Enzymatic Degradation of Lichenan Layers Adsorbed onto Regenerated Cellulose Surfaces Travis DePriest1, Xiao Zhang2 and Alan R. Esker2 1 Department of Physics and 2Department of Chemistry, Virginia Tech Blacksburg, Virginia, 24061, United States leeone@vt.edu Introduction Generally, secondary plant cell walls are composed of a network of cellulose, hemicelluloses, lignin, structural proteins, and a few lesser substances such as pectins and enzymes.1 Hemicelluloses are defined as plant polysaccharides in the cell wall with β-(1-4)-linked backbones in an equatorial conformation. 2 Hemicelluloses are grouped into xyloglucans, xylans, mannans, glucomannans, and β-(1→3,1→4)-glucans or mixed linkage glucans (MLGs).3 Grasses, cereals, and lichens are the main sources of MLGs which differ in structure and abundance among sources. Lichenans are MLGs with a linear chain isolated from varying species of lichens.3 Over 20,000 species of lichens are present over a wide range of habitats throughout the world.4 The most abundant source of lichenan is Cetraria islandica, where lichenan is 42% of the dry mass.3 As such, lichenan could be a successful alternative bioethanol feedstock in the absence of commercialized and centralized lignocellulosic technologies.4 Lichenan is made up of cellulose-like oligosaccharides of D-(1→4) linked glucose monomers that range from three to nine repeat units, separated by D-(1→3) linkages.5 The D-(1→3) linkages make the molecules flexible and soluble, while the D-(1→4) linkages are responsible for insolubility in cold water.3 Lichenan is composed of 86.35 % trisaccharides and 2.71% tetrasaccharides, making the trisaccharide:tetrasaccharide ratio 31.9:1, much higher than cereal MLGs.5 Pentasaccharides account for 6.82 % of the mass, while the remaining 4.13 % of lichenan is attributed to longer segments with degrees of polymerization from 6 to 9.5 Only considering two states, trisaccharides and tetrasaccharides, simplifies the theoretical treatments of lichenan. For these idealized chains the two oligosaccharides are randomly dispersed within the chain, a feature that might differ in longer segments.6 The gelation mechanism and solution conformation of lichenan are only partially understood. One theory proposes lichenan chains form paracrystalline or sheetlike structures associated through weak intermolecular bonding when regions of three or more consecutive trisaccharides occur along the chain.7Intermolecular hydrogen bonds arise between glucose monomers at C6 positions. Studies of 6% by mass gels in water in the literature supported claims of these regions, or junction zones and concluded an increase in the trisaccharide:tetrasaccharide ratio increased the density of potential interaction points.5 The greater the trisaccharide content of the MLG, the more elastic the gels became. The rheological characteristics of MLG solutions and gels are dependent upon the distribution of cellulosic oligomers, linkage patterns, molar mass, temperature, and concentration.8 Fine structure that affects solution properties are important in drug delivery systems such as nanocapsules, microcapsules, hydrogels, matrices, scaffolds, and gels. Variations in fine structure of lichenan based biopolymer blends9 could allow controlled and sustained drug release of drugs with or without enzyme incorporation for unique degradation-delivery processes. Lichenase or 1,3-1,4-β-D-glucanase (E.C.3.2.1.73) is an enzyme which specifically cleaves β1,4 glycosidic bonds adjacent to D-(1→3) linkages.10 Lichenan subjected to lichenase yields cellulose-like oligosaccharides of 2 to 9 glucose units. Menon et al. found that subjecting lichenan to lichenase and then to βglucosidase results in complete hydrolysis to glucose. These hydrolysates can be fermented using thermotolerant yeast for complete conversion to bioethanol. 4 Thus, lichenan can be used as a decentralized lignocellulosic feedstock for bioethanol production in developing countries where farming is widely distributed.4 Understanding how the composition of lichenan, cellulose and other components of the cell wall affect enzyme accessibility is pivotal in this application, as well as determining optimal conditions for lichenase catalyzed hydrolysis of lichenan. Previous work by the authors on lichenan using a quartz crystal microbalance with dissipation monitoring (QCM-D) addressed lichenan adsorption onto regenerated cellulose (RC) surfaces and competitive adsorption of lichenan onto RC surfaces coated with hemicelluloses such as xyloglucan, arabinoxylan, and glucuronoarabinoxylan. These two studies provided quantitative information about how lichenan interacts with cellulose and hemicelluloses common in plant cell walls of Cetraria islandica. This work addresses the adsorption of lichenan solutions onto RC surfaces and the subsequent degradation of these lichenan layers with the enzyme lichenase. Interactions were studied by varying activity and pH of the enzyme, while keeping the surface, temperature and ionic strength constant. Experimental Materials Lichenan from Cetraria islandica (lot 70901) and lichenase from Bacillus subtilis (lot 60101a) were purchased from Megazyme. Lichenan had a molar mass of 90 kg·mol-1 and lichenase had a mass of 26.750 kg·mol-1.6 Sample solutions of 0.05% lichenan by mass were prepared in sodium acetate buffers (20 mM, pH 5.5). Sample solutions of lichenase were prepared in sodium phosphate buffers (20 mM, pH = 5.5, 6.5 and 7.5). Buffer component were purchased from Aldrich and dissolved in ultrapure water (Millipore Gradient A-10, 18.5 ·cm, < 5 ppb organic impurities). Preparation of Regenerated Cellulose Surfaces Regenerated cellulose surfaces were formed on cleaned sensors (AT-cut quartz crystals covered with 5 nm of chromium and 100 nm of gold, Q-Sense AB). The sensors were cleaned with a UV/ozone ProCleaner (Bioforce Nanoscience,Inc.) for 20 min followed immersion into a heated solution of 10 mL ammonium hydroxide (conc.), 10 mL hydrogen peroxide (30% v/v), and 50 mL of ultrapure water. The regenerated cellulose films were spincoated from a solution of trimethylsilyl cellulose (TMSC) in toluene at 2000 rpm for 60 s. Exposure of the TMSC surface to the vapor of an aqueous solution of hydrochloric acid (10 wt%) for 5 min yielded a smooth surface. 11 Quartz Crystal Microbalance with Dissipation Monitoring (QCM-D) Four sensors coated with regenerated cellulose were placed into the QCM-D flow cells. Adsorption profiles included scaled frequency changes (Δf/n) and a dissipation changes (ΔD) as a function of time from the intrinsic frequency (4.95 MHz for gold-coated quartz crystals) and several odd overtones (n = 3 - 13). First, water was introduced into the flow cells at a rate of 0.200 mL·min-1 at 50 °C (the same rate and temperature was used for all experiments) for several hours until a stable baseline was obtained. Lichenan solution was then introduced into the flow cells and Δf/n and ΔD were recorded as a function of time. The adsorption time for lichenan solution was ~ 1 h until plateaus in Δf/n and ΔD profiles were obtained. Water was reintroduced into the flow cells for an additional ten minutes to remove the loosely bound molecules and determine the amount of irreversibly adsorbed material. Next, phosphate buffer was introduced into the flow cells for ten minutes in preparation for enzyme degradation. Once a stable phosphate buffer baseline was obtained, enzyme solution was introduced for one hour. Finally, phosphate buffer was reintroduced into the enzyme for the removal of reversibly bound material. Experiments were run in triplicate and reported values indicate the average ± one standard deviation. Voigt-based Viscoelastic Modeling For rigid surface layers (D small), the adsorbed surface concentration () is directly proportional to f/n through the Sauerbrey equation.12For soft films (D large), the Sauerbrey equation is invalid. As this was the case for adsorbed lichenen layers, a Voigt-based viscoelastic model was utilized in order to deduce , elasticity, viscosity, and the density of the absorbed layer.13 The modeling assumed the regenerated cellulose layer was an extension of the purely elastic quartz crystal and the surrounding solution was a purely viscous, semi – infinite Newtonian fluid. Atomic Force Microscopy (AFM) Lichenan layers obtained before and after enzyme degradation were dried in a vacuum oven at 50 °C overnight and then imaged with an MFP-3D-Bio atomic force microscope (MFP-3D-BIO, Asylum Research) in tapping mode. AFM images were collected under ambient conditions using a silicon tip (OMCL-AC 160TS, Olympus Corp.). The reported roughnesses are root-mean-square (RMS) values determined from a 2 μm × 2 μm scan area. Results and Discussion Adsorption of Lichenan onto RC surface Irreversible adsorption of lichenan onto RC was observed for adsorption from 0.05% by mass lichenan solutions in sodium acetate buffer (pH = 5.5) at 50 °C. The adsorption resulted in irreversible Δf/n = -55 ± 1 Hz and D = (6.9 ± 0.2) x 10-6. Voigt-based viscoelastic modeling of the lichenan layer irreversibly adsorbed onto RC surfaces using overtones n = 7 - 13 was consistent with a layer having  = 21 ± 1 mg·m-2, a density of 1050 kg·m-3, an elastic shear modulus of (0.32 ± 0.04) x 10-5 N·m-2 and a shear viscosity of (0.98 ± 0.03).The adsorbed lichenen layers had deduced thicknesses of 19 ± 1 nm, lacked large aggregates and were only marginally rougher than the RC surface root-mean-square (rms) roughnesses of ~1.3 vs. ~1.0 nm (Figure 1). RC 1.0 nm Lichenan 1.3 nm Figure 1. 2 m x 2 m AFM images of the RC surface before and after lichenan adsorption. Numbers on the images represent the rms roughnesses. Figure 2 shows f/n and D as a function of time for a lichenan film adsorbed onto RC at pH = 6.5 upon exposure to enzyme at different concentrations. The initial rate of hydrolysis increased with concentration until a limiting value was obtained around 5 U·mL-1. For solutions with concentrations of 0.5 U·mL-1 and higher, f/n and D were comparable after ~ 1 h. The increase in f/n (≈ 40 Hz), was ≈ 15 Hz less than the amount of lichenan that initially adsorbed onto the surface. This observation could mean that hydrolysis was incomplete, some enzyme remained adsorbed on the cellulose, or both. Figure 3 shows a control experiment in which lichenase was directly adsorbed onto a RC suface from a 10 U·mL-1 solution. As seen in Figure 3, some enzyme irreversibly adsorbs onto the RC film, f/n ≈ - 5 Hz after flushing the system with buffer. However, the amount is too small to account for all of the ”missing” f/n in Figure 2. As such, it appears the hydrolysis of lichenan by lichenase is incomplete. Figure 4. QCM-D degradation profiles for the 5th overtone for lichenan layers exposed to 5 U·mL-1 lichenase solutions at different pH. Conclusions th Figure 2. QCM-D degradation profiles from the 5 overtone for lichenan layers exposed to lichenase solutions at pH = 6.5. For almost all lichenase concentrations, nearly complete hydrolysis of the adsorbed lichenan films resulted after about 1 h. Increasing lichenase concentration led to an increase in the initial rate of hydrolysis of lichenan films adsorbed onto cellulose until the concentrations exceeded 5 U·mL-1. Comparable degradation occurred for the entire pH range (5.5 to 7.5) studied. Acknowledgements Figure 3. QCM-D adsorption profiles from the 5th overtone for lichenase adsorbed onto RC surfaces from 10 U·mL-1 solutions at pH = 6.5. In general, enzyme activity is a strong function of pH and temperature. While the effect of temperature was not considered in this study, the effect of pH was. Figure 4 shows f/n and D as a function of time for a lichenan film adsorbed onto RC upon exposure to enzyme at different pH for lichenase concentration of 1 U·mL-1. As seen in Figure 4, the variation in enzyme activity over the pH range of 5.5 to 7.5 is small to insignificant. The optimal pH for this enzyme is 6.5 with an optimal temperature of 60 °C.14 Another important factor influencing the activity of lichenase is the presence of calcium ions. The enzyme possesses a calcium binding site remote from the active site. Other studies have shown that calcium ions are important for the thermal and pH stability of lichenase, though sodium does bind to the same site with a less stable geometry.15 The authors are grateful for the financial support from NSF under Contract DMR-0805179 and the Center for LignoCellulose Structure and Formation, an Energy Frontier Research Center funded by the U.S. Department of Energy, Office of Science, Office of Basic Energy Sciences under Award Number DE-SC0001090. References (1) Heredia, A.; Jiménez, A.; Guillén, R. Composition of plant cell walls. Zeitschrift für Lebensmitteluntersuchung und -Forschung A 1995, 200, 24–31. (2) Scheller, H. V.; Ulvskov, P. Hemicelluloses. Annual Review of Plant Biology 2010, 61, 263–289. (3) Podterob, A. Chemical composition of lichens and their medical applications. Pharmaceutical Chemistry Journal 2008, 42, 582–588. (4) Menon, V.; Divate, R.; Rao, M. Bioethanol production from renewable polymer lichenan using lichenase from an alkalothermophilic Thermomonospora sp. and thermotolerant yeast. Fuel Processing Technology 2011, 92, 401–406. (5) Tosh, S. M.; Brummer, Y.; Wood, P. J.; Wang, Q.; Weisz, J. Evaluation of structure in the formation of gels by structurally diverse (1→3)(1→4)-β-d-glucans from four cereal and one lichen species. Carbohydrate Polymers 2004, 57, 249–259. (6) Staudte, R. G.; Woodward, J. R.; Fincher, G. B.; Stone, B. A. Water-soluble (1→3), (1→4)-β-dglucans from barley (Hordeum vulgare) endosperm. III. Distribution of cellotriosyl and cellotetraosyl residues. Carbohydrate Polymers 1983, 3, 299–312. (7) Tvaroska, I.; Ogawa, K.; Deslandes, Y.; Marchessault, R. H. Crystalline conformation and structure of lichenan and barley β-glucan. Can. J. Chem. 1983, 61, 1608–1616. (8) Lazaridou, A.; Biliaderis, C. G. Molecular aspects of cereal β-glucan functionality: Physical properties, technological applications and physiological effects. Journal of Cereal Science 2007, 46, 101–118. (9) Reijonen, M. WO/2006/125857, December 1, 2006 (10) Malet, C.; Jimenez-Barbero, J.; Bernabe, M.; Brosa, C.; Planas, A. Stereochemical course and structure of the products of the enzymic action of endo-1,3-1,4beta-D-glucan 4-glucanohydrolase from Bacillus licheniformis. Biochem J 1993, 296, 753–758. (11) Kontturi, E.; Thüne, P. C.; Niemantsverdriet, J. W., Cellulose model surfaces - Simplified preparation by spin coating and characterization by X-ray photoelectron spectroscopy, infrared spectroscopy, and atomic force microscopy. Langmuir2003, 19 (14), 5735-5741. (12) Sauerbrey, G., The use of quartz oscillators for weighing thin layers and for microweighing. Z. Phys1959, 155, 206-222. (13) Voinova, M.; Rodahl, M.; Jonson, M.; Kasemo, B., Viscoelastic acoustic response of layered polymer films at fluid-solid interfaces: Continuum mechanics approach. Physica Scripta 1999, 59, 391-396. (14) McCleary, B. V., Lichenase from Bacillus subtilis. In Methods in Enzymology, Willis A. Wood, S. T. K., Ed. Academic Press: 1988; Vol. Volume 160, pp 572575. (15) Keitel, T.; Meldgaard, M.; Heinemann, U. Cation binding to a Bacillus (1,3–1,4)-β-glucanase Geometry, affinity and effect on protein stability. European Journal of Biochemistry 1994, 222, 203–214.