type Iα crystalline cellulose were observed at 105.2 (C1), 89.0 (C4), 74.7 (C3), and 65.4 ppm (C6). In all spectra (Figure 2), broad peaks due to the amorphous form were observed at ca. 84 (C4) and 62 ppm (C6). The amorphous peaks and content were broader and greater, respectively, for the IL-pretreated BC samples. This is because amorphous material was the main component of the C4 and C6 signals in the spectra of the IL-pretreated BC samples (Figure 2, red, green, blue, and purple spectra). Among the IL-pretreated BC samples, EmimAc and BmimAc displayed relatively sharp peaks in the C1, C4, and C6 regions (Figure 2, red, and green spectra). The spectral shapes were notably different in the C4 region between IL-pretreated BC samples. In the loading plot (Figure 3a, right), the differences between pre- and post-ILpretreatment (PC1) were marked at C2 (71.3 ppm, Iα), C4 (87−90 ppm), and C6 (65.4 ppm, Iα). The main effect of the ILs used for pretreatment (PC2) appeared at C4 and C6 for the type II crystalline (C4: 87.5 ppm, C6: 63.0 ppm) and amorphous (C4: 80−85 ppm, C6: around 61 ppm) peaks. As expected, PCA of 1D spectra (Figure 3a) revealed that C4 carbon signals behaved as chemical shift markers. Furthermore, in previous reports, the crystalline ratio of cellulose has been determined by conventional solid-state NMRan approach widely used for quantitative analyses of polymeric cellulose, especially of the C4 signal.23 Chemical shifts were assigned based on previously reported values.45,46 Statistical Analysis of 2D DARR Spectra. Two-dimensional DARR measurements were performed to obtain additional information on cellulose structure (Figure 3b). Such spectra have been used for spectral assignment and to obtain distance information.33−35 In the present study, PCA was carried out on the spectra with a mixing time of 1.25 ms, which provided information over relatively short distances. In the score plot (Figure 3b, left), PC1 and PC2 showed trends similar to those of PCA results for the 1D spectra (Figure 3a, left). The exception was EmimDEP (Figure 3a, left, blue circle), which was clustered into a group different from EmimAc and BmimAc (Figure 3a, left, pink and green circles, respectively) in 1D PCA. Namely, the clustering pattern of the score plot changed substantially between 1D and 2D PCA (Figure 3a,b). The 2D spectra are composed of f1 and f2 directions, that is, expansion of f2-1D spectra along the f1 direction. In the present study, the 70−80 ppm region was difficult to separate by using only 1D spectra, whereas the C2, C3, and C5 peaks in this region of f2 were clearly separated by 2D measurement. The PCA loading plots of 2D DARR spectra also revealed numerous loading components along the f1 direction (Figure 3b). In the score plot (Figure 3b), differences between the four kinds of IL were characterized along PC2. Considering the FTIR results and the C4 chemical shift of a previous paper,47 the negative and positive sides of PC2 show tendencies toward type II crystalline

Figure 2. (a) 13C CP/MAS spectra and (b) 13C DD/MAS spectra of 13 C BC (yellow) and 13C BC samples pretreated by four kinds of IL (red: EmimAc, green: BmimAc, purple: EmimCl, light blue: EmimDEP). The MAS frequency was fixed at 12 kHz. A spacer peak appeared at ca. 110 ppm of the DD/MAS spectra of ILpretreated BC samples.

(b) spectra of 13C BC and IL-pretreated BC samples. PCA was performed on the solid-state 1D NMR spectra (CP/MAS, DD/ MAS) (Figure 3a). Solid-state NMR spectra contain a great deal of information about the superstructure. The peak intensity of CP/MAS spectra reflects both the molecular amount and molecular mobility (and/or other interactions), whereas that of DD/MAS reflects the molecular amount more directly. As discussed and shown in a previous paper,19 spectral regions with some mobility, such as hydrated pectin in a plant cell wall, are expected to have high intensities in the same chemical shift in DD/MAS spectra (in the paper, single-pulse excitation with MAS, SPEMAS). Therefore, mobility changes can be evaluated by signal intensity differences between CP/ MAS and DD/MAS, whereas structural differences between polymorphs (crystalline types Iα, Iβ, and II) are expected to exhibit different chemical shifts. In this Article, we focused on changes of line shapes in the solid-state NMR spectra derived specifically from structural changes. To emphasize line shape changes as relative values, we normalized the spectra by the sum of the intensities. This approach distinguished the spectra 1326

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Figure 3. PCA results composed of score plots and loading plots for (a) CP/MAS and DD/MAS spectra, as well as for (b) 2D DARR spectra (score plots, left; loading plots, right). 2D DARR spectra are accompanied by observed NMR spectra (panel b, right, top-left half). 13C BC (yellow) and 13C BC samples pretreated by four kinds of IL (red: EmimAc, green: BmimAc, purple: EmimCl, light blue: EmimDEP). The MAS frequency was fixed at 12 kHz. The method of 2D PCA is shown in Figure S2 of the Supporting Information. In the score plot (panels a and b, left), PC1 separated 13C BC from the other samples, and the difference between ILs used for pretreatment was reflected in PC2. In the loading plot of 2D DARR (panel b, right, bottom-right half), the C2, C3, and C5 peaks in f2 were clearly separated by 2D measurement along the f1 direction. The clustering pattern of the score plot (panel b, left) changed substantially after separating this region along the f1 direction.

chemical shifts of the type II crystalline form were definitively assigned by Kono et al,47 those of the amorphous structure remain controversial. In the loading plot (Figure 3b), the negative color (orange) corresponds to type II crystalline signals of the original spectra, and the positive color (pale blue) is attributed to the amorphous signals. IL-pretreated BC includes the polymorphs described above, and the loading plot of the 2D DARR spectrum shows each structural component as a separate color. PCA of the 2D DARR spectra is particularly useful for assigning signals from unknown components in complex systems (e.g., biomass polymers). In previous studies of biomass, especially of plant cell walls, there have been a number of attempts to separate the peak components in 1D solid-state NMR spectra by using relaxationedited CP/MAS17,18,20,21 with different relaxation times for atoms in different structural or chemical environments. In plant cell walls, the solid-state NMR signals of polysaccharides such as cellulose, hemicelluloses, and pectin largely overlap.19 However, as investigated in a model system mixing α-cellulose with pectin, “rigid” cellulose is associated with a long T1H or T1ρH, whereas pectin and hemicelluloses are associated with short T1H or T1ρH.18 The relaxation times of plant cell walls have been reported to differ among components; for example, for the primary cell wall of Arabidopsis thaliana, the T1ρH (C6) values of cellulose and hemicellulose have been reported as 21.6

and amorphous (EmimCl) forms, respectively, although such differences in score plots may largely be caused by structural differences in C2, C3, and C5 of amorphous forms. Also, from the relaxation point of view, the tendency of the PC2 (Figure 3) was suggested as described above. The relaxation times (T1H), which become lower with increase in molecular mobility in the solid state, of the main component in BC, EmimAc, BmimAc, EmimCl, and EmimDEP-pretreated BC sample were 554.9, 713.1, 791.7, 375.6, and 694.5 ms, respectively (Table S1 of the Supporting Information). Applicability of 2D PCA to 2D Spectral Separation. In the observed 2D DARR spectra (Figure 3b, right, top-left half), there were peaks of type I and II crystalline and amorphous forms. In the 2D spectrum of BC (Figure 3b, right, top-left half, gold line), peaks corresponding to the type Iα crystalline form were observed clearly. In the spectra of IL-pretreated BC samples, peaks corresponding to the type II crystalline form were observed, and peaks that were expected as amorphous peaks from their C4 chemical shifts were also observed. In the loading plot (Figure 3b, right, bottom-right half), the values of PC1 range from red (negative) to black (positive), and those of PC2 range from orange (negative) to blue (positive). Peaks corresponding to 13C BC resulted in red for PC1, and those corresponding to type II crystalline47 and amorphous forms resulted in orange and blue, respectively, for PC2. Although 1327

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Figure 4. HSQC (blue) and HSQC-NOESY36 (red) spectra of 13C BC in EmimAc/DMSO-d6 (13.90 wt %) at 120 °C. (a) C2, 3, 4, and 5 regions (mixing time = 250 ms) of cellulose with spectral assignments. (b) Observed NOE region (mixing time = 50 ms) between C−H of cellulose and 2H of Emim. (c) NOE peak intensity. The intensity of C6−H is shown at half value because there are two hydrogens (2H) at C6 of cellulose. Two NOE signals due to JCC coupling with its neighboring carbon were observed for cellulose C2−H. The lower NOE peak intensity of the cellulose C2−H is shown in panel c.

Table 1. Chemical Shift Assignments of Cellulose in EmimAc at 120°Ca δH / ppm δC / ppm a

C1

C2-1

C2-2

4.74 106.66

3.28 78.55

3.28 78.02

C3 3.68 78.90

3.68 79.42

C4

C5

C6-1

C6-2

3.66 82.94

3.52 79.95

3.99 64.49

3.84 64.67

Reference is TMS.

and 11.0 ms, and the corresponding T1C (C4) values have been reported as 4.2 and 3.0 s, respectively.48 In a previous study, a method that uses differences of the proton or carbon relaxation times that depend on domain structures and dynamics was applied to the model plant Arabidopsis thaliana.20 In addition, in 2D spectra, pulse sequences considering differences in molecular mobility have been used to observe components that are difficult to detect in CP.48 Therefore, a 2D sequential technique based on differences in relaxation times is also suggested to be valid for separating components. A statistical approach to NMR spectral analysis called statistical total correlation spectroscopy (STOCSY) is also well-known in the metabonomics field for the rapid visualization and identification of biomarkers.49 STOCSY has also been applied to diffusion ordered spectroscopy (DOSY) spectra that give a physicochemical property of diffusion constant.50 A method in which a relaxation-edited approach and a statistical approach such as STOCSY are combined would be valid for extracting latent components in solid-state NMR spectra more effectively. In this Article, we used PCA solely as a statistical approach to separate components and did not focus on the correlative changes with relaxation time. A preliminary investigation of a statistical method for edited spectra based on a relaxation measurement has just been initiated elsewhere.51 IL Solubilization State of BC. Solution-state analysis by NMR (Figure 4) was explored to identify the mechanism that changed BC into a different structure (Figures 1b and Figure 2) after regeneration from the various ILs. Differences in the

profile of the solid-state NMR spectra (Figure 3) were likely derived from differences in hydrogen bonding. During preparation of the solid samples, BC was dissolved in each IL and then regenerated; hydrogen bonding of the isolated BC pretreated with the IL may be related to the state of BC in the IL. Among the ILs used for BC pretreatment, EmimAc yielded the most typical properties. Therefore, we investigated the state of BC dissolved in an IL by using EmimAc as a representative example. Chemical Shift Assignment of Solubilized Cellulose by Solution-State NMR. Chemical shifts of polymeric cellulose were assigned from the HSQC spectrum measured at 120 °C.52 The C3−H and C5−H peaks for terminal residues were found between 79 and 80.5 ppm along the f1 direction (Figure 4a, blue). The blue peaks in Figure 4a show the HSQC spectrum in the range of C2 to C5 in the dissolved BC. H2, H3, H4, and H5 were observed at the 1H chemical shifts of 3.28, 3.68, 3.66, and 3.52 ppm, respectively; the assigned chemical shifts are listed in Table 1. The chemical shift of C1 was at the lowest field. In our analysis, 1H and 13C chemical shifts were assigned for native polymeric cellulose. An investigation of the interactions between BC and IL became possible because of these chemical shift assignments. Molecular Interaction Between BC and IL by Observation of Intermolecular NOEs. HSQC-NOESY measurements of 13C BC in EmimAc/DMSO-d6 were performed at 120 °C (Figure 4a−c). The mixing times for the measurements were 50 (Figure 4b), 125, 250 (Figure 4a), 500, and 1000 ms. This experiment 1328

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was performed to acquire structural information between BC and IL because NOE signals can offer information on the approximate distance between the 13C−1H of an HSQC spectrum and another 1H, provided that the location of the latter proton is near that of the former proton. A longer mixing time increases the likelihood of detecting NOE signals between distant atoms in an HSQC-NOESY experiment. The red peaks in Figure 4a show the HSQC-NOESY spectrum in the region of C2 to C5 at a mixing time of 250 ms. The observed NOE peaks in Figure 4a are H6-1/H4 (δH = 3.99 ppm/δC = 83.10 ppm), H6-2/H4 (3.84/82.94), H5/H4 (3.52/ 83.10), H2/H4 (3.27/82.94), H6-1/H5 (3.99/79.82), H6-2/ H5 (3.84/79.80), H3 or H4/H5 (3.66/79.78), H2/H5 (3.28/ 79.78), and H4/H2 (3.67/78.16). The red peaks in Figure 4b show the HSQC-NOESY spectrum at a mixing time of 50 ms, in which NOEs occurred between the C−H of BC and the 2H of the Emim cation. The observed NOE peaks in Figure 4b are Emim-H2/cellulose-H1 (δH = 10.31 ppm/δC = 106.47 ppm), Emim-H2/cellulose-H4 (10.31/82.94), Emim-H2/cellulose-H5 (10.31/79.95), EmimH2/cellulose-H2-1 (10.31/78.54), and Emim-H2/celluloseH2-2 (10.31/78.01). The peaks of the HSQC-NOESY spectrum were assigned based on Table 1. In the C2−C5 region (Figure 4a), NOEs of the same pyranose ring (H2/H4 and H6/H5) were observed along with other NOEs from cellulose (i.e., H5/H4, H2/H5). Therefore, NOEs between neighboring residues were likely observed at a mixing time of 250 ms. The high NOE intensity peaks of the HSQC-NOESY experiments with a mixing time of 125 ms relate to H1/H3, H1/H4, and H1/H5 (Figure S4 of the Supporting Information), which belong to the same pyranose ring and 1→4 glycoside linkage. Figure 4b shows NOEs between BC and the 2H of the 1ethyl-3-methylimidazolium (Emim) cation at a mixing time of 50 ms. NOEs linked to the Emim cation were observed at C6− H, C2−H, C5−H, and C1−H of BC. Therefore, all NOEs indicated that the 2H atom of the Emim cation is near C2−H, C1−H, and C4−H of cellulose. This is also supported by the spectra at the short mixing times of 125 and 250 ms. The NOE of C2−H with the 2H of the Emim cation had the highest intensity at a mixing time of 50 ms; in contrast, at a mixing time of 125 ms, the NOE for C4−H with the 2H of the Emim cation had the highest NOE intensity. However, if the lower intensities of two cellulose C2−H peaks were excluded, then the NOE for C2−H with the 2H of the Emim cation had the highest intensity (Figure S4 (c) of the Supporting Information); at mixing times of 250 to 1000 ms, the NOEs for C1−H and C4−H with the 2H of the Emim cation had relatively high NOE intensities (Figure 4c). This means that C2−H, C1−H, and C4−H of cellulose are closer to the 2H of the Emim cation than to other cellulose protons. In previous studies, the mechanism of BC dissolution by ILs was shown to be related to the nucleophilic property of the Cl− ion and the electrophilic property of imidazole.12,53−55 Distance data from NOESY experiments provide information on the structural features involved in the breaking of hydrogen bonds in BC. The NOEs of 2H-Emim were observed for C2−H, C4− H, and C1−H of BC (Figure 4). The results showed that the 2H of Emim was located nearest the 1H of BC and that the C2−H distance was the shortest. This suggests that Emim may participate in breaking the intrachain O6···H2 hydrogen bonds or in stabilizing the approach of the anion to O6···H2 by its van der Waals forces and π-CH stacking near the C2−H of BC.55,56

Each IL used in this study contained the imidazole group, whose state in solution should not greatly change between ILs. Further studies on the effect of the anion in the BC dissolution process are warranted. The keys to understand the mechanisms by which regenerated structures change may reside not only in the dissolved state but also in the regeneration and drying steps.

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CONCLUSIONS In summary, we monitored the IL-induced structural modification of cellulose I (BC) to a complex of cellulose II and amorphous forms with spectroscopic instruments such as FTIR and solid-state NMR. The ratio of components differed between the ILs used for pretreatment. An overview of the spectra by statistical analysis would highlight its characteristic regions systematically. Therefore, by performing statistical analysis (PCA), we characterized the spectral differences induced by IL-pretreatment in score plots and emphasized the characteristic marker regions in loading plots, consistent with previous reports. A statistical analysis of 2D solid-state NMR spectra also showed potential support for the assignment of unknown peaks. We aimed to clarify the mechanism of ILinduced changes so that ILs could be used to change the structure of cellulose structures systematically. The solutionstate NMR results indicated the possibility that the cation of an IL is related to the breaking of hydrogen bonds. We believe that analyses similar to those in the present study may lead to the creation of engineering materials with specific properties by controlling the hydrogen bonds without the need for chemical derivatization.

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ASSOCIATED CONTENT

S Supporting Information *

T1H values measured by solid-state NMR; the method of 2D data profiling (i.e., how to change 2D spectra into numerical matrices); PCA results with the observed spectra for ATR-MIR and NIR, respectively; and HSQC and HSQC-NOESY spectra (mixing time = 125 ms), including the peaks of C6−H and C1−H as well as additional information on NOE peak intensities. This material is available free of charge via the Internet at http://pubs.acs.org.

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AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS We thank Mr. Tetsuya Mori, Mr. Takanori Komatsu, and Ms. Yuuri Tsuboi for helpful discussions and providing highly washed bacteria cellulose. This research was partially supported by Grants-in-Aid for Scientific Research for challenging exploratory research (J.K.) and from the Ministry of Education, Culture, Sports, Science, and Technology, Japan. This work was also partially supported by grants from the New Energy and Industrial Technology Development Organization (NEDO to J.K.).

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REFERENCES

(1) Roy, D.; Semsarilar, M.; Guthrie, J. T.; Perrier, S. Chem. Soc. Rev. 2009, 38, 2046−2064. (2) Putaux, J. L. Macromol. Symp. 2005, 229, 66−71. 1329

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(3) Atalla, R. H.; Vanderhart, D. L. Science 1984, 223, 283−285. (4) Williamson, R. E.; Burn, J. E.; Hocart, C. H. Trends Plant Sci. 2002, 7, 461−467. (5) Klemm, D.; Schumann, D.; Kramer, F.; Hessler, N.; Hornung, M.; Schmauder, H. P.; Marsch, S. Adv. Polym. Sci. 2006, 205, 49−96. (6) El Seoud, O. A.; Koschella, A.; Fidale, L. C.; Dorn, S.; Heinze, T. Biomacromolecules 2007, 8, 2629−2647. (7) Mora-Pale, M.; Meli, L.; Doherty, T. V.; Linhardt, R. J.; Dordick, J. S. Biotechnol. Bioeng. 2011, 108, 1229−1245. (8) Xie, H.; King, A.; Kilpelainen, I.; Granstrom, M.; Argyropoulos, D. S. Biomacromolecules 2007, 8, 3740−3748. (9) Kilpelainen, I.; Xie, H.; King, A.; Granstrom, M.; Heikkinen, S.; Argyropoulos, D. S. J. Agric. Food Chem. 2007, 55, 9142−9148. (10) Ohno, H.; Fukaya, Y. Chem. Lett. 2009, 38, 2−7. (11) Remsing, R. C.; Liu, Z.; Sergeyev, I.; Moyna, G. J. Phys. Chem. B 2008, 112, 7363−7369. (12) Remsing, R. C.; Swatloski, R. P.; Rogers, R. D.; Moyna, G. Chem. Commun. (Cambridge, U. K.) 2006, 1271−1273. (13) Liu, H.; Sale, K. L.; Holmes, B. M.; Simmons, B. A.; Singh, S. J. Phys. Chem. B 2010, 114, 4293−4301. (14) Sekiyama, Y.; Chikayama, E.; Kikuchi, J. Anal. Chem. (Washington, DC, U. S.) 2010, 82, 1643−1652. (15) Sekiyama, Y.; Kikuchi, J. Phytochemistry (Elsevier) 2007, 68, 2320−2329. (16) Sekiyama, Y.; Chikayama, E.; Kikuchi, J. Anal. Chem. (Washington, DC, U. S.) 2011, 83, 719−726. (17) Tekely, P.; Vignon, M. R. J. Polym. Sci., Part C: Polym. Lett. 1987, 25, 257−261. (18) Tang, H. R.; Wang, Y. L.; Belton, P. S. Solid State Nucl. Magn. Reson. 2000, 15, 239−248. (19) Tang, H. R.; Belton, P. S.; Ng, A.; Ryden, P. J. Agric. Food Chem. 1999, 47, 510−517. (20) Newman, R. H.; Davies, L. M.; Harris, P. J. Plant Physiol. 1996, 111, 475−485. (21) Bootten, T. J.; Harris, P. J.; Melton, L. D.; Newman, R. H. Biomacromolecules 2009, 10, 2961−2967. (22) Akerholm, M.; Hinterstoisser, B.; Salmen, L. Carbohydr. Res. 2004, 339, 569−578. (23) Malm, E.; Bulone, V.; Wickholm, K.; Larsson, P. T.; Iversen, T. Carbohydr. Res. 2010, 345, 97−100. (24) Hestrin, S.; Schramm, M. Biochem. J. 1954, 58, 345−352. (25) Dadi, A. P.; Varanasi, S.; Schall, C. A. Biotechnol. Bioeng. 2006, 95, 904−910. (26) Dadi, A. P.; Schall, C. A.; Varanasi, S. Appl. Biochem. Biotechnol. 2007, 137, 407−421. (27) Ha, S. H.; Ngoc, L. M.; An, G. M.; Koo, Y. M. Bioresour. Technol. 2011, 102, 1214−1219. (28) Moulthrop, J. S.; Swatloski, R. P.; Moyna, G.; Rogers, R. D. Chem. Commun. (Cambridge, U. K.) 2005, 1557−1559. (29) Watanabe, A.; Morita, S.; Ozaki, Y. Biomacromolecules 2007, 8, 2969−2975. (30) Watanabe, A.; Morita, S.; Ozaki, Y. Biomacromolecules 2006, 7, 3164−3170. (31) Pines, A.; Shattuck, T. W. J. Chem. Phys. 1974, 61, 1255−1256. (32) Vanderhart, D. L.; Earl, W. L.; Garroway, A. N. J. Magn. Reson. (1969−1992) 1981, 44, 361−401. (33) Ohashi, R.; Takegoshi, K. J. Chem. Phys. 2006, 125, 214503. (34) Takegoshi, K.; Nakamura, S.; Terao, T. J. Chem. Phys. 2003, 118, 2325−2341. (35) Takegoshi, K.; Nakamura, S.; Terao, T. Chem. Phys. Lett. 2001, 344, 631−637. (36) Majumdar, A.; Zuiderweg, E. R. P. J. Magn. Reson., Ser. B 1993, 102, 242−244. (37) Nakanishi, Y.; Fukuda, S.; Chikayama, E.; Kimura, Y.; Ohno, H.; Kikuchi, J. J. Proteome Res. 2011, 10, 824−836. (38) Fukuda, S.; Nakanishi, Y.; Chikayama, E.; Ohno, H.; Hino, T.; Kikuchi, J. PLoS One 2009, 4, e4893. (39) Tian, C.; Chikayama, E.; Tsuboi, Y.; Kuromori, T.; Shinozaki, K.; Kikuchi, J.; Hirayama, T. J. Biol. Chem. 2007, 282, 18532−18541.

(40) Delaglio, F.; Grzesiek, S.; Vuister, G. W.; Zhu, G.; Pfeifer, J.; Bax, A. J. Biomol. NMR 1995, 6, 277−293. (41) Michell, A. J. Carbohydr. Res. 1988, 173, 185−195. (42) Marchessault, R. H.; Liang, C. Y. J. Polym. Sci. 1960, 43, 71−84. (43) Sugiyama, J.; Horikawa, Y. Mokuzai Gakkaishi 2008, 54, 49−57. (44) Tsuchikawa, S.; Siesler, H. W. Appl. Spectrosc. 2003, 57, 667− 674. (45) Kono, H.; Erata, T.; Takai, M. Macromolecules (Washington, DC, U. S.) 2003, 36, 5131−5138. (46) Kono, H.; Numata, Y.; Erata, T.; Takai, M. Macromolecules (Washington, DC, U. S.) 2004, 37, 5310−5316. (47) Kono, H.; Numata, Y. Polymer 2004, 45, 4541−4547. (48) Dick-Perez, M.; Zhang, Y. A.; Hayes, J.; Salazar, A.; Zabotina, O. A.; Hong, M. Biochemistry 2011, 50, 989−1000. (49) Cloarec, O.; Dumas, M. E.; Craig, A.; Barton, R. H.; Trygg, J.; Hudson, J.; Blancher, C.; Gauguier, D.; Lindon, J. C.; Holmes, E.; Nicholson, J. Anal. Chem. (Washington, DC, U. S.) 2005, 77, 1282− 1289. (50) Smith, L. M.; Maher, A. D.; Cloarec, O.; Rantalainen, M.; Tang, H. R.; Elliott, P.; Stamler, J.; Lindon, J. C.; Holmes, E.; Nicholson, J. K. Anal. Chem. (Washington, DC, U. S.) 2007, 79, 5682−5689. (51) Okushita, K.; Komatsu, T.; Chikayama, E.; Kikuchi, J. Polym. J. 2012, in press. (52) Flugge, L. A.; Blank, J. T.; Petillo, P. A. J. Am. Chem. Soc. 1999, 121, 7228−7238. (53) Swatloski, R. P.; Spear, S. K.; Holbrey, J. D.; Rogers, R. D. J. Am. Chem. Soc. 2002, 124, 4974−4975. (54) Zhang, H.; Wu, J.; Zhang, J.; He, J. S. Macromolecules (Washington, DC, U. S.) 2005, 38, 8272−8277. (55) Youngs, T. G.; Hardacre, C.; Holbrey, J. D. J. Phys. Chem. B 2007, 111, 13765−13774. (56) Fernandez, M. D.; Canada, F. J.; Jimenez-Barbero, J.; Cuevas, G. J. Am. Chem. Soc. 2005, 127, 7379−7386.

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