Article pubs.acs.org/Biomac
Comparative Analysis of Crystallinity Changes in Cellulose I Polymers Using ATR-FTIR, X-ray Diffraction, and Carbohydrate-Binding Module Probes Alenka Kljun,†,⊥ Thomas A. S. Benians,‡,⊥ Florence Goubet,§ Frank Meulewaeter,§ J. Paul Knox,‡ and Richard S. Blackburn*,† †
Sustainable Materials Research Group, Centre for Technical Textiles, University of Leeds, Leeds LS2 9JT, U.K. Centre for Plant Sciences, Faculty of Biological Sciences, University of Leeds, Leeds LS2 9JT, U.K. § Bayer BioScience N.V., Technologiepark 38, Gent, Belgium ‡
ABSTRACT: Cotton fiber cellulose is highly crystalline and oriented; when native cellulose (cellulose I) is treated with certain alkali concentrations, intermolecular hydrogen bonds are broken and Na-cellulose I is formed. At higher alkali concentrations Na-cellulose II forms, wherein intermolecular and intramolecular hydrogen bonds are broken, ultimately resulting in cellulose II polymers. Crystallinity changes in cotton fibers were observed and assigned using attenuated total reflectance Fourier transform infrared (ATR FT-IR) spectroscopy and X-ray diffraction (XRD) subsequent to sodium hydroxide treatment and compared with an in situ protein-binding methodology using cellulose-directed carbohydrate-binding modules (CBMs). Crystallinity changes observed using CBM probes for crystalline cellulose (CBM2a, CBM3a) and amorphous cellulose (CBM4-1, CBM17) displayed close agreement with changes in crystallinity observed with ATR-FTIR techniques, but it is notable that crystallinity changes observed with CBMs are observed at lower NaOH concentrations (2.0 mol dm −3), indicating these probes may be more sensitive in detecting crystallinity changes than those calculated using FTIR indices. It was observed that the concentration of NaOH at which crystallinity changes occur as analyzed using the CBM labeling techniques are also lower than those observed using X-ray diffraction techniques. Analysis of crystallinity changes in cellulose using CBMs offers a new and advantageous method of qualitative and quantitative assessment of changes to the structure of cellulose that occur with sodium hydroxide treatment.
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aqueous sodium hydroxide above a certain concentration is able to penetrate the cellulose crystalline lattice to produce a series of well-defined crystalline complexes holding a number of sodium ions and water molecules within the lattice.6 Two hypotheses have been proposed for the transformation mechanism of cellulose I to cellulose II: namely, chain-polarity transformation (parallel to antiparallel)2,7 and chain-conformation transformation (bent to bent-twisted).8 Okano and Sarko9 presented that a total of five unique alkali-celluloses (Na-celluloses) could be generated reproducibly, depending on the alkali concentration used. These five structures were named Na-celluloses I, IIA, IIB, III, and IV. Studies of cellulose crystallinity and morphology have developed significantly during the past two decades, using new methods of structural analysis. Many researchers have employed infrared spectroscopy to study crystallinity in native cotton and regenerated cellulose. O’Connor et al.10 developed an empirical “crystallinity index”, but Hurtubise and Krässig11 observed that this “crystallinity index” was dependent not only
INTRODUCTION
Cellulose is a linear polymer composed of glucose units that are linked by β-1,4 glycosidic bonds formed between the carbon atoms C(1) and C(4) of adjacent glucose units. Cotton fibers are one of the purest sources of cellulose and the most important industrial natural fibers.1 The supramolecular structure of cellulosic fibers can be described by a two-phase model with regions of high orientation (crystalline) and low orientation (amorphous).2,3 Alkali has a significant effect on morphological, molecular, and supramolecular properties of cellulose, and it is well-known that when native celluloses are treated with strongly alkaline solutions, cellulose adopts a modified crystal structure, irreversibly forming cellulose II,2,4 which is easily understandable from the concept that the initial parallel chain crystal structure (cellulose I) changes into more stable antiparallel chains (cellulose II).2,3 These changes in the structure of cellulose upon alkali treatment are exploited extensively in the textile industry, usually in a process known as mercerization. Concentration of sodium hydroxide, temperature, tension, degree of polymerization, source of the cellulose, and physical state of the cellulose have an effect on the properties and degree of change upon treatment.4,5 In its interaction with cellulose, © 2011 American Chemical Society
Received: August 22, 2011 Revised: October 5, 2011 Published: October 7, 2011 4121
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on crystallinity but also on the degree of mercerization and suggested a more appropriate designation using their “lateral order index” (LOI). Using mixtures of unmercerized and mercerized cotton, they showed that the LOI varied linearly with the percentage of mercerized cellulose in the mixture. Nelson and O’Connor12 developed the “total crystallinity index” (TCI) and Nada et al.13 introduced the empirical “hydrogenbond intensity” (HBI), which is related to the crystal system and the degree of intermolecular regularity as well as the amount of bound water. Although the crystallinity of cellulose has been investigated for almost a century using X-ray diffraction (XRD) techniques,14−20 there are few publications that use XRD to quantify changes in the crystallinity of cotton fibers; one example is the work of Richter et al.21 on cotton linters. XRD presents several problems in quantitative analysis of crystallinity changes, particularly in relation to sample preparation. Often cotton fibers are unsuitable for measurement by XRD and have to be ground to a powder for analysis, which itself causes a change in crystallinity.22 Carbohydrate-binding modules (CBMs) are distinct protein domains, without catalytic activity, found in many carbohydrate-active enzymes that act on plant cell walls.23−25 CBMs promote the association of catalytic domains with substrate, and their main function is to increase the catalytic efficiency of the carbohydrate-active enzymes against soluble and/or insoluble substrates. CBMs with the capacity to bind cellulose are associated with enzymes that hydrolyze both cellulose and other plant cell wall polymers such as xylan, mannan, and noncellulosic glucans.26−28 Type A CBMs are able to bind to crystalline cellulose,29,30 while type B CBMs recognize different substructures within amorphous cellulose (see Experimental Section).31,32 It has been previously demonstrated that CBMs directly coupled to fluorophores or with His-tag appendages can be used to investigate aspects of cellulose structure in the context of cell wall composites.33,34 Here we report the use of CBMs in an indirect in situ labeling technique to monitor changes in the crystallinity of cellulose during the mercerization of cotton fibers. The results are compared with the analysis of crystallinity changes monitored using attenuated total reflectance Fouriertransform infrared (ATR FT-IR) spectroscopy and XRD.
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Table 1. Cellulose-Directed CBMs Used in This Study CBM probe
type
ligand
ref
CBM2a CBM3a CBM4-1 CBM17
A A B B
crystalline cellulose crystalline cellulose amorphous cellulose amorphous cellulose
37 38 39 40
ligand recognition site is conserved presenting a flat surface comprising predominantly aromatic residues, which interact with the multiple planar cellulose chains found in crystalline cellulose.29,30 Type B CBMs do not bind to the planar surface of crystalline polysaccharides but recognize individual saccharide chains. Three well-characterized examples of cellulose-binding type B CBMs are found in families 4, 17, and 28. The ligand-binding sites in these protein modules comprise extended clefts or grooves that accommodate individual glycan chains in noncrystalline regions of cellulose;32,33 CBMs from these families display similar cellulose binding properties, although competition studies indicate that these modules recognize different substructures within amorphous cellulose.36 Sodium Hydroxide Treatment. Cotton fibers were treated with water or one of a series of eight aqueous sodium hydroxide solutions from 1.0 to 8.0 mol dm−3, using a liquor ratio of 100:1, in an oscillating water bath (Grant OLS 200) at 25 °C and 45 rpm for 30 min. The fibers were then rinsed in clean water, dried in air, and then conditioned as detailed below prior to analysis. Attenuated Total Reflectance Fourier-Transform Infrared (ATR FT-IR). Samples were subjected to FTIR spectroscopy (at four different points in the sample) using a Perkin-Elmer Spectrum BX spotlight spectrophotometer with diamond ATR attachment. Scanning was conducted from 4000 to 600 cm−1 with 64 repetitious scans averaged for each spectrum. Resolution was 4 cm−1, and interval scanning was 2 cm−1. Prior to measurement, samples were conditioned in a standard atmosphere of 65 ± 2% relative humidity and 20 ± 2 °C for 48 h and then held in a desiccator over P2O5 to maintain the same atmosphere as the FTIR measurement equipment not located in the same place. Obtained spectra were normalized to the absorbance of the O−H in-plane deformation band at 1336 cm−1 due to any obtained changes in this band among all examined samples. LOI (α 1429/893) and HBI (α 3336/1336) were calculated as described in ref 41. X-ray Diffraction. Prior to measurement, samples were conditioned in a standard atmosphere of 65 ± 2% relative humidity and 20 ± 2 °C for 48 h and then held in a desiccator over P2O5 to maintain the same atmosphere. Cotton fibers for X-ray diffraction (XRD) were prepared in the form of flat layers of fibers laid parallel to each other, the layer being 16 mm in diameter and 3 mm deep. To keep fibers flat, pressure-sensitive tape was laid over the top of the sample holder. A Panalytical MPD diffractometer was used employing Cu Kα radiation (λ = 1.5406 Å), generated at a voltage of 40 kV and current of 40 mA. Diffractograms were made by continuous scanning over the range of diffraction angle 2θ from 5° to 30° for 7 min scanning time. The unit cell of both cellulose I and cellulose II is monoclinic; Miller indices for the three principal planes of reflection are (11 ̅0), (110), and (200) for cellulose I and (110̅ ), (110), and (020) for cellulose II. Crystallinity index (CrI) was calculated using either eq 1 (for cellulose I) or eq 2 (when cellulose I had transformed to cellulose II) from an internal reference method of Segal et al.:42
EXPERIMENTAL SECTION
Materials. Fibers of cotton (Gossypium hirsutum L.) lines Cooker (C312) and FM966 were used in this study. The cotton fibers used had the following characteristics. Cooker: mean fiber length 26.65 mm; strength 30.8 g tex−1; elongation at break 6.0%. FM966: mean fiber length 30.48 mm; strength 31.1 g tex−1; elongation at break 3.8%. These two different fibers were selected to determine if there were any differences in observed effects between fiber types. Before use the wax content of the cotton fibers was removed by incubating fibers at room temperature with excess ethanol for 90 min and again for 90 min in refreshed ethanol. The fibers were then incubated with acetone for 60 min, followed by incubation in ether for 60 min. The fibers were then left to air-dry. The CBMs used in this research, listed in Table 1, have previously been used to label plant materials24 and were supplied by Prof. Harry Gilbert, Department of Biological and Nutritional Sciences, Newcastle University. All other chemicals were supplied by Sigma-Aldrich. CBMs are grouped into sequence-based families in the CAZy database35 and are named after the family in which they are located (e.g., a family 4 CBM is designated CBM4). CBMs from families 1, 2a, 3a, 5, and 10 are classified as type A CBMs, which bind to crystalline polysaccharides, predominantly cellulose,25 as the topography of the
(1)
(2) where I200 is the maximum intensity of crystalline scatter at the 200 reflection (used in the case of cellulose I at 2θ = 22.5°), I11̅0 is the maximum intensity of the 110̅ reflection (used in the case of cellulose II at 2θ = 19.8°), and Iam is intensity of diffraction at 2θ = 18.0° for cellulose I and 2θ = 16.0° for cellulose II.42,43 4122
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Figure 1. In situ fluorescence imaging of the binding of four cellulose-directed CBMs to Cooker and FM966 cotton fibers after treatments with a range of NaOH concentrations (0−8 M). In Situ Fluorescence Imaging of CBM Recognition of Cotton Fibers. Prior to analysis of CBM binding to the surface of cotton fibers, the outer pectin layer of the fibers was removed by use of pectate lyase24 to increase access to cellulose. The pectate lyase treatment will not additionally affect the crystallinity as the conditions encountered are not sufficiently alkaline to effect a change (see ref 24). All CBMs used contain His-tags and CBM recognition and binding to fiber surfaces was analyzed using a three-step immuno-based procedure. Fiber samples (5 mg) were incubated in 1 cm 3 5% (w/v) milk powder/phosphate-buffered saline (MP/PBS) for 30 min and then washed with PBS. CBMs (20 μg cm−3) were incubated with fiber samples for at least 90 min at room temperature with shaking. Samples were washed three times with PBS, 5 min per wash. A secondary antibody (mouse antihis) was added, diluted 1 in 1000 in MP/PBS, and incubated for at least 90 min. After washing, fibers were incubated with a tertiary antibody (anti-mouse linked to fluorescein isothiocyanate [FITC]) diluted 1 in 50 in MP/PBS for at least 90 min. Samples were washed three times with PBS, 5 min per wash. Samples were mounted in Citifluor antifade to prevent fluorescence fading. Fluorescence imaging of fibers used an Olympus BX61 microscope equipped with epifluorescence irradiation and Volocity software. Fluorescence quantification was done using Image J software. Fluorescence intensity measurements were made of regions of representative fiber surfaces using a consistent ellipse shape. Three measurements were made from each of three micrographs for each treatment. Values were normalized to a value between 0 and 1 by dividing the data points by the maximum intensity reading.
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significant amount of crystalline cellulose I, the band is toward 1430 cm−1 and the amounts of cellulose II and amorphous cellulose decrease.4 Hydrogen bond intensity (HBI) can also be used to interpret qualitative changes in crystallinity in cellulose; generally, as HBI increases crystallinity decreases.13 An increase in HBI represents an increase in hydrogen bonding between certain hydroxyl functions in the cellulose, which is typical of the conversion of cellulose I to cellulose IIeven though this represents a decrease in overall % crystallinity, an increase in this particular hydrogen bonding increases. Nelson et al.12 found that the band at 1336 cm−1 showed the greatest differences between the crystalline cellulose II and the amorphous cellulose, and Kondo et al.45 demonstrated that these bands were not characteristic of the amorphous area. CBMs, with distinct molecular recognition capacities, have been used to locate in situ the presence of crystalline and amorphous regions of cellulose in plant materials.24 However, such biological techniques have not been used to quantify molecular changes in polymers subject to treatments. Figure 1 shows indirect immunofluorescence detection of CBMs binding to the surface of Cooker and FM966 cotton fibers after NaOH treatments equivalent to that used for the FT-IR analyses above. Four different CBMs were used to visualize the changes in crystallinity with increasing NaOH concentrations for both the Cooker and the FM966 fibers. CBM2a and CBM3a bind to crystalline cellulose, and it is observed in Figure 1 that the binding intensity of these CBMs to the fibers decreases with increasing NaOH treatment concentration, reflecting the decrease in crystallinity. It is notable that the major decrease in fluorescence signal was observed between 1.0 and 2.0 mol dm−3 NaOH treatments for both CBM2a and CBM3a. In contrast the recognition of fibers by CBM17 and CBM4-1, directed to amorphous cellulose, increases with increasing
RESULTS AND DISCUSSION
Lateral order index (LOI) can be used to interpret qualitative changes in cellulose crystallinity and is based on the ratio of absorbance bands at specific wavenumbers; generally, as LOI decreases crystallinity also decreases.4,10,11,44 The 1430 (1429) and 893 cm−1 absorption bands can be used to study the type of crystalline cellulose and the crystallinity changes because the crystalline cellulose I spectrum differs clearly in this band from cellulose II and amorphous cellulose.44 If a cellulose fiber has a 4123
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Figure 2. Quantification of CBM binding to cotton fibers and FTIR analyses in response to NaOH treatments. (a) Crystalline-directed CBM2a and CBM3a binding to Cooker cotton fibers with increasing NaOH concentration in comparison with LOI (ATR-FTIR). (b) Crystalline-directed CBM2a and CBM3a binding to FM966 cotton fibers with increasing NaOH concentration in comparison with LOI (ATR-FTIR). (c) Amorphous cellulose-directed CBM4-1 and CBM17 binding to Cooker cotton fibers with increasing NaOH treatment in comparison with HBI (ATR-FTIR). (d) Amorphous cellulose-directed CBM4-1 and CBM17 binding to FM966 cotton fibers with increasing NaOH treatment in comparison with HBI (ATR-FTIR). Error bars show ± standard deviation of fluorescence quantification.
2 mol dm−3 NaOH lower than that observed by LOI and also indicates a steeper transition between crystalline and amorphous forms. Figures 2c and 2d show changes in HBI with increasing NaOH concentrations, where it is observed that significant changes in HBI occur between 3.0 and 4.0 mol dm−3 NaOH, which is associated with a decrease in crystallinity and a transformation of cellulose I to cellulose II. Figures 2c and 2d also show that the quantification of amorphous-cellulose directed CBM4-1 and CBM17 binding in a comparison with the HBI data. In this case the two CBMs generated the same trend with each of two fibers in that there was a steady increase in binding with increasing concentrations of alkali particularly between 2.0 and 6.0 mol dm−3 NaOH. In this case the comparison with the HBI value, derived from FTIR analyses, indicated a less abrupt transition and loss of crystallinity as evidenced by the CBM recognition profiles. The CBMs are, of course, binding to the surface of the cotton fibers and cannot access the fiber interiors. In contrast the ATR-FTIR techniques are assaying throughout the fibers. It is of considerable interest that the binding of the crystalline cellulose-directed CBMs are particularly sensitive to changes resulting from the NaOH treatment presumably reflecting cellulose rearrangements with 2.0 mol dm−3 and thus indicating an absolute requirement for crystalline structures. In contrast, the amorphous-cellulose directed CBMs are also sensitive to
NaOH concentration treatments which again reflects the decrease in crystallinity. These observations are in agreement with the observations made using FT-IR as the analysis method and also will reflect the conversion of cellulose I (more crystalline) to the less crystalline cellulose II. Accordingly, the results suggest that CBM2a and CBM3a bind to both crystalline cellulose I and crystalline cellulose II. However, these micrographs shown in Figure 1 only provide qualitative analysis, and in order to provide quantitative analysis of these visualized changes in cellulose, crystallinity image analysis was used to convert fluorescence signals for each treatment into relative values. Figure 2 shows quantitative changes in binding of the four CBMs with increasing NaOH treatment concentrations, for Cooker and FM966 cotton fibers, respectively. Figures 2a and 2b show changes in LOI of cotton fibers with increasing NaOH concentration, where it is observed that significant changes in LOI occur with treatments between 2.0 and 4.0 mol dm−3 NaOH, which is associated with a decrease in crystallinity and a transformation of cellulose I to cellulose II. Above 4.0 mol dm−3 NaOH, no further crystallinity change was observed, probably as a result of cellulose being in the most energetically favorable state (cellulose II). In the case of CBM probes for crystalline cellulose (CBM2a, CBM3a) for both fibers analyzed (Figure 2a,b), the observed decline in binding reflects the crystallinity change as indicted by LOI but the loss of crystallinity is observed with treatment with up to 4124
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Figure 3. Crystallinity of cotton fibers with increasing sodium hydroxide concentration in comparison with crystallinity changes measured using X-ray diffraction: (a) Cooker cotton fibers using CBMs for crystalline cellulose (CBM2a, CBM3a); (b) FM966 cotton fibers using CBMs for crystalline cellulose (CBM2a, CBM3a). Error bars show ± standard deviation for fluorescence intensity values.
Author Contributions ⊥ These two authors contributed equally to the work.
the lower concentrations of alkali but also continue to increase in capacity to bind to the fiber surfaces in line with increasing NaOH concentrations. These two cellulose recognition capabilities are therefore quite distinct and do not merely appear to recognize two directly interconverted cellulose structures. These binding profiles also indicate the capacity for a range of states of cellulose structures at cotton fiber surfaces. Figure 3 shows the same crystalline cellulose directed CBM binding data in comparison with crystallinity changes observed for CrI values calculated after measurement using XRD techniques. Crystallinity values derived from XRD and the indication of the transition to noncrystalline cellulose with increasing NaOH concentration closely reflect those obtained from the FTIR analyses.
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ACKNOWLEDGMENTS The authors thank Bayer BioScience N.V., Belgium, for the provision of a scholarship to AK. The authors also thank the UK Biotechnology and Biological Sciences Research Council (BBSRC) and Bayer BioScience N.V. for the provision of a BBSRC CASE Award to T.A.S.B. We thank Harry Gilbert for the provision of the CBMs and useful discussions.
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REFERENCES
(1) Hsieh, Y. L. In Cotton: Science and Technology; Gordon, S., Hsieh, Y. L., Eds.; Woodhead: Cambridge, 2007. (2) Krässig, H. A. Cellulose: Structure, Accessibility and Reactivity; Krässig, H. A., Ed.; Gordon and Breach Science Publishers S.A.: Yverdon, 1993. (3) Bredereck, K.; Hermanutz, F. Rev. Prog. Color. Relat. Top. 2005, 35, 59−75. (4) Colom, X.; Carrillo, F. Eur. Polym. J. 2002, 38, 2225−2230. (5) Jaturapiree, A.; Manian, A. P.; Bechtold, T. Cellulose 2006, 13, 647−654. (6) Porro, F.; Bédué, O.; Chanzy, H.; Heux, L. Biomacromolecules 2007, 8, 2586−2593. (7) Okano, T.; Sarko, A. J. Appl. Polym. Sci. 1985, 30, 325−332. (8) Hayashi, J.; Yaginuma, Y. In Wood and Cellulosics; Kennedy, J. F., Phillips, G. O., Williams, P. A., Eds.; Ellis Horwood Ltd.: Chichester, 1987. (9) Okano, T.; Sarko, A. J. Appl. Polym. Sci. 1984, 29, 4175−4182. (10) O’Connor, R. T.; DuPré, E. F.; Mitcham, D. Text. Res. J. 1958, 28, 382−392. (11) Hurtubise, F. G.; Krässig, H. Anal. Chem. 1960, 32, 177−181. (12) Nelson, M. L.; O’Connor, R. T. J. Appl. Polym. Sci. 1964, 8, 1325−1341. (13) Nada, A.-A. M. A.; Kamel, S.; El-Sakhawy, M. Polym. Deg. Stab. 2000, 70, 347−355. (14) Nishikawa, S.; Ono, S. Proc. Tokyo Math. Phys. Soc. 1913, 7, 131−138. (15) Meyer, K. H.; Mark, H. Ber. Dtsch. Chem. Ges. 1928, 61, 593− 614. (16) Meyer, K. H.; Misch, L. Helv. Chim. Acta 1937, 20, 232−244. (17) Gardner, K. H.; Blackwell, J. Biopolymers 1974, 13, 1975−2001. (18) Sarko, A.; Muggli, R. Macromolecules 1974, 7, 486−494. (19) Nishiyama, Y.; Langan, P.; Chanzy, H. J. Am. Chem. Soc. 2002, 124, 9074−9082.
CONCLUSIONS
Using FT-IR and XRD in conjunction with series of cellulosedirected binding modules can provide useful assessments of the crystallinity status of cellulose in cotton fibers. The pair of crystalline-cellulose directed CBMs and a pair of amorphous cellulose-directed CBMs indicate that the crystallinity status of cotton fibers can be rapidly determined. As the CBMs bind to fiber surfaces, the modulation of binding is sensitive to crystallinity changes in the cotton at lower concentrations of alkali treatment and are thus more sensitive to detecting such crystallinity changes in comparison with values derived using FTIR or XRD indices. Analysis of crystallinity changes in cellulose using probing with CBMs offers a new method of qualitative and quantitative measurement of changes to the structure of cellulose that occur with sodium hydroxide treatment. The technique offers the advantages of not changing the crystallinity of the fiber in sample preparation (as seen with X-ray diffraction), the potential for increased sensitivity, and the ability to spatially analyze and map the surface of individual cotton fibers for crystallinity changes rather than having to evaluate the bulk change in a sample (as seen with ATR-FTIR).
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AUTHOR INFORMATION Corresponding Author *Tel: +44 113 343 3757. Fax: +44 113 343 3704. E-mail:
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(20) Nishiyama, Y.; Sugiyama, J.; Chanzy, H.; Langan, P. J. Am. Chem. Soc. 2003, 125, 14300−14306. (21) Richter, U.; Krause, T.; Schempp, W. Angew. Makromol. Chem. 1991, 185, 155−167. (22) Chidambareswaran, P. K.; Sreenivasan, S.; Patil, N. B. Text. Res. J. 1987, 57, 219−222. (23) Hashimoto, H. Cell. Mol. Life Sci. 2006, 63, 2954−2967. (24) Blake, A. W.; McCartney, L.; Flint, J. E.; Bolam, D. N.; Boraston, A. B.; Gilbert, H. J.; Knox, J. P. J. Biol. Chem. 2006, 281, 29321−29329. (25) Boraston, A. B.; Bolam, D. N.; Gilbert, H. J.; Davies, G. J. Biochem. J. 2004, 382, 769−781. (26) Kellett, L. E.; Poole, D. M.; Ferreira, L. M.; Durrant, A. J.; Hazlewood, G. P.; Gilbert, H. J. Biochem. J. 1990, 272, 369−376. ́ (27) Hogg, D.; Pell, G.; Dupree, P.; Goubet, F.; Marti n-Orú e, S. M.; Armand, S.; Gilbert, H. J. Biochem. J. 2003, 371, 1027−1043. (28) McKie, V. A.; Vincken, J. P.; Voragen, A. G.; van den Broek, L. A.; Stimson, E.; Gilbert, H. J. Biochem. J. 2001, 355, 167−177. (29) Xu, G.-Y.; Ong, E.; Gilkes, N. R.; Kilburn, D. G.; Muhandiram, D. R.; Harris-Brandts, M.; Carver, J. P.; Kay, L. E.; Harvey, T. S. Biochemistry 1995, 34, 6993−7009. (30) Raghothama, S.; Simpson, P. J.; Szabó, L.; Nagy, T.; Gilbert, H. J.; Williamson, M. P. Biochemistry 2000, 39, 978−984. (31) Boraston, A. B.; Nurizzo, D.; Notenboom, V.; Ducros, V.; Rose, D. R.; Kilburn, D. G.; Davies, G. J. J. Mol. Biol. 2002, 319, 1143−1156. (32) Jamal, S.; Nurizzo, D.; Boraston, A. B.; Davies, G. J. J. Mol. Biol. 2004, 339, 253−258. (33) McCartney, L.; Gilbert, H. J.; Bolam, D. N.; Boraston, A. B.; Knox, J. P. Anal. Biochem. 2004, 326, 49−54. (34) McCartney, L.; Blake, A. W.; Flint, J.; Bolam, D. N.; Boraston, A. B.; Gilbert, H. J.; Knox, J. P. Proc. Natl. Acad. Sci. U. S. A. 2006, 103, 4765−4770. (35) Cantarel, B. L.; Coutinho, P. M.; Rancurel, C.; Bernard, T.; Lombard, V.; Henrissat, B. Nucleic Acids Res. 2009, 37, D233−238. (36) McLean, B. W.; Boraston, A. B.; Brouwer, D.; Sanaie, N.; Fyfe, C. A.; Warren, R. A.; Kilburn, D. G.; Haynes, C. A. J. Biol. Chem. 2002, 277, 50245−50254. (37) Bolam, D. N.; Ciruela, A.; McQueen-Mason, S.; Simpson, P.; Williamson, M. P.; Rixon, J. E.; Boraston, A.; Hazlewood, G. P.; Gilbert, H. J. Biochem. J. 1998, 331, 775−778. (38) Tormo, J.; Lamed, R.; Chirino, A. J.; Morag, E.; Bayer, E. A.; Shoham, Y.; Steitz, T. A. EMBO J. 1996, 15, 5739−5751. (39) Tomme, P.; Creagh, A. L.; Kilburn, D. G.; Haynes, C. A. Biochemistry 1996, 35, 13885−13894. (40) Boraston, A. B.; Chiu, P.; Warren, R. A.; Kilburn, D. G. Biochemistry 2000, 39, 11129−11136. (41) Široký, J.; Blackburn, R. S.; Bechtold, T.; Taylor, J.; White, P. Cellulose 2010, 17, 103−115. (42) Segal, L.; Creely, J. J.; Martin, A. E. Jr.; Conrad, C. M. Text. Res. J. 1959, 29, 786−794. (43) Krässig, H. A. Cellulose: Structure, Accessibility and Reactivity; Gordon and Breach Science Publishers S.A.: Yverdon, 1993. (44) Carrillo, F.; Colom, X.; Suñol, J. J.; Saurina, J. Eur. Polym. J. 2004, 40, 2229−2234. (45) Kondo, T.; Sawatari, C. Polymer 1996, 31, 393−399.
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