Forestry, Faculty of
DSpace
UBCV
Gourlay, Keith Ian
Doctor of Philosophy - PhD
University of British Columbia
Agricultural and forestry-derived fibres can be converted into fuels and chemicals via a biorefinery. However, the densely-packed fibrillar architecture of lignocellulosic biomass makes the cellulose inherently inaccessible to the enzymes involved in this bioconversion process. This limits the efficiency of enzymatic deconstruction and necessitates relatively high enzyme/protein loadings, which decreases the economic viability of the overall process. It has previously been suggested that the rate-limiting step of cellulose hydrolysis is not the depolymerisation of the carbohydrate chains, but rather the rate at which the enzymes can gain access to the cellulose buried within the biomass. Recently, several proteins such as the Expansins, Swollenin and Loosenin have been shown to disrupt the cellulosic structure without directly depolymerizing the carbohydrates. This protein-induced “amorphogenesis” is thought to occur as a delamination, splitting, peeling, swelling, or decrystallizing of the biomass, thereby enhancing accessibility of the entrenched carbohydrates to the depolymerizing enzymes. However, a key challenge when studying these amorphogenesis-inducing proteins involves quantifying their disruptive effects. While depolymerizing enzymes can be readily quantified by measuring the amount of liberated soluble sugars, amorphogenesis-inducing proteins are thought to promote a variety of disruptive effects without releasing soluble products. As the undefined nature of the amorphogenesis end product makes quantification challenging, one of the initial goals of the work was to refine/develop techniques to better quantify amorphogenesis. Two distinct carbohydrate binding modules (CBMs), one of which preferentially binds to crystalline cellulose and the other to amorphous cellulose were used to track changes in cellulose accessibility and surface morphology. When various substrates were treated with the amorphogenesis-inducing protein, Swollenin, CBM adsorption revealed that Swollenin promoted the dispersal and disruption of the more amorphous regions of biomass, increasing the access of the depolymerizing enzymes to the cellulose component. Subsequent work involving the fluorescent tagging of these CBMs and confocal microscopy further suggested that Swollenin was targeting the less-ordered regions of the cellulosic substrate. When Swollenin was assessed for its ability to disrupt an industrially-relevant substrate, steam pretreated corn stover, it primarily targeted amorphous regions where it synergised strongly with xylanases (~300%), promoting the release of hemicellulosic oligomers.
The Role of Amorphogenesis in the Enzymatic Deconstruction of Lignocellulosic Biomass by Keith Ian Gourlay MRes, The Imperial College of Science, Technology and Medicine, 2009 BSc, Queen’s University, 2008 A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY in THE FACULTY OF GRADUATE AND POSTDOCTORAL STUDIES (FORESTRY) THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver) OCTOBER 2014 © Keith Ian Gourlay, 2014 ii Abstract Agricultural and forestry-derived fibres can be converted into fuels and chemicals via a biorefinery. However, the densely-packed fibrillar architecture of lignocellulosic biomass makes the cellulose inherently inaccessible to the enzymes involved in this bioconversion process. This limits the efficiency of enzymatic deconstruction and necessitates relatively high enzyme/protein loadings, which decreases the economic viability of the overall process. It has previously been suggested that the rate-limiting step of cellulose hydrolysis is not the depolymerisation of the carbohydrate chains, but rather the rate at which the enzymes can gain access to the cellulose buried within the biomass. Recently, several proteins such as the Expansins, Swollenin and Loosenin have been shown to disrupt the cellulosic structure without directly depolymerizing the carbohydrates. This protein-induced “amorphogenesis” is thought to occur as a delamination, splitting, peeling, swelling, or decrystallizing of the biomass, thereby enhancing accessibility of the entrenched carbohydrates to the depolymerizing enzymes. However, a key challenge when studying these amorphogenesis-inducing proteins involves quantifying their disruptive effects. While depolymerizing enzymes can be readily quantified by measuring the amount of liberated soluble sugars, amorphogenesis-inducing proteins are thought to promote a variety of disruptive effects without releasing soluble products. As the undefined nature of the amorphogenesis end product makes quantification challenging, one of the initial goals of the work was to refine/develop techniques to better quantify amorphogenesis. Two distinct carbohydrate binding modules (CBMs), one of which preferentially binds to crystalline cellulose and the other to amorphous cellulose were used to track changes in cellulose accessibility and surface morphology. When various substrates were treated with the amorphogenesis-inducing protein, Swollenin, CBM adsorption revealed that Swollenin promoted the dispersal and disruption of the more iii amorphous regions of biomass, increasing the access of the depolymerizing enzymes to the cellulose component. Subsequent work involving the fluorescent tagging of these CBMs and confocal microscopy further suggested that Swollenin was targeting the less-ordered regions of the cellulosic substrate. When Swollenin was assessed for its ability to disrupt an industrially-relevant substrate, steam pretreated corn stover, it primarily targeted amorphous regions where it synergised strongly with xylanases (~300%), promoting the release of hemicellulosic oligomers. iv Preface A version of Section 3.2 was published as Gourlay, K., Arantes, V., and J. Saddler, 2012. Use of substructure-specific carbohydrate binding modules to track changes in cellulose accessibility and surface morphology during the amorphogenesis step of enzymatic hydrolysis. Biotechnology for Biofuels 5:51. I performed the experimental work and drafted the manuscript. Valdeir Arantes and Jack Saddler contributed to the planning, data interpretation and the writing of the manuscript. Data from Section 3.3 was incorporated into two publications, including Gourlay, K., Hu, J., Arantes, V., and J. Saddler (2014) Monitoring changes in cellulose fibre surface morphology and fragmentation during treatment with cellulases and Swollenin. In preparation (submitted). In this work, I performed the experimental work and drafted the manuscript. Jinguang Hu helped with the experimental work, while Valdeir Arantes and Jack Saddler contributed to the planning, data interpretation and writing of the manuscript. An additional publication from this section was Arantes, V., Gourlay, K., and J. Saddler (2014). The Enzymatic Hydrolysis of Pretreated Pulp Fibres Predominantly Involves “Peeling/Eroding” Modes of Action. (Biotechnology for Biofuels 2014, 7:87). In this work, experiments and manuscript drafting were carried out by myself and Valdeir Arantes. Jack Saddler contributed to the planning, data interpretation and writing of the manuscript. A version of Section 3.4 was published as Gourlay, K., Hu, J., Arantes, V., Andberg, M., Saloheimo, M., Pentillä, M., and J. Saddler (2013). Swollenin aids in the amorphogenesis step during the enzymatic hydrolysis of pretreated biomass. Bioresource Technology 142:498-593. I performed the experimental work and drafted the manuscript. Jinguang Hu provided the purified hydrolases. Martina Andberg, Markku Saloheimo, Merja Pentillä provided the Swollenin enzyme. Valdeir Arantes and Jack Saddler contributed to the planning, data interpretation and the writing of the manuscript. v Additional publications resulting from research carried out during this thesis project include: • Hu, J., Arantes, V., Pribowo, A., Gourlay, K., and J. Saddler (2014) Substrate factors that influence the synergistic interaction of AA9 and cellulases during the enzymatic hydrolysis of biomass. Energy & Environmental Science. 7, 2308-2315 • Chandra, R., Gourlay, K., Chung, P., and J. Saddler (2014) Two-stage, acid-catalyzed steam pretreatment of wood results in good hemicellulose recovery and effective enzymatic hydrolysis of the cellulosic component. In preparation, submitted. • Hu, J., Gourlay, K., Arantes, V., Pribowo, A, and J. Saddler (2014) Synergistic cooperation between cellulase components during lignocellulose deconstruction. In preparation. • Hu, J., Arantes, V., Gourlay, K., Pribowo, A., and J. Saddler (2014) Can synergistic cooperation between cellulase and accessory enzymes enhance the high solid loading hydrolysis of pretreated lignocellulose? In preparation. vi Table of contents Abstract .................................................................................................................................................. ii Preface ................................................................................................................................................... iv Table of contents.................................................................................................................................... vi List of tables ............................................................................................................................................ x List of figures .......................................................................................................................................... xi List of abbreviations .............................................................................................................................. xvi Acknowledgements ................................................................................................................................xx 1. Introduction........................................................................................................................................ 1 1.1 Structural features of lignocellulosic biomass influencing enzymatic hydrolyzability ................. 7 1.1.1 A brief history on the structure of cellulose .......................................................................... 7 1.1.2 The multilevel organization of lignocellulose ...................................................................... 12 1.1.3 Crystallinity ........................................................................................................................ 13 1.1.4 Hemicelluloses ................................................................................................................... 18 1.1.5 Lignin ................................................................................................................................. 21 1.1.6 Cellulose accessibility ......................................................................................................... 25 1.1.7 Fibre dislocations ............................................................................................................... 27 1.1.8 Effects of pretreatments on the structure of lignocellulosic biomass .................................. 29 1.2 Enzymatic deconstruction of lignocellulosic substrates ........................................................... 32 1.2.1 Enzymes involved in lignocellulose deconstruction ............................................................. 33 1.2.1.1 Hydrolytic cellulases ................................................................................................... 35 1.2.1.2 Oxidative and accessory enzymes ............................................................................... 37 1.2.1.3 Amorphogenesis-inducing proteins............................................................................. 38 1.3 Evidence that amorphogenesis does occur ............................................................................. 40 1.3.1 Macroscopic disruption/fragmentation .............................................................................. 40 1.3.2 Microscopic/ultrastructural disruption ............................................................................... 43 1.3.3 Nanoscopic disruption ........................................................................................................ 44 1.4 Focus on Swollenin ................................................................................................................. 48 1.5 Enhancement of biomass deconstruction by amorphogenesis-inducing proteins.................... 49 1.6 Current techniques used for measuring amorphogenesis ....................................................... 53 1.7 Techniques for measuring accessibility with potential for quantifying amorphogenesis .......... 56 1.7.1 Water retention value ....................................................................................................... 57 1.7.2 Fibre quality analyzer (FQA) ............................................................................................... 58 vii 1.7.3 Nitrogen adsorption and mercury porosimetry .................................................................. 58 1.7.4 Solute exclusion ................................................................................................................. 59 1.7.5 Differential scanning calorimetry and time-domain nuclear magnetic resonance ............... 60 1.7.6 Particle scattering .............................................................................................................. 63 1.7.7 Simons’ stain ..................................................................................................................... 63 1.7.8 Cellulase adsorption .......................................................................................................... 65 1.7.9 CBM adsorption ................................................................................................................. 66 1.7.10 Summary of techniques ..................................................................................................... 71 1.8 Thesis overview and research approach ................................................................................. 71 1.8.1 Summary ............................................................................................................................ 71 1.8.2 Research approach ............................................................................................................. 73 2. Materials and methods .................................................................................................................. 77 2.1 Proteins and enzymes ............................................................................................................ 77 2.2 Biomass preparation .............................................................................................................. 78 2.3 Determining pretreated biomass composition........................................................................ 79 2.4 Enzymatic hydrolysis and Swollenin treatments ..................................................................... 80 2.5 Accessibility measurements ................................................................................................... 82 2.5.1 Simons’ staining ................................................................................................................. 82 2.5.2 FQA and settlability ............................................................................................................ 83 2.5.3 Cellulase adsorption ........................................................................................................... 83 2.5.4 Nitrogen adsorption ........................................................................................................... 84 2.5.5 CBM adsorption ................................................................................................................. 84 2.6 CBM adsorption for Scatchard plots ....................................................................................... 85 2.7 SEM imaging .......................................................................................................................... 86 2.8 X-ray diffraction ..................................................................................................................... 86 2.9 Confocal microscopy .............................................................................................................. 86 2.10 Monomer and oligomer analysis after Swollenin treatment ................................................... 87 2.10.1 High performance liquid chromatography (HPLC) ........................................................... 87 2.10.2 Polyacrylamide carbohydrate electrophoresis (PACE) ..................................................... 87 3. Results and discussion ................................................................................................................... 89 3.1 The role of accessibility in enzymatic hydrolysis ..................................................................... 89 3.1.1 Background ........................................................................................................................ 89 3.1.2 Substrate preparation ........................................................................................................ 91 3.1.3 Correlating accessibility with enzymatic hydrolyzability ...................................................... 91 3.1.4 CBM adsorption as a tool for quantifying accessibility ...................................................... 102 viii 3.1.5 Conclusions ...................................................................................................................... 106 3.2 Development and application of a new technique for quantifying amorphogenesis ............. 107 3.2.1 Background ...................................................................................................................... 107 3.2.2 Measuring cellulose accessibility ...................................................................................... 112 3.2.2.1 Adsorption of Simons’ stain to model cellulosic substrates ....................................... 112 3.2.2.2 Adsorption of CBM2a and CBM44 to dissolving pulp ................................................. 115 3.2.2.3 Adsorption of CBMs to model cellulosic substrates ................................................... 117 3.2.3 Quantification of acid-induced cellulose disruption .......................................................... 120 3.2.3.1 CBM adsorption ........................................................................................................ 120 3.2.3.2 Relationship between initial hydrolysis rate and CBM adsorption ............................. 123 3.2.4 Tracking changes in cellulose morphology over the course of hydrolysis .......................... 125 3.2.5 Quantification of Swollenin-induced changes in cellulose accessibility .............................. 128 3.2.6 Conclusions ...................................................................................................................... 133 3.3 Amorphogenesis and fibre fragmentation ............................................................................ 133 3.3.1 Background ...................................................................................................................... 133 3.3.2 Quantifying macroscopic fibre properties during enzymatic deconstruction ..................... 139 3.3.2.1 Fibre length .............................................................................................................. 139 3.3.2.2 Fibre width ............................................................................................................... 141 3.3.3 Quantifying Swollenin-induced macroscopic manifestations of amorphogenesis on lignocellulosic biomass ..................................................................................................... 144 3.3.4 Quantifying Swollenin-induced macroscopic manifestations of amorphogenesis on a model cellulosic substrate ........................................................................................................... 145 3.3.5 Conclusions ...................................................................................................................... 154 3.4 The effects of Swollenin on an industrially-relevant biomass substrate, steam pretreated corn stover ..................................................................................................................................... 155 3.4.1 Background ...................................................................................................................... 155 3.4.2 Macroscopic effects of Swollenin on steam pretreated corn stover .................................. 156 3.4.3 Microscopic effects of Swollenin on steam pretreated corn stover ................................... 158 3.4.3.1 Small particle release ................................................................................................ 158 3.4.3.2 Quantification of oligomer release by Swollenin ....................................................... 159 3.4.3.2.1 High performance liquid chromatography analysis ............................................... 159 3.4.3.2.2 Polyacrylamide carbohydrate electrophoresis analysis ......................................... 161 3.4.3.3 CBM adsorption to Swollenin-treated SPCS............................................................... 163 3.4.4 Synergism between Swollenin and hydrolytic cellulase and xylanase monocomponents... 164 3.4.5 Conclusions ...................................................................................................................... 168 4. Conclusions and future work .................................................................................................... 169 4.1 Conclusions .......................................................................................................................... 169 4.2 Future work ......................................................................................................................... 171 ix 4.2.1 Screening for effective combinations of biomass, amorphogenesis-inducing proteins, and hydrolytic/oxidative enzymes ........................................................................................... 171 4.2.2 Broader application of tagged CBMs as molecular probes ................................................ 172 4.2.3 Pulp and paper applications ............................................................................................. 172 References .......................................................................................................................................... 177 x List of tables Table 1: Amorphogenesis-inducing proteins identified to date, along with their putative function and level of disruption. ................................................................................................................................ 46 Table 2: Pretreatment conditions and substrate composition after pretreatment for a range of substrates ............................................................................................................................................. 92 Table 3: Amount of protein required to achieve 70% conversion after 72 hours at 2% consistency. ....... 93 Table 4: Pretreatment conditions and substrate compositions for the biomass samples used in the CBM adsorption experiments. ..................................................................................................................... 103 Table 5: The degree of synergism (DS) between various purified hydrolytic enzymes and Swollenin on SPCS. Samples were run in triplicate and σDS represents one standard deviation. ............................... 166 xi List of figures Figure 1: Model of protein-induced amorphogenesis of biomass occurring at the nanoscopic (A1), microscopic (B1), and macroscopic (C1) levels. Manifestations include the nanoscopic pitting of microfibrils (A2), microscopic dispersion, swelling, weakening and loosening of cell walls (B2), and the dispersion and disaggregation of macroscopic associations of fibre cells (C2). ........................................ 47 Figure 2: Derivatization process of sugars or oligomers with ANTS in the presence of NaBH3CN. ........... 88 Figure 3: Representative conversion curve of glucan conversion over time for a typical enzymatic deconstruction reaction. ....................................................................................................................... 93 Figure 4: Glucan content of pretreated biomass samples relative to Simons’ Stain dye ratio. Both the steam- and organosolv-pretreated corn fibre samples (SPCF, OPCF) were excluded from the correlation coefficient calculation (adapted from Arantes and Saddler, 2011). ........................................................ 95 Figure 5: Minimum enzyme loading plotted against Simons’ Stain dye ratio. The corn fibre samples were excluded from the regression analysis (adapted from Arantes and Saddler, 2011). ................................ 96 Figure 6: Minimum enzyme loading required for 70% hydrolysis plotted against the average fibre length. The corn fibre samples were excluded from the regression analysis (adapted from Arantes and Saddler, 2011). .................................................................................................................................................... 98 Figure 7: Minimum enzyme loading required to achieve 70% conversion plotted against nanoscopic accessibility as measured by Nitrogen BET. ............................................................................................ 99 Figure 8: Minimum protein loading required for 70% conversion is plotted against the maximum protein adsorption for a range of pretreated biomass samples. Correlation coefficients were calculated for samples pretreated with either steam or organosolv pretreatment (adapted from Arantes and Saddler, 2011). .................................................................................................................................................. 100 Figure 9: Adsorption of CBM2a and CBM44 to steam-pretreated biomass samples. Samples were run in triplicate and error bars represent one standard deviation from the mean. ......................................... 104 Figure 10: Conversion of three steam pretreated samples plotted against the amount of bound CBM2a (squares), CBM44 (triangles) and the sum of both CBMs (diamonds). .................................................. 104 Figure 11: Representative crystal structures of a Type A CBM (Left) and a Type B CBM (Right). The CBMs illustrated here are the Family 1 CBM from T. reesei Cel7A (Left) (PDB code 1CBH; (Kraulis et al., 1989), and the Family 4 CBM from C. fimi endo-1,4-glucanase C (Right) (PDB code 1GU3; (Boraston et al., 2002b). Reproduced from (Guillén et al., 2010). .................................................................................. 109 xii Figure 12: Adsorption of Simons’ Stain DO dye to a range of model cellulosic substrates. The least ordered substrates (PASC and Cellulose II) were found to adsorb more dye than other substrates. CNC and Cellulose III adsorbed the least dye. PASC: Phosphoric Acid Swollen Cellulose, DSP: Dissolving Pulp, CNC: Cellulose Nanocrystals. ............................................................................................................... 113 Figure 13: Percent conversion after 48 hours for a range of model cellulosic substrates plotted against adsorption of Simons’ Stain DO dye. In general, the more disordered celluloses adsorbed more dye and hydrolysed more readily. The Cellulose III data point was considered an outlier and was excluded from the regression analysis. PASC: Phosphoric Acid Swollen Cellulose, DSP: Dissolving Pulp, CNC: Cellulose Nanocrystals. ....................................................................................................................................... 115 Figure 14: Scatchard plots of CBM2a and CBM44 bound to dissolving pulp. Best fit lines were calculated for the linear sections of the plots only. Emax values represent the maximum amount of CBM adsorbed at equilibrium per mg of cellulose........................................................................................................ 116 Figure 15: Adsorption of CBMs to a range of model cellulosic substrates. Samples were run in triplicate and error bars represent one standard deviation from the mean. PASC: Phosphoric Acid Swollen Cellulose, DSP: Dissolving Pulp, CNC: Cellulose Nanocrystals................................................................ 117 Figure 16: Ratio of CBM2a to CBM44 adsorbed to a range of model cellulosic substrates. Samples were run in triplicate and error bars represent one standard deviation from the mean. PASC: Phosphoric Acid Swollen Cellulose, DSP: Dissolving Pulp, CNC: Cellulose Nanocrystals................................................... 118 Figure 17: Percent conversion after 48 hours of a range of model cellulosic substrates plotted against CBM adsorption. Adsorption of CBM2a is represented by diamonds while CBM44 is represented by squares. Filled shapes represent those included in the regression analysis, while empty shapes represent outliers. Samples were run in triplicate and error bars represent one standard deviation from the mean............................................................................................................................................................. 119 Figure 18: Scanning electron microscopy of cotton fibres after treatment with a range of o-phosphoric acid concentrations. All images were taken at x1200 magnification and each image depicts a representative fibre for the indicated acid concentration. ................................................................... 121 Figure 19: Adsorption of crystalline cellulose-binding CBM2a and amorphous cellulose-binding CBM44 to cotton fibres treated with a range of o-phosphoric acid concentrations. Experiments were run in triplicate and error bars represent one standard deviation from the mean. ......................................... 122 Figure 20: Initial hydrolysis rate vs CBM adsorption. Initial hydrolysis rate (calculated after 30 minutes of hydrolysis) of acid-disrupted cotton fibres increased with increasing CBM adsorption. Each data point represents a cotton fibre sample treated with a different concentration of o-phosphoric acid and xiii hydrolyzed with the same enzyme loading. Experiments were run in triplicate and error bars represent one standard deviation from the mean................................................................................................ 125 Figure 21: Adsorption of CBM2a (squares, solid line) and CBM44 (triangles, dashed line) over the course of a 6 hour hydrolysis, plotted alongside glucose release (diamonds, dotted line). A rapid drop in the amount of accessible amorphous cellulose occurred within the first 5-15 minutes of hydrolysis. ......... 126 Figure 22: Crystallinity index as determined by XRD over the course of enzymatic hydrolysis. ............. 127 Figure 23: Adsorption of CBMs to Swollenin-treated cotton fibres. Swollenin promoted an increase in binding of CBM44 and, particularly, CBM2a. A BSA negative protein control was found to have no significant effect on the extent of binding of either CBM. At least three replicates were performed for each sample and error bars represent one standard deviation from the mean. ................................... 129 Figure 24: Effect of Swollenin on mercerized cotton fibres imaged by SEM. The surface of the control fibre (Left) appears roughened due to the mercerization treatment. The rough features on the surface of the mercerized cotton fibres appear to have been sloughed off by the action of Swollenin (Right). Images are of representative fibres for the indicated treatment. ..................................................................... 130 Figure 25: Representation of quantifying specific manifestations of amorphogenesis using CBM adsorption. In the example illustrated here, the untreated cellulose (top) binds approximately equal amounts of CBM2a and CBM44. After protein-induced amorphogenesis, the possible manifestations of amorphogenesis can be separated between sloughing/splitting/delaminating/peeling effects (Left) and surface decrystallization effects (Right). If the ratio of bound CBM2a:CBM44 increases or remains unchanged during amorphogenesis, this suggests the splitting/delaminating/peeling manifestation, whereas if this ratio decreases, a manifestation of amorphogenesis involving the direct surface decrystalization is occurring. In the case of Swollenin, this ratio increased, consistent with the scenario on the left representing a sloughing/splitting/delaminating/peeling effect. Cr = Crystalline-binding, Am = Amorphous-binding. ............................................................................................................................ 132 Figure 26: Enzymatic hydrolysis profiles of pretreated poplar and lodgepole pine. No correlation was observed between initial fibre length and enzymatic hydrolyzability. SPP180: Steam Pretreated Poplar pretreated at 180°C, SPP200: Steam Pretreated Poplar pretreated at 200°C, OPLP: Organosolv Pretreated Lodepole Pine. ................................................................................................................... 140 Figure 27: Average fibre lengths of fibres from two pretreated hardwood and one pretreated softwood substrates over the course of enzymatic hydrolysis. SPP180: Steam Pretreated Poplar pretreated at 180°C, SPP200: Steam Pretreated Poplar pretreated at 200°C, OPLP: Organosolv Pretreated Lodepole Pine. .................................................................................................................................................... 140 Figure 28: Average fibre width (midpoint of each bar) and standard deviation of the width (height of each bar) for SPP180, SPP200 and OPLP over the course of hydrolysis. SPP180: Steam Pretreated Poplar xiv pretreated at 180°C, SPP200: Steam Pretreated Poplar pretreated at 200°C, OPLP: Organosolv Pretreated Lodepole Pine. ................................................................................................................... 142 Figure 29: Representative SEM micrographs of SPP200 samples treated with either buffer (A,B) or cellulase (C,D). Fibre peeling can be observed at both low (C) and high (D) magnifications. ................. 143 Figure 30: Average fibre width of pretreated hardwood, softwood and dissolving pulp substrates after treatment with either a BSA control or Swollenin at 10µg protein/mg substrate. All samples were run in triplicate and error bars represent one standard deviation from the mean. ......................................... 144 Figure 31: Average fibre length of various pretreated lignocellulosic pulps and dissolving pulp after treatment with either buffer or Swollenin at 10µg protein/mg substrate. DSP: Dissolving Pulp, SPCS: Steam Pretreated Corn Stover, OPCS: Organosolv Pretreated Corn Stover, SPP: Steam Pretreated Poplar, OPP: Organosolv Pretreated Poplar, SPLP: Steam Pretreated Lodgepole Pine, OPLP: Organosolv Pretreated Lodgepole Pine. All samples were run in triplicate and error bars represent one standard deviation from the mean. .................................................................................................................... 145 Figure 32: Average fibre length of dissolving pulp fibres treated overnight at 50°C with 10µg purified enzyme per mg substrate. BSA: Bovine Serum Albumin, Xyn: Xylanase GH10, CBH: Cel7A, EG: Cel5A, Swo: Swollenin. ................................................................................................................................... 146 Figure 33: Sugar release and average fibre length during enzymatic hydrolysis of dissolving pulp using 15mg enzyme/g glucan. All reactions were performed in triplicate and error bars represent one standard deviation from the mean. .................................................................................................................... 148 Figure 34: ‘Settlability’ of dissolving pulp fibres over the course of hydrolysis at 15mg enzyme/g glucan. A dramatic increase in settlability is observed within the first 5 minutes, followed by a subsequent step-up in settlability between 60 and 90 minutes. Fibres were stained with Direct Orange dye and washed thoroughly prior to settling overnight in order to enhance the visibility of the fibres. .......................... 149 Figure 35: A representative dissolving pulp fibre imaged without excitation light (A), with excitation of CBM2a-AMCA-X (B), and with excitation of CBM44-OG (C). The colored pixels from B and C were extracted, combined, and overlaid onto the original fibre image to produce figure D. The colored pixels were then enhanced (E). In general, CBM2a (blue) localized to well-ordered regions of the fibre surface while CBM44 (green) was predominantly bound within fibre dislocations. A full description of the enhancement process is provided in the preceding paragraphs. .......................................................... 150 Figure 36: Fibre length of steam-pretreated (SPCS) and organosolv pretreated (OPCS) corn stover after treatment with either a buffer control or Swollenin............................................................................. 157 Figure 37: Turbidity of the supernatants of Swollenin and control-treated SPCS. Swollenin appears to decrease the turbidity of the supernatants. The slight increase in turbidity in the presence of the BSA xv protein control likely corresponds to protein aggregation, also observed in the sample containing just BSA and buffer in the absence of SPCS. BSA: Bovine Serum Albumin, SPCS: Steam Pretreated Corn Stover.................................................................................................................................................. 158 Figure 38: Sugars detected after incubation of SPCS with either Swollenin or a BSA control. (A) Oligomeric and monomeric glucose and xylose release. (B) Amount of oligomeric arabinose, galactose and mannose detected after incubation with Swollenin. These sugars were not detected in monomeric form, and oligomers were undetectable in the BSA-treated samples. (C) Percent release of cellulosic and hemicellulosic sugars in a soluble form after incubation with Swollenin relative to the original glucan and total hemicellulose content of the pretreated substrate. Values are relative to a BSA control. All reactions were run in triplicate and error bars represent one standard deviation from the mean. ....... 160 Figure 39: Amount of monomeric and oligomeric glucose and xylose detected after incubation of Swollenin with SPCS between 20°C and 50°C. Total sugar represents the sum of all oligomeric and monomeric glucose and xylose. Values are relative to a BSA control, reactions were run in triplicate, and error bars represent one standard deviation from the mean. .............................................................. 160 Figure 40: PACE analysis of released carbohydrates. (A) Carbohydrates released after incubation of Swollenin and a BSA control with SPCS. Standards are labeled; HMW represents the higher molecular weight fraction. (B) Densitometry analysis and peak selection. (C) Quantification of the relative increase in sugars released by Swollenin compared to the control. Values represent the percent increase in peak area for each sugar. ............................................................................................................................. 162 Figure 41: Change in CBM adsorption to SPCS after Swollenin treatment. Absolute change in CBM binding is shown on the left, while percent change is shown on the right. Samples were run in triplicate and error bars represent one standard deviation from the mean. ....................................................... 164 Figure 42: The amount of glucose (A) and xylose (B) released from SPCS after incubation of various purified hydrolytic enzymes with either a BSA control or Swollenin. Reactions were run in triplicate and error bars represent one standard deviation from the mean. .............................................................. 165 xvi List of abbreviations σ error (standard deviation) µL microliter AFEX ammonia fibre expansion AFM atomic force microscopy AIL acid-insoluble lignin Am amorphous AMCA-X (6-((7-Amino-4-Methylcoumarin-3-Acetyl)amino)Hexanoic Acid ANOVA analysis of variance ANTS 8-aminonaphthalene-1,3,6-trisulfonic acid Ara arabinose ASL acid-soluble lignin BDL below detectable level BET Brunauer–Emmett–Teller BSA bovine serum albumin CBH cellobiohydrolase CBM carbohydrate binding module CBU cellobiohydrolase units CD catalytic domain CFP cyan fluorescent protein CNC cellulose nanocrystals Cr crystalline CrI crystallinity index DB direct blue dye DEAE diethylaminoethanol xvii DMSO dimethyl sulfoxide DNS dinitrosalicylic DO direct orange dye DS degree of synergism DSC differential scanning calorimetry DSP dissolving pulp EO ethanol organosolv EtOH ethanol FITC fluorescein isothiocyanate FFP fibril forming protein FNIII fibronectin type III FPU filter paper units FQA fibre quality analyzer FTIR Fourier-transform infrared g gram G guaiacyl Gal galactose Glu glucose H2SO4 sulfuric acid H hydroxyphenyl HCl hydrochloric acid HMW high molecular weight HPAEC-PAD High Performance Anion Exchange Chromatography with Pulsed Amperometric Detection HPLC high-performance liquid chromatography IPTG Isopropyl-β-D-1-thiogalactopyranoside xviii kDa kilo-Dalton kg kilogram L litre LCC lignin carbohydrate complex LPMO lytic polysaccharide mono-oxygenase m meter Man mannose mg milligram mL millilitre mM millimole per litre mmol millimole M mole per litre MW molecular weight Na2CO3 sodium carbonate NaCNBH3 sodium cyanoborohydride NaOH sodium hydroxide nm nanometer NMR nuclear magnetic resonance OD optical density OG Oregon Green 514 OPCF organosolv pretreated corn fibre OPCS organosolv pretreated corn stover OPLP organosolve pretreated lodgepole pine PACE polyacrylamide carbohydrate electrophoresis PASC phosphoric acid swollen cellulose xix PDB protein data bank s second(s) S syringyl SANS small angle neutron scattering SAXS small angle x-ray scattering SEM scanning electron microscopy SS Simon’s Stain SDS-PAGE sodium dodecyl sulfate polyacrylamide gel electrophoresis SO2 sulphur dioxide SPCF steam pretreated corn fibre SPCS steam pretreated corn stover SPDF steam pretreated douglas fir SPLP steam pretreated lodgepole pine SPP steam pretreated poplar t time TD-NMR time domain nuclear magnetic resonance USD United States dollars v/v volume/volume w/v weight/volume w/w weight/weight WAXD wide angle x-ray diffraction WRV water retention value XRD x-ray diffraction Xyl xylose Xyn xylanase xx Acknowledgements I would like to express my sincere thanks to my supervisor, Dr. Jack Saddler, for his mentorship, support, and guidance throughout my PhD studies, as well as my committee members Dr. Doug Kilburn and Dr. Paul McFarlane for their invaluable advice and input during this research. Special thanks also go to Dr. Valdeir Arantes for generously contributing many hours of his time and providing me with guidance and support. Dr. Jinguang Hu and Dr. Amadeus Pribowo were excellent collaborators during this research, and deserve recognition for the extensive support and friendship that they provided. I would also like to thank all members of the Forest Products Biotechnology/Bioenergy group for their ongoing help and for providing an enjoyable work environment, as well as the UBC 4-year fellowship program for funding this research. And last but absolutely not least, the years spent carrying out this research would have been far less enjoyable without the loving support of my family and the fantastic group of friends that have provided me with support and kept me grounded and loving life throughout the course of my PhD studies. You are all amazing and have made the past 5 years fly by in a blur of long days in the lab and long weekends in the sun. xxi “When you cannot measure it, your knowledge is of a meagre and unsatisfactory kind” Lord William Thomson Kelvin (1824-1907) 1 1. Introduction Global fossil fuel reserves are a finite resource. As the easily extracted portions of this resource become scarcer, the world’s focus has been shifting to reserves located in increasingly hard-to-reach places. Extraction from these sources, such as deep sea reserves in the arctic or from within the Canadian tar sands, is energy intensive and has negative environmental consequences (Charpentier et al., 2009; Kelly et al., 2009; Rooney et al., 2012). These drivers have led governments around the world to pursue the production of fuels and chemicals from renewable, sustainable resources. The production of corn and sugarcane ethanol in the USA and Brazil, respectively, has supplemented fossil fuel supply (Chum et al., 2014). However, there is some concern that corn ethanol provides only limited energy savings and may impact food supply (Hill et al., 2006). For these reasons, the world’s focus has been shifting towards developing advanced biorefineries, which make use of agro-industrial or forestry-derived waste residues for conversion to fuels and chemicals (Brown and Brown, 2013; Cheng and Timilsina, 2011). Typically, the production of biofuels via a biochemical pathway involves first breaking down the biomass into monomeric sugars, then fermenting these sugars to produce the desired fuel or chemical. Biomass historically used for producing biofuels through the biochemical pathway has included sugarcane and corn. In these feedstocks, the sugars are relatively easily extracted. For example, with sugarcane, the sugars simply need to be extracted from the cane before being fermented. In crops such as corn or wheat, the sugars are present as starch, a simple, relatively unstructured, homogenous polymer used as temporary energy storage by the plants. The starch chains are predominantly composed of glucose molecules linked via α-1,4 linkages, which induce a slight twist along the length of the chain. The twist at each α-1,4 bond induces the formation of a helix, which, combined with the significant branching found in starch polymers, results in a relatively disordered and loosely packed 2 tertiary structure of starch (French, 1973; Warren, 1996). This description is somewhat simplistic, as starch in nature can be found in a number of ‘resistant starch’ structures, including physically inaccessible starch (RS1), granular starch (RS2), retrograded starch (RS3), and starch which contains additional crosslinking (RS4) (Sajilata et al., 2006). Despite these resistant forms, starch overall is still much more amorphous and accessible than cellulose. This accessibility of starch enables amylase enzymes (which hydrolyze α-1,4 linkages, as well as other linkages making up the starch structure) to easily access the sugar chains and break them down (Naik et al., 2010). Thanks to this ease of access, starch hydrolysis requires relatively low amounts of enzyme to break the starch chains down to monomeric sugars for subsequent fermentation to ethanol. Thus, so called ‘first generation’, or what the IEA has termed ‘conventional’ biofuels, are in many cases, such as in Brazil and some regions within the US, cost competitive with gasoline (Demirbas, 2011; Goldemberg, 2007). Unlike sugar and starch crops, the sugars found within biomass feedstocks used for producing advanced biofuels are locked away within a complex, heterogeneous lignocellulosic matrix. Plants have evolved to polymerize sugars into starch as an energy storage polymer for subsequent rapid depolymerisation back to monomeric sugars. Contrarily, the lignocellulosic matrix making up the structural material of plants has evolved over billions of years to maximize strength while resisting microbial attack. Thus the lignocellulosic fraction of plant biomass is far more difficult to break down into its component sugars than starch. Lignocellulosic biomass has three main components: cellulose, hemicelluloses and lignin (Sjöström, 1993). Cellulose is composed of β-1,4-linked glucose molecules, which unlike the α-1,4 linkages found in starch, form linear, untwisted, unbranched chains. These chains are stacked together into densely-packed, crystalline structures which greatly limit the accessibility of the cellulose to the enzymes involved in the biochemical deconstruction of biomass (the biochemical versus 3 thermochemical pathways are discussed below) (Arantes and Saddler, 2010). Additionally, these crystalline bundles are wrapped in a complex matrix of hemicelluloses, which are branched, heterogeneous polymers made up of a variety of sugars linked via a diverse range of bonds. Finally, the cellulose and hemicellulose fraction is encased within a crosslinked matrix of lignin, a heterogeneous phenolic macromolecule (Somerville et al., 2004). A more detailed discussion on the structure of lignocellulosic biomass is found in Section 1.1. To summarize, producing conventional biofuels from sugar or starch crops is relatively easy. However, producing advanced biofuels from the lignocellulosic portion of biomass is more difficult, and requires additional process steps, higher enzyme loadings, and a greater number of enzyme activities. The conversion of lignocellulosic materials to fuels and chemicals can be achieved through either thermochemical (i.e. gasification or pyrolysis) or biochemical (i.e. enzymatic/microbial) pathways. There are a number of pros and cons associated with each of these pathways (Foust et al., 2009; Wright and Brown, 2007). However, the biochemical route appears to be proceeding more rapidly toward commercialization (Janssen et al., 2013). This could be due, in part, to the fact that this pathway realizes economic viability at smaller scales (Stephen et al., 2010), which is desirable for biomass to biofuel facilities due to the low energy density of the biomass, which makes transporting large volumes of biomass across long distances particularly costly (Shabani et al., 2013; Stephen et al., 2010). In the biochemical conversion pathway, the cost of the enzymes required for the hydrolysis step of biomass processing is thought to be the third largest contributor to the cost of establishing a cellulosic biofuels facility, with only capital cost and feedstock cost contributing more (Isola, 2013). The development of more efficient protein production techniques has successfully brought down the price of producing cellulase mixtures to about $10.14/kg protein (Klein-Marcuschamer et al., 2012), which is on par with the $10.40/kg production costs of starch-degrading enzymes (Castro et al., 2010). However, 4 It has been suggested that further reductions in these enzyme production costs might be difficult to realize (Klein-Marcuschamer et al., 2012). Despite similar enzyme production costs, the more recalcitrant nature of lignocellulosic biomass compared to starch means that much more enzyme is required to catalyze cellulose hydrolysis to sugars. Recent advances in cellulase production techniques, as well as increases in the efficiency of cellulase enzyme cocktails, have helped to successfully lower the cost of enzymes (per litre of cellulosic ethanol produced) by about 72% between 2008 and 2012 (Isola, 2013). Despite this decrease, the cost of the enzyme component for producing cellulosic ethanol is still far higher than that for producing starch-derived ethanol. Estimated enzyme component costs for producing cellulosic ethanol range from $0.13-0.38 USD/L ethanol (Klein-Marcuschamer et al., 2012), compared to $~0.01 USD/L corn ethanol (McAloon et al., 2000). This difference in enzyme costs arises not so much from differences in enzyme production costs, but rather from the amount of enzyme required for biomass deconstruction, with ~10-50g enzyme per kg cellulose required, compared to just 0.2-0.3g enzyme per kg starch (Arantes and Saddler, 2011; McAloon et al., 2000). The main reason for the large amount of enzyme required to break down lignocellulosic biomass is the heterogeneity and hierarchical structural organization of this substrate. Because this substrate is composed of a variety of different polymers tightly associated with one another, a large number of different enzymes with a range of activities on the different bonds present in the biomass are required. For example, in Trichoderma reesei, a fungal strain widely used for the industrial production of cellulolytic enzymes, more than 80 different enzymes are secreted (Vinzant et al., 2001). These enzymes include hydrolases involved in depolymerizing cellulose and hemicellulose which are complimented by the activities of lytic polysaccharide monooxygenases (LPMOs), which have been shown to cleave cellulose chains within the more crystalline regions of the biomass (Harris et al., 2010; Vaaje-Kolstad et 5 al., 2010), as well as to aid in the depolymerisation of some hemicelluloses (Agger et al., 2014). A more thorough review of these enzymes can be found in Section 1.2.1. In addition to the hydrolases and LPMOs involved in lignocellulose degradation, a third class of proteins has been implicated in biomass depolymerisation. These proteins appear to have no hydrolytic or oxidative activities, and have been termed ‘disruptive proteins’ or ‘amorphogenesis-inducing’ proteins (Reviewed in Arantes and Saddler, 2010). In contrast to the hydrolases and LPMOs, these amorphogenesis-inducing proteins do not appear to be directly capable of cleaving the covalent bonds between sugars. Instead, they appear to be involved in opening up and disrupting the biomass, possibly through the disruption of hydrogen bonding between the polymers (McQueen-Mason and Cosgrove, 1994). This promotion of amorphogenesis within the biomass is thought to play a role in enhancing the accessibility of the carbohydrate polymers to the activities of the hydrolases and LPMOs, thus enhancing the overall efficiency of the enzyme mixture (Arantes and Saddler, 2010). In the decades following the first report of these potentially disruptive proteins in 1991 (Din et al., 1991), surprisingly little has been discovered regarding exactly how these proteins disrupt and open up biomass. Since this initial report, a number of proteins from organisms spanning the Plantae, Fungi, and Bacteria kingdoms have been identified. A review of these proteins was recently published (Arantes and Saddler, 2010), and a summary of the amorphogenesis-inducing proteins identified to date can be found in Section 1.2.1.3 of this introduction. While a number of these proteins have been identified, little is known about the mechanisms that are employed to perform these disruptive activities and what role these proteins might play in possible synergism with the other families of enzymes involved in cellulose hydrolysis. The main reason for this lack of understanding stems from issues surrounding measuring and quantifying the effects of disruptive proteins on pure cellulosic or lignocellulosic biomass. When working 6 with hydrolases, the end products, soluble sugars, are relatively easy to measure through techniques such as high performance liquid chromatography (HPLC), the Glucose Oxidase assay (Berezin et al., 1977; Berlin et al., 2006a), or colorimetric assays such as the Dinitrosalicylic (DNS) acid assay (Miller, 1959). However, when working with amorphogenesis-inducing proteins, the end product is hard to describe, let alone quantify. For example, some of the terms put forward to describe the various effects of disruptive proteins have included delamination, fibrillation, decrystallization, swelling, loosening, splitting, roughening, pitting, or weakening (Arantes and Saddler, 2010). Adding to this ambiguity in the terms used to describe the end product of biomass disruption is the fact that these manifestations of amorphogenesis can occur at scales several orders of magnitude apart, from the nanoscopic decrystallization within a single elementary fibril (Gao et al., 2001), to the macroscopic dispersion of filter paper aggregates (Jäger et al., 2011). As a result of the lack of a defined end product and the great differences in scale between various disruptive effects, choosing and developing a technique to quantify the disruptive effects of the various amorphogenesis-inducing proteins is challenging. The review of applicable techniques that is covered in Section 1.7 describes the difficulties in quantifying protein-induced amorphogenesis and how this has hindered the development of an in-depth understanding of the roles of these proteins in cellulose hydrolysis. By developing a better understanding of the effects of amorphogenesis-inducing proteins on lignocellulosic structures, either through the application of tried and tested techniques or through the development of new ones, it should be possible to gain insights into the roles of these proteins in the overall process of enzymatic lignocellulose deconstruction. These insights should lead to the development of more efficient enzyme cocktails, able to rapidly reduce the carbohydrate polymers within biomass to their component sugars for subsequent conversion to fuels or chemicals. This will ultimately lower the amount (and therefore cost) of the enzymes required for biomass deconstruction, 7 thereby improving the economics of the biomass to fuels and chemicals process and narrowing the cost gap between producing starch-derived versus lignocellulose-derived fuels. In the following section, the structural properties of lignocellulosic biomass are discussed, with a focus on the specific factors that limit the efficiency of enzymatic hydrolysis. 1.1 Structural features of lignocellulosic biomass influencing enzymatic hydrolyzability The structure and composition of lignocellulosic biomass, unlike the relatively simple and homogenous structure of starch, is a complex, heterogeneous matrix (Himmel et al., 2007; Tester et al., 2004). This matrix is arranged into progressively more complex organizations at increasing size scales, from the nanoscopic packing and crystallization of individual cellulose chains, to the microscopic organization of microfibrils within the cell wall, to the macroscopic association of individual fibre cells into structures easily visible to the naked eye (Himmel et al., 2007; Mansfield et al., 1999; Sjöström, 1993; Somerville et al., 2004). In this section, the structural features of lignocellulosic biomass influencing enzymatic deconstruction are outlined, starting with a brief history on the origins of our understanding of cellulose structure. 1.1.1 A brief history on the structure of cellulose With regard to biomass-derived fuels and chemicals, cellulose is the most pertinent polymer within the lignocellulosic matrix, partly because it is the most abundant polymer within most biomass sources, and also because it is comprised of the hexose sugar glucose, which is readily fermented to ethanol or other products by industrial yeast strains (Aristidou and Penttilä, 2000). The term ‘cellulose’ was first put forward by the French Academy in 1839 by Brongniart et al., after Anselme Payen discovered in 1838 that all plant tissues yielded a resistant, fibrous substance when purified by acid-ammonia treatment followed by extraction with water, alcohol and ether (Brongniart et 8 al., 1839; Payen, 1838). This fibrous substance was named cellulose. Shortly after the discovery of cellulose, Carl von Nägeli, in 1858, suggested that bundles of cellulose formed crystalline structures, ubiquitously present among plant cell walls (Nägeli and Cramer, 1858; Wilkie, 1961). It wasn’t until 1937 that the structure of crystalline plant cellulose was first put forward (Meyer and Misch, 1937). Since then, a number of different crystalline allomorphs of cellulose have been identified, including Cellulose Iα, Iβ, II, III, and IV. Cellulose Iα and Iβ are by far the most abundant cellulose allomorphs found in nature, with the other allomorphs only produced artificially (although it has been suggested that some Cellulose II might exist in rare cases in nature (Roberts et al., 1989)). For this reason, only cellulose Iα and Iβ will be discussed here. With regard to these ‘native’ plant celluloses, a large number of X-ray diffraction, electron diffraction and neutron diffraction experiments have been performed, resulting in a widely accepted crystal structure for plant cellulose (Nishiyama et al., 2002). Briefly, this structure is made up of individual cellulose chains organized into sheets, which are in turn stacked upon each other. The cellulose chains themselves are exceptionally rigid and planar, due to the intramolecular hydrogen bonds formed between the exocyclic O6 primary alcohol hydroxyl group to the O2 secondary alcohol hydroxyl group of the subsequent residue, and from the O3 hydroxyl group to the O5 ring oxygen of the next sugar residue (Nishiyama et al., 2002). Sheets of parallel cellulose chains are associated laterally by Coulombic forces (predominantly hydrogen bonding) through equatorial hydroxyl groups. Finally, these cellulose sheets are stacked on top of each other, primarily associated via Van der Waals forces. Although these individual Van der Waals forces are weak, the large number of interactions between adjacent sheets leads to a relatively strong cumulative force (Pizzi and Eaton, 1985). In short, the cellulose crystal is made up of stacked sheets, where the dominant intraplane forces are Coulombic interactions, and the dominant interplane forces are Van der Waals interactions (Heiner et al., 1995). The combined intra- and intermolecular hydrogen bonds within cellulose result in the complete utilization of the potential hydrogen-bonding moieties, leading to an almost complete lack of hydrogen 9 bonding between sheets (Nishiyama et al., 2002). The interplane surfaces are thus relatively hydrophobic, which has implications for enzymatic hydrolysis, whereby not even water, let alone enzymes, can fit into the hydrophobic interplane spaces (Krässing, 1993), making enzymatic penetration into microfibrils challenging. Additionally, certain outer surfaces of the cellulose elementary- and micro-fibrils are hydrophobic, requiring enzymes active on these faces to contain a hydrophobic binding face (Creagh et al., 1996). The role of crystallinity in determining enzymatic hydrolysis rate is discussed in more detail below. When attempting to elucidate the crystal structure of native cellulose, it was found that for all plant-derived celluloses, diffraction patterns could not be fitted exactly to any single simple unit cell. Thus it was proposed that the crystalline regions of plant cellulose were composed of mixtures of two distinct crystal structures (Atalla and Vanderhart, 1984). The presence of two distinct, overlapping crystal structures within the basic crystalline unit of cellulose is one of the key factors which has impeded elucidation of the precise crystal structure of native plant cellulose, and which has led to over a century of debate on this topic. It is now known that crystalline plant cellulose is composed of a mixture of two allomorphs, Cellulose Iα, which has a triclinic unit cell containing one chain, and Cellulose Iβ, comprising a monoclinic unit cell containing two parallel chains (Reviewed in Nishiyama et al. 2002). The only physical difference between these two crystal forms is in the precise stacking pattern between the cellulose sheets. The planes themselves remain unchanged between Cellulose Iα and Iβ. In plant cellulose, specifically wheat straw, it has now been shown by high resolution atomic force microscopy (AFM) that Cellulose Iα and Iβ are intimately associated in the cell wall (Li et al., 2006). Shortly after it became recognized that cellulose took on a crystalline structure in nature, debate turned to how the elementary crystal structures were formed into the larger microfibrils that could be seen under scanning electron microscopy (SEM). As early as the 1950s it was suggested that small crystalline bundles, known as elementary fibrils, with postulated diameters of 7-9 nm, were 10 associated together to form larger structures containing both amorphous and crystalline cellulose. These larger structures were termed microfibrils, and were thought to have diameters of 15-25 nanometers (Frey-Wyssling, 1953). We now know that cellulose in higher plants is primarily produced by rosettes of cellulose synthase enzymes at the plasma membrane (Doblin et al., 2002). These rosettes are often composed of 30-36 subunits, producing 30-36 parallel chains of cellulose, which rapidly crystallize into elementary fibrils with approximately square cross-sections with diameters of ~3.5nm (Reviewed in Doblin et al. 2002). Very recent research has suggested that the cellulose at the center of elementary fibrils is highly ordered, with this order decreasing toward the outer layer of the elementary fibrils (Langan et al., 2014). These outer layers of the fibrils appear to be at least partially solvated, and in aggregates of elementary fibrils there does appear to be some degree of solvation between the individual elementary fibrils organized into the larger bundle (Langan et al., 2014). Despite decades of research, it is still unclear precisely how these elementary fibrils are associated together, along with amorphous regions of cellulose, to form microfibrils. Highlighting this, there still appears to be confusion in the literature regarding the definition of a microfibril, with some researchers using the term to refer to the smallest bundles of cellulose with diameters of ~3.5nm (referred to in this thesis as elementary fibrils), while other researchers use the term microfibril to describe bundles of elementary fibrils associated together with diameters of ~15-25nm (Lynd et al., 2002; Mansfield et al., 1999). In this work, the term elementary fibril is used to describe only the smallest cellulose fibrils, while the term microfibril is used to describe these larger associations of elementary fibrils. Although the general structures of elementary fibrils and microfibrils are now relatively well understood, there remains some disagreement over the precise distribution of crystalline and amorphous cellulose within microfibrils. In particular, it is unclear if amorphous cellulose is present in small quantities evenly distributed throughout the length of the microfibrils, or if amorphous cellulose is 11 localized to specific regions within the length of the microfibril (i.e. at specific ‘weak spots’, or ‘periodic disordered regions’) (Hidayat et al., 2012). The presence of these periodic disordered regions have been shown to occur within flax (Astley and Donald, 2001) and ramie (Nishiyama et al., 2003) fibres. Although the large difference in the period between these disordered regions (6-7nm for flax, ~150nm for ramie) raises some doubts as to the significance of these findings, it is possible that different species produce microfibrils with different periods between disordered regions. It is also possible that the 6-7nm periodic disorders reported for flax fibres could represent regions of very slightly misaligned residues, whereas the 150nm periodic disorders for ramie fibres could represent more significant weak spots within the elementary fibrils. In comparison, the recent commercial production of nanocrystalline cellulose particles has resulted in a relatively uniform length of the smallest cellulose particles produced after strong acid treatments. These particles are generally 100-200nm in length, and could represent the true length of the purely crystalline regions present within elementary cellulose fibrils. This 100-200nm periodicity has been demonstrated for hardwoods, softwoods, and cotton (Habibi et al., 2010), implying comparable ‘minimum crystalline unit lengths’ within the elementary fibrils of higher plants. Cellulose from organisms employing alternative cellulose synthesis regimes, such as bacteria, algae and tunicates, has been shown to form much longer nanocrystalline units (1000s of nm) (Habibi et al., 2010). This reinforces the probability that the length of the truly crystalline cellulose within the elementary fibrils of higher plants is an inherent property imparted during cellulose synthesis. After more than a century of research, the underlying cellulosic structures making up lignocellulosic biomass are relatively well characterized. However, an even better elucidation of the underlying structure is still required if we are to better understand how cellulose is degraded. In particular, how non-hydrolytic disruptive proteins might be acting to open up this structure. In the following sections, the organization of these basic cellulose structures within the lignocellulosic matrix is discussed, followed by an overview of the enzymes involved in degrading and disrupting this biomass. 12 1.1.2 The multilevel organization of lignocellulose In order to better understand the disruptive effects of non-hydrolytic proteins on lignocellulosic materials, the structural architecture of these materials can be categorized into three levels of organization: macroscopic, microscopic, and nanoscopic. At the macroscopic level, the cells of native plant material are bound together through the middle lamella, a glue-like composite rich in pectates and lignin, while in processed materials such as filter paper adjacent cells are held together primarily through hydrogen bonding (Sjöström, 1993). The macroscopic entity resulting from the association of adjacent cells is easily visible to the naked eye. The microscopic/ultrastructural organization of the cell wall, visible through techniques such as electron or confocal microscopy, consists primarily of bundles of 30-36 cellulose chains encased within hemicelluloses and lignin, forming structures known as microfibrils (although values of up to 200 associated chains have also been reported) (Somerville et al., 2004; Wyman et al., 2005). These microfibrils are arranged into sheets within the plant cell wall, with adjacent sheets having different microfibril angles (the angle between the longitudinal axis of the microfibril and the longitudinal axis of the cell) (Sjöström, 1993). Within these sheets, adjacent microfibrils are held together through complex interactions involving hemicelluloses and lignin, discussed in more detail below (Scheller and Ulvskov, 2010; Somerville et al., 2004). Finally, at the nanoscopic level, visible only through techniques such as atomic force microscopy (AFM) (Ding and Himmel, 2006; Harris et al., 2010), individual cellulose chains are organized into crystalline bundles, known as elementary fibrils, found within the microfibrils. At this level, the regular repeating units of the β-(1,4) linked D-glucose units (Purves, 1954) allow adjacent cellulose chains to associate, via hydrogen bonding and Van der Waals forces, into extremely compact structures (Heiner et al., 1995; Nishiyama et al., 2002; Pizzi and Eaton, 1985). The cellulosic cores of these microfibrils are 13 known to contain regions of extremely well ordered cellulose (known as crystalline regions), as well as less ordered regions (known as paracrystalline or amorphous regions) (Langan et al., 2014; Sjöström, 1993). In order to understand the mechanisms by which non-hydrolytic proteins might disrupt lignocellulosic substrates, it is necessary to develop a sound understanding of the molecular architecture at the various levels of organization within the substrate, from the nanoscopic structures of the smallest molecular building blocks, to the microscopic heterogeneous complexes resulting from the association of microfibrils, to the macroscopic structures arising from the association of adjacent fibre cells. The specific substrate characteristics contributing to the recalcitrance of native lignocellulosic materials to enzymatic degradation are discussed below. 1.1.3 Crystallinity Although there is now a tentative consensus over the crystal structures of the various allomorphs of cellulose, the influence of the degree of cellulose crystallinity (known as the crystallinity index, CrI) on enzymatic hydrolyzability of cellulose is a more contentious issue. Some authors argue that cellulose crystallinity is a key determinant of enzymatic hydrolysis rates (Chen et al., 2007; Fan et al., 1980; Hall et al., 2010; Lee et al., 1983), while others suggest that crystallinity has little effect on hydrolysis, particularly on lignocellulosic substrates (Agarwal et al., 2013; Converse, 1993; Kawakubo et al., 2010; Lynd et al., 2002; Mansfield et al., 1999; Ramos et al., 1999; Zhang and Lynd, 2004). One of the early theories on the influence of crystallinity on enzymatic hydrolyzability suggested that, if crystalline regions of cellulose were more recalcitrant to hydrolysis than amorphous regions, then a given substrate, containing both crystalline and amorphous regions, would increase in crystallinity over the course of hydrolysis as the more easily hydrolyzed amorphous regions were solubilized. This would leave behind the more crystalline regions (Betrabet and Paralikar, 1977). 14 Reaching a consensus on the effects of enzymatic hydrolysis on cellulose crystallinity has proved difficult, with several researchers finding that crystallinity did not appear to change (Ohmine et al., 1983; Puls and Wood, 1991), and others finding slight to moderate increases in cellulose crystallinity over the course of hydrolysis (Chen et al., 2007; Fan et al., 1980; Lee et al., 1983; Del Rio et al., 2012), and still others showing a brief increase in crystallinity at the early stages of hydrolysis, followed by a decrease and subsequent levelling off (Sinitsyn et al., 1989). Ramos et al. (1999) showed that a variety of enzyme mixtures did not appear to increase the crystallinity of fully bleached Kraft pulp, and Hall et al. (2010) found that Avicel crystallinity did not increase during enzymatic hydrolysis, while Chen et al. (2007), Fan et al. (1980), and Lee et al. (1983) did see an increase in the crystallinity of cellulose during hydrolysis. Contrary to this theory, Sun et al. (2011) actually observed a decrease in substrate crystallinity during enzymatic hydrolysis of pretreated corn cobs, as measured by wide angle X-ray diffraction (WAXD) (Sun et al., 2011). However, due to difficulties with using WAXD to differentiate between amorphous biomass in general and amorphous cellulose in particular, this result may simply be due to the relative increase in the amount of amorphous lignin and hemicellulose in the substrate, as the cellulose (both amorphous and crystalline) are hydrolyzed (Sun et al., 2011). Overall, it has been difficult to conclude unequivocally whether the cellulosic fraction of lignocellulosic biomass does become enriched in crystalline regions as hydrolysis preferentially removes the amorphous regions. The majority of past work assessing changes in cellulose crystallinity over the course of hydrolysis, particularly for lignocellulosic biomass, has not found a significant increase in cellulose crystallinity (Agarwal et al., 2013; Converse, 1993; Ramos et al., 1999; Zhang and Lynd, 2004). In addition to the idea that cellulose crystallinity should increase during enzymatic hydrolysis, it has been suggested that substrates with higher initial cellulose crystallinity should be more recalcitrant to enzymatic deconstruction than those with a lower initial crystallinity (Bertran and Dale, 1985; Reese et al., 1950). As with the theory suggesting that crystalline cellulose should become enriched during 15 hydrolysis, the idea that initial hydrolysis rate should correlate well with initial cellulose crystallinity has been difficult to prove, with a number of contradictory results in the literature. For example, in the case of pure cellulose (Agarwal et al., 2013; Hall et al., 2010), and some lignocellulosic materials (Fan et al., 1980; Sinitsyn et al., 1991) several studies have demonstrated that samples with lower crystallinity have higher initial hydrolysis rates than those with higher crystallinity. However, when working with heterogeneous lignocellulosic biomass, it is challenging to reduce the crystallinity of the biomass (by ball-milling, for example) without altering any other substrate properties, such as the accessibility, surface morphology and particle size of the biomass (Caulfield and Moore, 1974; Howell and Stuck, 1975; Lee and Fan, 1982; Rivers and Emert, 1987; Saddler et al., 1982; Schwald et al., 1988; Yoshida et al., 2008). Thus it is difficult to tell whether enhancements in hydrolysis after treatment to reduce crystallinity are indeed due to the reduction in crystallinity, or are in fact due to changes to other substrate characteristics intrinsically linked to crystallinity (Caulfield and Moore, 1974; Howell and Stuck, 1975; Jeoh et al., 2007; Lee and Fan, 1982). It should be noted that when dealing with lignocellulosic rather than pure cellulosic substrates, the majority of studies aimed at correlating lignocellulosic substrate properties with enzymatic hydrolyzability have found that factors such as lignin content and accessibility play a more significant role in enzymatic hydrolyzability than cellulose crystallinity (Agarwal et al., 2013; Kawakubo et al., 2010; Puri, 1984; Ramos et al., 1993, 1999). For example, Kawakubo et al., (2010) demonstrated that the initial crystallinity index for Japanese cedar wood pretreated under a variety of different conditions did not correlate with overall saccharification yield. These authors did observe a correlation between saccharification yield and cellulose accessibility measured using cellulose-specific carbohydrate binding modules (CBMs) as probes (Kawakubo et al., 2010). Research by Hall et al. (2010) attempted to determine whether accessibility or crystallinity played a more significant role in the slowdown of enzymatic hydrolysis rate at longer hydrolysis times. In this work, the authors treated Avicel to increasing concentrations of phosphoric acid to create a range of 16 cellulosic substrates with varying crystallinity and accessibility. When correlating the initial hydrolysis rate with the amount of enzyme adsorbed to the cellulose (accessibility), the correlation only held at CrI values below ~40%. However, when initial hydrolysis rate was correlated with CrI values, lower crystallinity led to higher initial hydrolysis yields for all CrI values tested. In other words, the crystallinity, and not the amount of adsorbed enzyme, appeared to be the determining factor for initial hydrolysis rate (Hall et al., 2010). When studied in isolation on pure or almost pure cellulosic substrates, it appears that cellulose crystallinity does play a role in determining enzymatic hydrolysis rate (Agarwal et al., 2013; Hall et al., 2010). However, when heterogeneous pretreated lignocellulosic substrates are used, other factors, such as the type, distribution, and amount of lignin and hemicellulose, and in particular the accessibility of cellulose to cellulase, likely play a more significant role in determining the enzymatic hydrolyzability of the biomass (Agarwal et al., 2013; Arantes and Saddler, 2010; Converse, 1993; Lynd et al., 2002; Ramos et al., 1999; Sinitsyn et al., 1991; Zhang and Lynd, 2004). For example, Sinitsyn et al. (1991) showed a linear correlation between the crystallinity and hydrolyzability of a range of pure cellulosic substrates, but for pretreated lignocellulosic substrates, a linear correlation was found between hydrolyzability and accessible surface area, but not with crystallinity. This idea is supported by Agarwal et al. (2013) in particular, who state that cellulose crystallinity and composition are not as important as the ultrastructural changes caused by the disruption of the tightly packed regions of the cell wall. One reason why there appears to be so much contradiction in the literature regarding the role of crystallinity in the hydrolysis of biomass, even when working with pure cellulose, are the techniques used to measure crystallinity. There are no standardized protocols or techniques for measuring crystallinity, so comparison between crystallinity values published by different groups is not easy (Park et al., 2010). In addition, these techniques generally require substrate drying prior to analysis which can 17 alter the crystallinity of cellulosic samples (Newman, 2004; Park et al., 2009; Rebuzzi and Evtuguin, 2005). For example, even when research groups applied the same XRD technique for measuring crystallinity to a simple, standardized substrate (in this case Avicel PH-101), widely different values were observed, ranging from 62% to 87.6% using the peak height method (De Souza et al., 2002; Thygesen et al., 2005), and from 39% to 75.3% using various other methods (Ardizzone et al., 1999; Thygesen et al., 2005). This variability in measured crystallinity indices for a substrate as simple and standardized as Avicel highlights the difficulties involved in getting reproducible crystallinity indices between different research groups. These difficulties are compounded further when complex, heterogeneous lignocellulosic substrates are used in place of Avicel. While it is difficult to determine exactly how important cellulose crystallinity itself is to enzymatic hydrolysis of lignocellulosic biomass, it is clear that the degree of crystallinity of a given substrate is intrinsically linked to other substrate properties, such as cellulose accessibility. For example, Jeoh et al. (2007) suggest that, rather than thinking of crystallinity and accessibility as two separate factors, crystallinity should in fact be thought of as just one of the many factors influencing the accessibility of cellulose to cellulases. This idea makes sense when considering cellulose at the nanoscopic level of individual chains, as a crystalline bundle of cellulose would in essence simply be a region where the outer cellulose chains block access to the chains in the interior of the bundle. Past research assessing the relationship between cellulose crystallinity and ease of enzymatic deconstruction, specifically in the case of pure cellulosic substrates, suggests that crystallinity likely does play a role in determining enzymatic hydrolysis (Fan et al., 1980; Hall et al., 2010; Lee et al., 1983). When working with lignocellulosic biomass, it is likely that, once the enzymes gain access to the cellulose buried within the lignocellulosic matrix, the crystallinity of that cellulose probably does play a role in how easily the enzymes can hydrolyze it. However, factors such as lignin and hemicellulose content, 18 ultrastructural organization, and in particular the influence that these compounds have in determining the cellulose accessibility to cellulase, all appear to affect the efficiency of enzymatic deconstruction to a greater extent than does just cellulose crystallinity. The influence of hemicelluloses, lignin, and accessibility on enzymatic hydrolysis is discussed below. 1.1.4 Hemicelluloses Hemicelluloses are the second most abundant source of carbohydrates, after cellulose. While the term hemicellulose has traditionally referred to all non-cellulosic, non-pectin carbohydrate polymers in biomass, it has recently been proposed that this definition should be narrowed to include only those carbohydrate polymers comprised of a backbone consisting of β-1,4 linked glucose, xylose or mannose (Scheller and Ulvskov, 2010). Since these sugars all have the same equatorial position at C1 and C4, this gives significant structural similarity to the backbones of this heterogeneous family of carbohydrates (Scheller and Ulvskov, 2010). Unlike the linear unbranched glucopyranose homopolymeric structure of cellulose, hemicelluloses can be composed of a variety of different sugar monomers, and the backbones are often substituted with side chains. The following information on hemicelluloses is a simplification of the complex heterogeneous hemicellulosic structures, and the exact composition and degree of substitution of the particular hemicelluloses varies between species and cell types. For an in-depth review of hemicellulose structure, see Scheller and Ulvskov (2010). The predominant forms of hemicellulose found in secondary plant cell walls are glucuronoxylan, glucuronoarabinoxylan, galactoglucomannan and, to a lesser extent, glucomannan. Glucuronoxylan is predominantly found in dicots such as hardwoods, making up 20-30% of the secondary cell wall, and is composed of a β-1,4 linked xylose backbone substituted mostly with glucuronic acid, but also with minor amounts of O-methylated glucuronic acid and arabinose (Pinto et al., 2005; Sjöström, 1993). Glucuronoarabinoxylan is found in significant amounts in grasses and conifers, making up 40-50% and 5-19 15%, respectively, of these secondary cell walls (Ishii, 1997; Scheller and Ulvskov, 2010). The backbone of glucuronoarabinoxylan is composed of β-1,4-linked xylose, and this backbone can be substituted with a variety of different moieties, including arabinose, feruloylated arabinose, glucuronic acid, O-methylated glucuronic acid, and acetyl groups (Scheller and Ulvskov, 2010). In addition, some of the backbone xyloses can be substituted at multiple sites, adding further complexity to this polymer. Galactoglucomannan is predominantly found in conifer secondary cell walls, making up 10-30% of these walls, but can also be found in low amounts (0-3%) in dicots (Hamilton et al., 1960; Scheller and Ulvskov, 2010; Sjöström, 1993). This polymer has a backbone composed mostly of β-1,4 linked mannans, but also containing some glucose residues (within the β-1,4 linked backbone). This heteropolymeric backbone is substituted with galactose, but to a lesser extent than the degree of substitution of glucuronoxylan and glucuronoarabinoxylan. Finally, glucomannan, which has a similar structure to galactoglucomannan but with less/no substitution, makes up 2-5% of dicot and 0-5% of grass secondary cell walls (Scheller and Ulvskov, 2010). In the plant cell wall, hemicelluloses are tightly associated with both the cellulose and lignin components. Hemicelluloses interact with cellulose predominantly through hydrogen bonding to the surface of cellulosic elementary fibrils. However, in addition to these surface interactions, it is thought that soluble hemicellulose in close proximity to the cellulose synthase complexes at the plasma membrane can also get ‘zipped in’ to the interior of cellulosic elementary fibrils and microfibrils immediately after cellulose synthesis (Hackney et al., 1994; Iwata et al., 1998; Tokoh et al., 1998). These interactions between hemicelluloses and cellulose result in the formation of a partial hemicellulosic coating around the cellulosic cores of microfibrils, which shields the interior cellulose from enzymatic attack (Hayashi et al. 1987; Vincken et al. 1995). In addition to the interactions with cellulose, hemicelluloses have also been shown to interact with lignin through electrostatic and covalent interactions. In particular, ester bonds between lignin and methylglucuronic acid residues have been 20 documented, as well as ether bonds between arabinosyl residues and lignin (Eriksson et al., 1980). Low molecular weight phenolic compounds such as ferulic acid and p-coumaric acid have also been found covalently linked to hemicelluloses (Hartley and Ford, 1989). Although each of these individual interactions documented to date were observed on a single biomass source, it seems likely that similar interactions between hemicelluloses and cellulose/lignin are present in most plant species. In short, hemicellulose effectively masks the cellulosic fraction from enzymatic attack by hydrogen bonding directly to the cellulose, and by covalently linking to lignin, facilitating the formation of a hemicellulose-lignin shield around cellulose microfibrils. In addition to the steric effects imparted by hemicelluloses hindering the access of cellulases to the cellulosic fraction, some of the side groups on hemicelluloses have specific inhibitory effects on enzyme action. For example, ester-linked acetyl groups on hemicelluloses have been found to block enzymatic hydrolysis of the hemicellulosic fraction, with deacetylation of xylan improving enzymatic digestibility by 5-7 fold (Biely et al. 1985; Chen et al. 2012; Chen et al. 2014; Grohmann et al. 1989). This improvement in xylan digestibility in turn enhanced the cellulose digestibility by 2-3 fold (Grohmann et al., 1989). In general, the presence of side chains and acetylated residues requires the application of additional enzymes to remove these residues prior to the depolymerisation of the hemicellulose backbone (Decker et al., 2008). Thus, the heterogeneity of hemicelluloses, and the presence of a variety of side chains, hinders the enzymatic deconstruction of the hemicellulosic fraction of the biomass. The factors influencing biodegradability of the hemicellulosic fraction are of particular importance due to the role that hemicelluloses play in shielding the cellulose. In other words, factors which hinder enzymatic hydrolysis of the hemicellulosic fraction will in turn prevent rapid hydrolysis of the cellulosic fraction. A number of studies have emphasised the importance of hemicelluloses in influencing the ease of enzymatic hydrolysis of cellulose in industrially-relevant pretreated lignocellulosic substrates. For 21 example, Hu et al. (2011) demonstrated that replacing a portion of the cellulase with a commercial xylanase-rich enzyme preparation during the hydrolysis of steam pretreated corn stover (SPCS) dramatically enhanced the hydrolysis of the cellulosic fraction even though less cellulase was present in the system. In subsequent work, it was found that purified xylanases enhanced the catalytic efficiency of the cellulase Cel7A by more than 80% on a range of xylan-containing substrates, including steam pretreated corn stover, sugarcane bagasse, and poplar (Hu et al., 2013). In addition to this work, several other papers have demonstrated that hemicelluloses play a major role in inhibiting cellulose hydrolysis (Grohmann et al., 1989; Kumar and Wyman, 2009; Ohgren et al., 2007). In particular, Kumar and Wyman (2009) found an almost linear correlation between the amount of xylan removed from Ammonia Fibre Expansion (AFEX) treated corn stover and subsequent glucose released during hydrolysis, while Ohgren et al. (2007) demonstrated that near-theoretical conversion levels of the cellulosic fraction of steam pretreated corn stover could be obtained only when the cellulase mixture was supplemented with xylanase. In summary, the effective modification or removal of hemicellulose is crucial in order to achieve efficient hydrolysis of the cellulosic fraction of industrially relevant pretreated lignocellulosic substrates. 1.1.5 Lignin Lignin macromolecules are derived from the oxidative polymerization of phenylpropane monolignols, the most prevalent of which are p-coumaryl alcohol, coniferyl alcohol, and sinapyl alcohol, which are each composed of a methoxylated aromatic ring attached to a 3-carbon aliphatic tail (Glasser, 1980; Sarkanen and Ludwig, 1971; Sjöström, 1993). During the formation of secondary plant cell walls, these monolignols are secreted into the cell wall, then oxidatively dehydrogenated to form crosslinked high molecular weight, heterogeneous macromolecules (Boerjan et al., 2003). While most researchers consider this polymerization event to be a predominantly random process (known as the Random 22 Coupling Model), the discovery of ‘dirigent’ proteins, which have been shown to catalyze the formation of specific bonds between lignin monomers have prompted some to propose the Dirigent Protein Model, whereby the bonds formed during polymerization of lignin are suggested to be catalyzed via the specific action of dedicated enzymes (Davin and Lewis, 2000; Davin et al., 1997; Hatfield and Vermerris, 2001). However, the inability to find enzymes with the ability to catalyze the formation of the β-O-4 linkage (the most common linkage in most lignin macromolecules), has raised doubts that lignin polymerization is predominantly catalyzed by dirigent proteins (Boerjan et al., 2003; Hatfield and Vermerris, 2001). There does not appear to be a current consensus on the precise mechanism of lignin formation in the literature, although most researchers do appear to agree that there is some degree of randomness in the formation of the lignin macromolecules (Boerjan et al., 2003; Davin et al., 2008; Hatfield and Vermerris, 2001). One proposal which can at least partially reconcile these two theories is the suggestion that dirigent proteins may be involved in the initiation of lignin polymerization, the bulk of which is then subsequently carried out via the random coupling pathway (Davin and Lewis, 2000; Hatfield and Vermerris, 2001). After polymerization, the monolignols mentioned above are found in the form of p-hydroxyphenyl (H), guaiacyl (G), and syringyl (S) lignins, respectively. These monolignols differ in the number of methoxyl groups found on the benzene ring, with H monomers containing no methoxyl group, G containing one, and S containing two (Reviewed in (Vanholme et al., 2010)). The presence and distribution of these methoxyl groups on the rings affect the recalcitrance of the biomass to enzymatic deconstruction (Santos et al., 2012). This is because each additional methoxyl group takes up a potential site for crosslinking to the aromatic ring, effectively blocking this site from forming covalent linkages to adjacent lignin or carbohydrate moieties (Fengel and Wegener, 1983; Glasser, 1980). Thus, lignin rich in monomers with a lower number of methoxyl groups, such as guaiacyl lignin (one methoxyl group), are capable of forming a more condensed, hydrophobic, recalcitrant lignin macromolecule than those with 23 more methoxyl groups, such as syringyl lignin (two methoxyl groups). The precise ratios of these three building blocks vary dramatically between species. For example, softwood lignin tends to be composed almost entirely of G units, with low amounts of H units, resulting in the formation of a particularly condensed, hydrophobic and recalcitrant lignin. On the other hand, hardwoods have a higher proportion of S units, which leads to a less condensed and less recalcitrant lignin structure (Fengel and Wegener, 1983; Sjöström, 1993). In addition to linkages forming between lignin moieties during polymerization, linkages are also formed between lignin and certain carbohydrates within the biomass (Fengel and Wegener, 1983; Grabber, 2005; Jin et al., 2006; Scheller and Ulvskov, 2010; Sjöström, 1993; Whetten and Sederoff, 1995). For example, linkages to carbohydrates can occur through ester linkages to 4-O-methylglucuronic acid groups present in some xylans or through ether linkages to arabinose or mannose present in glucuronoarabinoxylan or glucomannan, respectively (Sjöström, 1993). Additionally, further linkages between hemicellulose and lignin can occur through the ferulic and possibly p-coumaric acid esters associated with grass xylans (Scheller and Ulvskov, 2010). Thus, lignin macromolecules are comprised of a covalently crosslinked, very high molecular weight structure, which in turn is covalently linked to the carbohydrate fraction of the biomass. There are three main mechanisms through which lignin has been suggested to contribute to the recalcitrance of lignocellulosic biomass: through sterically blocking access of the depolymerizing enzymes to the cellulosic and hemicellulosic fractions of the biomass (Kumar et al., 2012; Mooney et al., 1998), by inactivation of these enzymes via non-productive binding to the lignin via hydrophobic interactions (Berlin et al., 2005, 2006b; Eriksson et al., 2002; Nakagame et al., 2011a, 2011b; Palonen et al., 2004; Sutcliffe and Saddler, 1986), effectively removing them from the pool of active enzymes in the system, and through the inhibitory effects of soluble lignin derivatives on the depolymerizing enzymes 24 (Kim et al., 2011; Ximenes et al., 2011). Modifying the surface of lignin within pretreated lignocellulosic biomass, either through sulfonation or surfactant treatment, reduces non-specific enzyme binding to lignin while not significantly affecting the steric effects of the lignin (Eriksson et al. 2002; Mooney et al. 1998). These treatments have been shown to enhance the release of glucose by cellulases (Eriksson et al. 2002; Mooney et al. 1998). Additionally, work by Kumar et al. (Kumar et al., 2012) demonstrated the relative effects of steric blocking and non-productive adsorption by completely delignifying a substrate then adding back the lignin. This reduced the steric blocking effects of the lignin without mitigating the non-productive binding to the enzymes (Kumar et al. 2012), and revealed that the steric blocking effect of lignin likely played a more inhibitory role in enzymatic deconstruction than the non-productive adsorption effect. Application of these approaches by various researchers have demonstrated that both non-productive binding and steric effects of lignin play a role in inhibiting enzymatic cellulose hydrolysis (Eriksson et al., 2002; Kumar et al., 2012; Mooney et al., 1998; Palonen et al., 2004). However, it has been difficult to determine which of these mechanisms contributes more to recalcitrance. The relative inhibitory effects of these two mechanisms have been found to be dependent on enzyme loading, where low enzyme loadings were found to be more inhibited by non-productive adsorption, while reactions carried out at higher enzyme loadings were found to be more inhibited by the restrictions on cellulose accessibility imposed by the lignin (Kumar et al., 2012). Overall, it appears that the steric effects of lignin likely play a more inhibitory role than non-productive adsorption effects, as a number of researchers have found very good correlations between accessibility and hydrolyzability, while correlations between lignin content and hydrolyzability have been less consistent (Arantes and Saddler, 2011; Wiman et al., 2012). The role of accessibility in enzymatic hydrolysis is discussed in detail in the following section. 25 1.1.6 Cellulose accessibility Cellulose accessibility to cellulases has often been suggested to be the major determinant of enzymatic hydrolyzability on a broad range of pretreated lignocellulosic substrates (Arantes and Saddler, 2010, 2011; Chandra et al., 2008a; Esteghlalian et al., 2001a; Grethlein, 1985; Sinitsyn et al., 1991; Thompson et al., 1992; Wiman et al., 2012). Although cellulose is usually the most abundant component of the cell wall, enzymatic access to this cellulose is not trivial, due to the compact fibrillar architecture of the cellulose itself, as well as the tight association of this cellulose with other compounds, such as hemicellulose and lignin. As described above, the diffusion of cellulase enzymes to the cellulose is thwarted by substrate factors at various levels of organization. For example, at the macroscopic level, when deconstructing biomass particles comprised of aggregates of fibre cells, the cellulases must be capable of diffusing through/between the outer layer of cells to gain access to the fibre cells within the interior of the aggregate. At the microscopic level, it has been suggested that barriers to accessibility include the inability of cellulases to diffuse/penetrate past the outermost layer of microfibrils at the surface of the fibre cell (Mansfield et al., 1999). Finally, at the nanoscopic level, the extremely tight packing of individual cellulose chains into elementary fibrils prevents cellulases from accessing the cellulose chains interior to these bundles (Krässing, 1993). These many factors that limit the access of cellulases to the cellulose have been suggested to play a key role in preventing the rapid and complete enzymatic conversion of the cellulosic fraction of the biomass into sugars (Arantes and Saddler, 2010, 2011; Chandra et al., 2008a; Esteghlalian et al., 2001a; Grethlein, 1985; Sinitsyn et al., 1991; Thompson et al., 1992; Wiman et al., 2012). When working with lignocellulosic substrates, the hemicelluloses and lignin, and possibly the crystallinity of the cellulose itself, are all thought to play a role in limiting hydrolysis rate. However, all the factors which have been suggested to hinder enzymatic hydrolysis also affect the accessibility of the 26 cellulose to cellulases. For example, the masking of cellulose fibrils by a sheath of hemicellulose and lignin will reduce the accessible surface area of the cellulose. Additionally, crystalline bundles of cellulose will result in the interior chains being shielded from cellulases by the outermost chains of the crystalline bundle. Thus, of all the factors that might limit the rate and extent of enzymatic hydrolysis, the degree of cellulose accessibility to cellulases will likely provide the most accurate predictor of enzymatic hydrolysis as this is the substrate characteristic which best reflects the cumulative effects of many other substrate properties (such as hemicellulose/lignin masking, and cellulose crystallinity). While this concept is somewhat oversimplified (for example, as well as sterically sheathing cellulose, lignin is also involved in sequestering cellulases via non-productive adsorption), the relationship between cellulose accessibility and enzymatic hydrolyzability is one of the most well-documented correlations in the field of bioconversion. Rollin et al. (2011) demonstrated that, when working with switchgrass, increasing the accessibility of the cellulose to cellulases was a more important factor in enhancing enzymatic hydrolyzability than delignification. Additionally, work on other pretreated lignocellulosic substrates, such as corn stover (Jeoh et al., 2007; Zhu et al., 2009), spruce (Wiman et al., 2012) and Douglas fir (Mooney et al., 1998), to name a few, has shown that increasing the accessibility of the cellulose enhances the efficiency of enzymatic hydrolysis. Finally, even when comparing highly diverse substrates such as softwoods, hardwoods, and agricultural residues, strong correlations were observed between cellulose accessibility and enzymatic hydrolyzability when similar compromised pretreatment conditions were used (Arantes and Saddler, 2011). Although cellulose accessibility to cellulases does appear to be an important factor in determining enzymatic hydrolyzability, it must be noted that, just because cellulose is accessible to cellulase, this does not necessarily mean that this cellulose is readily hydrolyzable. This is partly due to the fibrillar organization of the cellulose within elementary fibrils, where different longitudinal faces of 27 the elementary fibrils present a different structural motif for cellulase recognition (Lehtiö et al., 2003; McLean et al., 2000; Tormo et al., 1996). This impacts recognition of the cellulose by cellulases, which are often targeted to the cellulose surface by CBMs which recognize specific faces of the elementary fibril (McLean et al., 2002). For example, the CBM of the commercially important cellulase, Cel7A from T. reesei, specifically recognizes the hydrophobic 110 face of crystalline Valonia cellulose fibrils (Lehtiö et al., 2003). The importance of CBM specificity and surface morphology of cellulose will be discussed in more detail in Section 3.2.1 of this thesis. In addition to biomass composition, cellulose crystallinity and the overall accessibility of the cellulose to cellulases, certain physical characteristics of cellulosic fibres have been suggested to influence hydrolyzability. Specifically, the cellulose within fibre dislocations has been suggested to hydrolyze more readily than the cellulose present within the bulk of the fibre (Ander et al., 2008; Fernandez and Young, 1996; Lee et al., 1983). 1.1.7 Fibre dislocations The presence of fibre dislocations (also called kinks, microcompressions, irregularities, and slip planes (Nyholm, et al., 2001)) has been observed across a range of plant species, including softwood, hemp, flax, and wheat (Reviewed in (Hidayat et al., 2012). These dislocations are thought to be present in untreated biomass fibres, but are also induced during processing steps (Hidayat et al., 2012). Typically, the fibre structure within the dislocations contains surface features perpendicular to the direction of the microfibrils. These dislocations have been described as containing more amorphous cellulose with less order than the surrounding fibre (Ander et al., 2008; Fernandez and Young, 1996; Lee et al., 1983). These regions have also been described as a distortion of the crystalline cellulose microfibrils (Lee et al., 1983) although other work has suggested that microfibrils continue through these dislocations (Thygesen et al. 2007). Overall, it is generally accepted that dislocations within fibres 28 make up weak points of the fibre that are rapidly cleaved by cellulases. However, the precise reason why these regions are more easily hydrolyzed is still incompletely understood. Several previous studies have used CBMs with different specificities for crystalline and amorphous regions of the cellulose to try and reveal the differences in surface morphology between the surrounding fibre and the cellulose within the dislocations (Ding et al., 2006; Hidayat et al., 2012; Kawakubo et al., 2010). In general, CBMs can be grouped into three Types. Type A CBMs which contain planar binding faces that preferentially bind to the flat surfaces present on crystalline arrangements of carbohydrates (i.e. crystalline cellulose), Type B CBMs which contain cleft-shaped binding sites that bind specifically to isolated carbohydrate chains, such as amorphous cellulose, and Type C CBMs which contain a binding pocket, that binds to the ends of carbohydrate chains, or to monomers and short oligomers (Boraston et al., 2004). Different CBMs within each Type can have specificities for different carbohydrate polymers. When CBMs which preferentially bind to crystalline cellulose (Type A) or to amorphous cellulose (Type B) were used to probe the dislocations present on cellulose fibres, no clear consensus was reached, with some Type A CBMs not binding to dislocations, while other Type A CBMs did bind at these dislocations. Interestingly, all Type B CBMs tested to date do appear to localize to the fibre dislocations (Ding et al., 2006; Kawakubo et al., 2010), suggesting that these regions are generally less ordered and contain more isolated carbohydrate chains that are not laterally associated into fibrils than the bulk of the fibre. However, although these dislocations appear to be inherently more disordered than the surrounding fibre, polarized light microscopy has demonstrate that these regions are still birefringent, implying that some crystalline structural organization is still present within these dislocations (Hidayat et al., 2012). Overall, fibre dislocations appear to be the weak points of the fibre, enriched in amorphous cellulose, and more susceptible to hydrolysis (acid or enzymatic) than the surrounding more ordered fibre structure. 29 Typically, to achieve effective cellulose hydrolysis some form of pretreatment is required to open up the biomass for subsequent enzymatic deconstruction. Much of the earlier discussion has focussed on the native structure of unpretreated plant cell walls. Although the fibrillar architecture of the lignocellulosic matrix remains relatively intact after pretreatment, this process is known to alter the structure of this biomass in a variety of ways. Thus pretreatment-induced changes in biomass structure and composition are briefly discussed in the following section. 1.1.8 Effects of pretreatments on the structure of lignocellulosic biomass While a thorough review of the effects of the various proposed pretreatments on the structure and properties of lignocellulosic biomass is beyond the scope of this thesis, this section will briefly summarize the effects of one of the leading pretreatment technologies, steam pretreatment, which was predominantly employed in this work (Brownell and Saddler, 1987; Galbe and Zacchi, 2007; Jørgensen et al., 2007). During steam pretreatment, the biomass is heated with steam under pressure in the presence or absence of an acid catalyst. At the end of a set residence time, usually 2-5 minutes, the pressure in the system is rapidly released, resulting in a forceful physical dispersion of biomass fibres. From a compositional point of view, steam pretreatment leaves essentially all of the lignin and cellulose in the biomass, while dissolving a portion of the hemicellulosic fraction. This dissolution of hemicellulose is enhanced in the presence of an acid catalyst, such as SO2 or H2SO4 (Söderström et al., 2003). In the case of uncatalyzed steam pretreatment, solubilisation of acetic acid via the release of the acetyl groups present on the hemicellulose results in a lowering of the pH. This solubilized acetic acid is then free to perform acid-catalyzed hydrolysis of a portion of the hemicellulose in a process known as autocatalysis (Kim et al., 2009). Overall, the compositional change in biomass after acid-catalyzed or autocatalyzed steam pretreatment is predominantly the reduction in hemicellulose content, which increases the proportion of lignin and cellulose in the pretreated biomass. 30 In addition to compositional changes, one of the most pronounced effects of steam pretreatment appears to be the redistribution and restructuring of the remaining biomass components. In particular, recent studies have suggested that the cellulosic fraction of the biomass actually becomes less hydrated and more ordered, with the coalescing of elementary fibrils into larger macrofibrils after pretreatment (Langan et al., 2014). This effect has been noted previously, and was suggested to be due to the removal of hemicellulose, resulting in alignment and aggregation of adjacent cellulose chains, leading to an increase in apparent crystallinity of the cellulose (Ramos et al., 1993). However, this recent work suggests that this increasing order of the cellulose after pretreatment could be due to the entropically unfavorable confinement of water at the surface of and in between relatively loosely associated bundles of elementary fibrils (Langan et al., 2014; Pingali et al., 2014). It appears that during the high-temperatures associated with steam pretreatment, where the entropic penalty of water confinement is increased, water is released from the system. This results in the irreversible dehydration of the elementary fibrils (Langan et al., 2014; Pingali et al., 2014). While this effect in itself is likely not beneficial to subsequent enzymatic hydrolysis, it appears that this reordering of the cellulose is an unavoidable consequence of the high temperatures associated with effective pretreatment. As well as this apparent dehydration and reordering of the cellulose, significant changes in the distribution and structure of the lignin and hemicellulose also occur during steam pretreatment. In particular, it has been shown that the lignin fraction of the biomass softens at the higher temperatures associated with steam pretreatment, enabling this lignin to seep out from within the biomass, forming globular structures on the surface of microfibrils (Donaldson et al., 1988; Selig et al., 2007). These redistributed lignaceous entities have been shown to also contain a significant amount of carbohydrates (Selig et al., 2007), suggesting that a portion of the hemicellulose (likely the portion closely associated with the lignin via lignin-carbohydrate complexes, discussed above) is also redistributed in this process. Thus, the key effects of steam pretreatment of biomass appear to be the decrease in particle size via the 31 rapid decompression of the pretreatment chamber, the solubilization of a portion of the hemicellulose (either through acid or auto-catalysis), the dehydration and reorganizing of the cellulose, and the softening and redistribution of the lignin (and some portion of the carbohydrates). Overall, these effects act to greatly increase the overall accessibility and digestibility of the cellulosic fraction to cellulases. An alternative pretreatment used in this thesis was a solvent-extraction based pretreatment, organosolv pulping (Chandra et al., 2007; Pan et al., 2006). This process involves heating biomass with an organic solvent such as ethanol in the presence or absence of an acid catalyst (Pan et al., 2006). Unlike steam pretreatment, organosolv pretreatment primarily solubilizes the lignin (Sewalt et al., 1997; Chandra et al., 2007). Although most pretreatments are effective in opening up and enhancing the accessibility of biomass, the pretreatment conditions used will always be a compromise. This is because very severe pretreatments (i.e. high temperature, pressure, and/or acid loading) promote the degradation of some (primarily hemicellulose derived) sugars within the biomass to compounds such as furfural and hydroxymethyl furfural, which are inhibitory to downstream processes such as enzymatic hydrolysis and fermentation (Ramos 2003). Additionally, higher severity pretreatments require greater energy and chemical inputs, while resulting in yield loss due to the degradation of potentially fermentable sugars, thereby significantly lowering the overall efficiency of the bioconversion process. For these reasons, pretreatment conditions are optimized to achieve maximum accessibility while promoting minimal sugar degradation, using the lowest energy and chemical inputs possible (Chandra et al., 2007). After pretreatment, the cellulosic fraction of the biomass is made significantly more accessible to the enzymes. However, the fibrillar structure of cellulose means that a considerable amount of this material must still be largely inaccessible to enzymatic attack. This implies that the enzyme mixture must be capable of disrupting and opening up this matrix, as well as the individual fibrils, in order to access 32 the cellulose chains buried within the fibrillar architecture of the cell wall. In addition to the suite of hydrolytic and oxidative enzymes known to be involved in directly depolymerizing the carbohydrate chains, there also appears to be a family of proteins specifically involved in opening up and disrupting the structure of the cell wall, in a process known as amorphogenesis (Arantes and Saddler, 2010). These proteins are thought to have the role of enhancing cellulose accessibility to cellulases, thereby improving the overall efficiency of the enzymatic hydrolysis step. The hydrolytic and oxidative enzymes involved in depolymerizing cell wall carbohydrates are briefly discussed below, followed by a more detailed description of the various non-hydrolytic, non-oxidative amorphogenesis-inducing proteins identified to date. 1.2 Enzymatic deconstruction of lignocellulosic substrates Due to the recalcitrant, heterogeneous nature of lignocellulosic materials, the enzymatic deconstruction of biomass is a complex process, involving a variety of different enzymes with different activities, and proceeding through a number of distinct steps during the deconstruction process. This process can be broadly broken down into three steps: the ‘amorphogenesis’ step, where the biomass is opened up and made more reactive to enzymatic attack, followed by the solubilisation step, where longer insoluble carbohydrate polymers are cleaved into shorter, soluble fragments, and finally the depolymerisation of these solubilized oligomers to monomers (Arantes and Saddler, 2010; Coughlan, 1985; Reese et al., 1950). Over the past several decades, a vast amount of research has been carried out to elucidate the roles of various enzymes in the process of biomass deconstruction. However, the majority of this work was carried out on enzymes known to play a role in the latter steps of this process. In other words, the enzymes involved in solubilisation and depolymerisation have been well studied, whereas those involved in the amorphogenesis step have remained elusive (Arantes and Saddler, 2010). Indeed, at this 33 point the enzymes known to be involved in opening up and swelling the biomass (promoting amorphogenesis) can be counted on both hands, while those known to be involved in solubilizing and depolymerizing the biomass number in the hundreds. Although this knowledge gap is at least somewhat excusable due to the challenges associated with measuring the effects of amorphogenesis-inducing proteins (discussed in more detail in the ‘techniques’ section), the lack of knowledge of the enzymes involved in promoting amorphogenesis is particularly vexing due to the highly influential role that cellulose accessibility plays in determining enzymatic hydrolysis rates (discussed in section 1.1.6 above). As discussed previously, a number of studies have suggested that cellulose accessibility to cellulase is one of, if not the single, most important factor in determining the rate of enzymatic solubilisation and depolymerisation of the biomass. As cellulose accessibility plays such a key role in enzymatic deconstruction rate, it would be useful if we could develop a better understanding of the roles of these amorphogenesis-inducing proteins play in opening up and disrupting the biomass. This action consequently facilitates subsequent enzymatic attack by the hydrolytic and oxidative enzymes. In the following sections, the various enzymes involved in biomass deconstruction are discussed briefly, followed by a more in-depth discussion on the non-hydrolytic, non-oxidative proteins that have been shown to promote amorphogenesis. 1.2.1 Enzymes involved in lignocellulose deconstruction The enzymatic deconstruction of biomass has traditionally been referred to as the “enzymatic hydrolysis” step of the bioconversion process. However, the relatively recent discovery of enzymes that cleave cellulose by an oxidative rather than a hydrolytic mechanism (Harris et al., 2010; Vaaje-Kolstad et al., 2010), as well as the identification of several proteins that disrupt the biomass without apparent hydrolytic or oxidative activity (Arantes and Saddler, 2010), suggests that the term “enzymatic hydrolysis” is not completely descriptive of the overall process. 34 The enzymes involved in deconstructing biomass include those that specifically break down cellulose either through hydrolytic or oxidative action, those that are involved in breaking down the other components of biomass (hemicellulose, lignin, lignin-carbohydrate complexes), and those that open up the biomass to facilitate the action of these first two groups. In general, biomass-degrading enzymes come in a variety of shapes and sizes, from very small (~8.5kDa) xylanases (Tenkanen et al., 1992) up to very large (1000s of kDa) multimeric biomass-degrading complexes known as cellulosomes (Doi et al., 2003). In general, fungi produce free enzyme systems, whereby individual enzymes with only one or two catalytic domains are secreted, while some bacteria produce multimeric cellulosomes containing a multiplicity of active sites and binding domains held together via a large scaffoldin protein (Reviewed in Lynd et al., 2002). As only fungal enzymes were used throughout the thesis work, the detailed structures and functions of bacterial cellulosomes are not discussed in detail here and can be accessed at (Bayer et al., 1994, 2004). While some of the biomass-degrading enzymes secreted by fungi simply consist of a single catalytic domain (CD), many of the secreted enzymes have a modular structure, containing at least one carbohydrate binding module (CBM) fused to the CD via a flexible, often glycosylated, linker region (Tomme et al., 1988). The primary role of the CBMs is thought to be the facilitated targeting of the CD to the desired polymer, thereby enhancing the local concentration of the enzyme at the substrate surface. However, other potential roles such as biomass disruption and cellulose decrystallization, have also been suggested (Guillén et al., 2010). Although most CBMs target the CD to the specific polymer that the CD is active against, some CBM-CD combinations have mismatched specificities. For example, some CDs that specifically hydrolyze xylan are fused to a cellulose-specific CBM, suggesting that these polymers are very tightly associated in the biomass structure (Ong et al., 1993). 35 1.2.1.1 Hydrolytic cellulases Hydrolytic cellulases are the most-studied of the enzymes involved in lignocellulose deconstruction. These enzymes generally have a modular structure, comprised of a catalytic domain joined via a flexible linker region to at least one carbohydrate binding module (Boraston et al., 2004; Davies and Henrissat, 1995; Gilbert et al., 1990; Hildén and Johansson, 2004; Ong et al., 1989). In the secretome of T. reesei, the most widely-used fungus for industrial cellulolytic enzyme production, a variety of different cellulases have been identified. These include endoglucanases, which cleave cellulose at seemingly random locations within the cellulose chain, and cellobiohydrolases, which bind to chain ends and progressively move down the cellulose chain releasing cellobiose units. Of the endoglucanases, Cel7B and Cel5A (previously known as EGI and EGII, respectively) are the most abundant, making up 5-10% and 1-10%, respectively, of the cellulase mixture secreted from wild-type QM6A T. reesei (Kleywegt et al., 1997; Kolbe and Kubicek, 1990; Markov et al., 2005). Endoglucanases have an open binding cleft which allows these enzymes to bind to and cleave cellulose distal from chain ends (Kleywegt et al., 1997; Nakazawa et al., 2008). These enzymes have been shown to preferentially bind to and hydrolyze the more disordered/amorphous regions of the cellulose, with relatively low activity on crystalline cellulose (Béguin and Aubert, 1994). Alternatively, cellobiohydrolases preferentially hydrolyze the more organized/crystalline regions of the cellulose (Béguin and Aubert, 1994). These enzymes, unlike endoglucanases, have a tunnel-shaped active site which requires cellulose chains to be fed into this tunnel end-wise (Divne et al., 1994; Rouvinen et al., 1990). This active site tunnel means that cellobiohydrolases cannot effectively hydrolyze cellulose from the middle of the chains (although there is some evidence that cellobiohydrolases do have weak endo activity (Ståhlberg et al., 1993), and must instead work from either the reducing or non-reducing end of the chains. In the case of T. reesei, the main cellobiohydrolases are Cel7A, which 36 hydrolyzes from the reducing ends, and Cel6A, which hydrolyzes from the non-reducing ends (Barr et al., 1996; Chanzy and Henrissat, 1985; Divne et al., 1994; Imai et al., 1998). Both of these enzymes then processively cleave off cellobiose units from the single cellulose chain being fed through the active site (Igarashi et al., 2009; Nummi et al., 1983). Under cellulase-inducing conditions, these two enzymes make up approximately 60-70% of the secreted cellulase in wild-type T. reesei, with Cel7A alone making up 40-50% (Sandgren et al., 2005; Shoemaker et al., 1983; Teeri et al., 1987). In addition to the endoglucanases and cellobiohydrolases, T. reesei also secrete β-glucosidase, albeit at much lower levels (Korotkova et al., 2009). Unlike the endoglucanases and cellobiohydrolases, β-glucosidase does not work on the insoluble cellulose, but rather cleaves shorter soluble cello-oligomers, including cellobiose (Halliwell and Griffin, 1973; Shewale, 1982; Sternberg, 1976). This enzyme plays a key role in reducing end-product inhibition, as cellobiose and other short soluble cello-oligomers have been shown to inhibit the hydrolase enzymes that act on the insoluble cellulose (Halliwell and Griffin, 1973). In native T. reesei, β-glucosidase is not expressed at sufficient levels to effectively prevent end-product inhibition, at least for the case of laboratory-condition hydrolysis experiments. For this reason previous versions of commercial cellulase mixtures derived from T. reesei made use of an exogenous source of β-glucosidase, such as from Aspergillus niger (Korotkova et al., 2009; Seidle et al., 2004). However, newer commercial cellulase mixtures now contain enhanced β-glucosidase activity to mitigate this end-product inhibition (Jiang et al., 2011). Although these cellobiohydrolases, endoglucanases, and β-glucosidase enzymes have different active site structures, different substrate targets and different roles in the enzyme mixture, they all cleave cellulose via a hydrolytic mechanism. 37 1.2.1.2 Oxidative and accessory enzymes Up until 2010, it was thought that essentially all enzymatic cellulose cleavage was carried out by hydrolytic enzymes. However, it was recently shown that a new family of enzymes, known as lytic polysaccharide mono-oxygenases (LPMOs), also contribute to the deconstruction of cellulose. These enzymes, which include the cellulose-active AA9 (auxiliary activity family 9, formerly known as GH61) (Harris et al., 2010; Langston et al., 2011; Vaaje-Kolstad et al., 2010) the chitin- and cellulose-active AA10 (formerly known as CBM33) (Forsberg et al., 2011), and the chitin-active AA11 (Hemsworth et al., 2014) have oxidative activity, whereby reactive oxygen species are produced proximal to the surface of the carbohydrates via a redox-active metal atom chelated at the active site (Reviewed in Horn et al., 2012). These oxidative enzymes require a redox-active cofactor such as the enzyme cellobiose dehydrogenase (Langston et al., 2011), compounds such as ascorbate or gallate (Quinlan et al., 2011), or soluble compounds present in the liquor produced during pretreatment of lignocellulosic materials (Hu et al., 2014; Quinlan et al., 2011). AA9 was first discovered as an enhancer of cellulase activity, and has since been shown to target the crystalline regions of cellulose (Beeson et al., 2012; Hu et al., 2014). As endoglucanases tend to have low activity on crystalline cellulose and cellobiohydrolases need free chain ends to act upon (Teeri, 1997), it seems that LPMOs such as AA9 provide the ‘missing activity’ in traditional hydrolytic enzyme mixtures, by cleaving cellulose chains within crystalline regions, thereby freeing up new chain ends for hydrolysis by cellobiohydrolases (Harris et al., 2010; Langston et al., 2011; Vaaje-Kolstad et al., 2010). Until recently, LPMOs were grouped along with ‘non-hydrolytic disruptive proteins’, however, since it is now known that these enzymes are part of a family of oxidative enzymes that can directly catalyze the cleavage of cellulose, it seems that these oxidative enzymes should be grouped separately from the non-hydrolytic, non-oxidative proteins which do not directly cleave cellulose. In addition to the 38 hydrolytic and oxidative enzymes that function to degrade the cellulosic fraction of biomass, there are many other enzymes involved in degrading the other bonds that exist within the complex lignocellulosic matrix. These include enzymes that break down hemicelluloses, lignin, and the interface between lignin and hemicelluloses. For example, a variety of endo-acting and exo-acting hemicellulases (such as xylanases and mannanases) break down the hemicellulose, while phenol oxidases (laccases) and peroxidases (lignin peroxidase and manganese peroxidase) degrade the lignin, and a variety of acetyl esterases and esterases that hydrolyze lignin-linked glycoside bonds, such as coumaric acid esterase and feruloyl acid esterase are involved in deconstructing lignin-carbohydrate complexes (Dashtban, 2009; Tenkanen et al., 1992). While there is broad diversity in the specific substrate targets and also in the structure and mechanism (hydrolytic vs oxidative) of the enzymes involved in biomass deconstruction, these enzymes all have a common need to diffuse into the substrate to reach their particular target bonds within the macromolecular organization of the biomass. The accessibility of these bonds to the enzymes has been shown to be one of the main factors contributing to the recalcitrance of lignocellulosic materials (Arantes and Saddler, 2010). It is therefore particularly interesting that there appears to be a family of proteins involved in opening up and disrupting cellulosic biomass, thereby facilitating the diffusion of all other enzymes to their target bonds within the biomass. 1.2.1.3 Amorphogenesis-inducing proteins In addition to the hydrolytic and oxidative enzymes that are involved in deconstructing biomass, it has been shown that a third family of proteins, which do not have hydrolytic or oxidative activities, are also involved. This family of proteins is fundamentally different from the enzymes discussed earlier because they appear to be capable of opening up and disrupting cellulosic/lignocellulosic biomass without releasing soluble sugars (Arantes and Saddler, 2010). It has been proposed that these proteins 39 are involved in weakening the lignocellulosic matrix without directly cleaving the polymers comprising the biomass. Although these proteins have traditionally been described as “non-hydrolytic disruptive proteins” (Din et al., 1991; Saloheimo et al., 2002), the identification of the oxidative family of deconstructing enzymes necessitates a more implicit descriptor for these proteins, such as “non-hydrolytic/non-oxidative disruptive proteins” or “amorphogenesis-inducing proteins”. The concept that a disruptive component is involved in the disruption of lignocellulosic substrates was first proposed by Reese et al. in 1950. They postulated that the factors involved in lignocellulose degradation could be broken down into the C1 (disruptive) and Cx (hydrolytic) components. This ‘C1-Cx’ theory suggested that the non-hydrolytic C1 component was involved in disrupting/loosening the substrate, thereby increasing the amount of cellulose accessible to the hydrolytic Cx components (Reese et al., 1950). Over the years, this theory has evolved, and the meanings of C1 and Cx have changed (Summarized in (Esteghlalian et al., 2001b). During the development of this theory, both exoglucanases (Wood and McCrae, 1972) and endoglucanases (Wood and McCrae, 1979) have been suggested to play the role of the disruptive component. However, the most recent iteration of this theory suggests that the C1 component is indeed non-hydrolytic (Arantes and Saddler, 2010; Esteghlalian et al., 2001b). This theory differs from the original theory put forward by Reese et al. in that it is now thought that the C1 activity is carried out not solely by a system distinct from the Cx component, but also by the CBMs of lignocellulolytic enzymes (Esteghlalian et al., 2001b). Evidence accumulated since Reese et al. first suggested the involvement of a disruptive component implies that non-hydrolytic proteins and protein domains are capable of disrupting the substrate at the macroscopic, microscopic, and even nanoscopic levels. For the purposes of this work, accessibility has been broken down into three categories, with ‘nanoscopic’ representing the sub-cellulase scale (i.e. <5nm), microscopic representing scales larger than cellulases but invisible to the naked eye (i.e. 5nm - 100µm), and macroscopic representing larger phenomenon visible to the naked eye (i.e. >100µm). 40 1.3 Evidence that amorphogenesis does occur As manifestations of amorphogenesis promoted by non-hydrolytic, non-oxidative proteins have been shown to occur at the macroscopic, microscopic, and nanoscopic levels of biomass organization, evidence for action at each of these scales is discussed below 1.3.1 Macroscopic disruption/fragmentation At the macroscopic level, where individual cells are associated in aggregates visible to the naked eye, disruption of these entities should be apparent by the release of individual fibres from the bulk of the substrate. Techniques for measuring macroscopic disruption have included measurements of particle size, image analysis of aggregate dispersion and measuring the extension of biomass using an extensometer (Jäger et al., 2011; McQueen-Mason et al., 1992). Several families of proteins have been implicated in loosening/disrupting cellulosic and lignocellulosic materials at the macroscopic level (Banka et al., 1998; Chen et al., 2010; Jäger et al., 2011; McQueen-Mason et al., 1992; Quiroz-Castañeda et al., 2011; Saloheimo et al., 2002). Although not technically a protein family, some cellulose-binding CBMs have been suggested to play a role in macroscopic disruption. There are currently (as of July 2014) 69 distinct families of CBM, with binding specificities to a range of insoluble polymers, including starch, chitin, pectin, several hemicelluloses (including xylan, mannan and others) and cellulose (www.cazy.org). These protein domains range in size from less than 40 residues (Family I CBMs) to up to 200 residues (CBM Families 11, 17 and 40). These families are derived from the structural folds of the CBMs. Thus the different Types of CBM (i.e. A, B, or C, described above in Section 1.1.7, where CBMs are broadly classified by binding face architecture) can contain a number of different CBM families containing different fold structure but exhibiting similar binding face architecture. 41 Typically, a single CBM is linked to a corresponding catalytic domain, although multiple CBMs can be found linked to one or more catalytic domains (Bolam et al., 2001; Boraston et al., 2002a; Tomme et al., 1995). Interestingly, CBMs with different specificities have been found linked to a single catalytic domain (e.g. a xylanase catalytic domain linked to a xylan-specific and a cellulose-specific CBM) (McLean et al., 2000). Overall, the primary role of CBMs appears to be in targeting their conjugated catalytic domains to the substrate surface (Hervé et al., 2010). Several cellulose-binding CBMs have been suggested to disrupt cellulosic and/or lignocellulosic substrates at various levels of organization (Din et al., 1991; Gilkes et al., 1993; Lee et al., 2000). In particular, Family 2 CBMs, (which are ~110 residues in length and exhibit cellulose-binding activity, have been isolated from both an exoglucanase/xylanase (Cex)) and an endoglucanase (CenA) from Cellulomonas fimi and have been demonstrated to cause the disaggregation/dispersion of filter paper and to prevent flocculation of bacterial microcrystalline cellulose (Din et al., 1991; Gilkes et al., 1993). As well as the CBMs, the plant expansin protein family has been implicated in the loosening of plant cell walls (Cosgrove 2000). The expansins were first discovered as mediators of acid-induced cell wall expansion in plants (McQueen-Mason et al., 1992), and have since been found to have a slight positive effect on the enzymatic hydrolysis of dilute-acid pretreated poplar (Baker et al., 2000). They are composed of two domains: an inactive glycosyl hydrolase family 45 domain, and a second domain putatively involved in polysaccharide binding (Yennawar et al., 2006). The macroscopic disruptive effects of these proteins are demonstrated by the stretching of plant material and filter paper when treated with expansins and analyzed using an extensometer (Cosgrove and Durachko 1994; Cosgrove 2000; McQueen-Mason et al. 1992). This effect is thought to be caused by a weakening of the hydrogen bonding between cellulose chains and/or between cellulose and other adjacent polysaccharides (Cosgrove 2000). There are two sub-families of expansins: the α-expansins and the β-expansins. The α-expansins have been implicated in acid-induced plant cell growth, while the β-expansins have been 42 found to loosen Type II plant cell walls (those of the grasses and other commelinoid monocots), suggesting that these proteins may act preferentially on the specific hemicelluloses of these plant families (Cosgrove 2000). As well as the bacterial and plant proteins putatively involved in macroscopic substrate disruption, the fungal protein Swollenin from Trichoderma is also thought to play a role in amorphogenesis. This protein was shown to facilitate the macroscopic substrate disruption of Valonia cell wall fragments, where treatment of these fragments with Swollenin resulted in a dispersion/dissaggregation of the cell wall agglomerates (Saloheimo et al., 2002). Additional work using the swollenin-like protein, AfSwo1, from Aspergillus fumigatus on Avicel PH-101 resulted in the size reduction of Avicel particles after incubation with AfSwo1 (Chen et al., 2010). It appeared that AfSwo1 promoted an almost 2-fold reduction in the size of the Avicel particles after a 72 hour incubation (Chen et al., 2010). Although this work did not make use of a protein control, the dependency of the size reduction effect on both temperature and pH suggested that the observed effect was indeed due to the specific activity of AfSwo1. In addition to the work by Chen et al., more recent work using laser diffraction has shown that Swollenin from Kluyveromyces lactis is also capable of reducing the size of filter paper fragments (disaggregation), as well as reducing the particle size of α-cellulose and Avicel (Jäger et al., 2011). Additionally, the Swollenin-related protein, Loosenin from Bjerkandera adusta, has been shown to promote the macroscopic dispersion of filter paper aggregates (Jäger et al., 2011; Quiroz-Castañeda et al., 2011). In addition to Swollenin and Loosenin, a third fungal protein, known as fibril forming protein (FFP) from T. reesei has been shown to enhance macroscopic substrate disruption. The FFP is a small (11.4kDa) protein capable of releasing fibrils from filter paper without any detectable release of reducing sugars (Banka et al., 1998). This protein exhibits pH and temperature optima close to those of cellulase, further supporting its role in substrate disruption. 43 Although there is significant evidence suggesting that these proteins have a macrosopic disruptive effect on cellulosic and/or lignocellulosic substrates, many of these experiments did not make use of a protein control, which would have indicated whether the effects observed in the presence of these proteins were due to their specific disruptive effects, or to a more general protein-mediated disruption/dispersion, such as that potentially induced by the surfactant properties of some proteins (Kerff et al. 2008; Lee et al. 2010; Yang and Wyman 2006). Therefore, except for the case of the expansins, where boiled expansin was used as a negative protein control, it has not yet been definitively proven that non-hydrolytic proteins are capable of causing macroscopic disruption of cellulosic or lignocellulosic substrates. 1.3.2 Microscopic/ultrastructural disruption At the microscopic/ultrastructural level (5nm - 100µm) protein-mediated disruption is evidenced by a loosening of adjacent microfibrils, the release of very small particles from the cell wall, and possibly complete disintegration of sections of the wall. Techniques for assessing microscopic disruption have primarily made use of microscopy techniques, either for directly visualizing disruption of cell walls, or for quantifying microscopic substrate dispersion/fragmentation via microscopy coupled with image analysis software (Din et al., 1991; Jäger et al., 2011; Pinto et al., 2004; Saloheimo et al., 2002). Some of the proteins described above as having macroscopic substrate loosening and/or weakening properties have also been shown to promote microscopic disruption of cellulosic and lignocellulosic substrates. For example, the C. fimi Family 2 CBM from CenA has been shown to disrupt the surface of ramie and cotton fibres (Din et al., 1991; Tomme et al., 1995). Additionally, microscopy studies on the effect of swollenin on cotton fibres suggest that it is able to swell cotton fibres without producing detectable amounts of reducing sugars (Saloheimo et al., 2002). Scanning Electron 44 Microscopy (SEM) has also been used to observe roughening of the surface of cotton fibres promoted by CBM2a from Cellulomonas fimi (Din et al., 1991, 1994), disaggregation of Whatman CF-11 cellulose fibres by CBM1 from T. reesei (Pinto et al., 2004), as well as fibrillation of filter paper fibres (Jäger et al., 2011). Finally, although the observed effect of expansin treatment (loosening and elongation of the plant material or filter paper) is itself a macroscopic observation, it is likely that the expansins are also promoting disruption of the substrate at the microscopic level through effects such as disrupting microfibril-microfibril interactions (Cosgrove 2000). 1.3.3 Nanoscopic disruption Substrate disruption at the nanoscopic level (<5nm) has been demonstrated through direct visualization (using AFM) of nano-scale pitting on the surface of microfibrils (Lee et al., 2000). Additionally, it has been suggested that crystallinity measurements could provide insight into nano-scale substrate disruption (Wang et al., 2008). Specifically, the ~40 residue cellulose-binding Family 1 CBMs have been implicated in this event. These CBMs are composed of a single, highly thermostable, β-sandwich and are found almost exclusively in fungi, with the only exception being an algal protein exhibiting four adjacent Family 1 CBM folds (www.cazy.org) (Kraulis et al., 1989). Evidence to support the role of these CBMs in disrupting the nanoscopic molecular architecture has been provided by Lee et al (2000) who produced a catalytically inactive Cel7A from T. reesei by treating the isolated enzyme with hexachloropalladate (Lassig et al., 1995; Shultz et al., 1995). These workers postulated that, by catalytically inactivating the enzyme, any observable effects of the protein could be attributed to the CBM. Subsequent AFM visualization of cotton fibres incubated with this inactive Cel7A revealed pitting of the cellulose surface, without detectable release of sugars. This suggested that Family 1 CBMs were capable of directly disrupting the surface of microfibrils (Lee et al., 2000). Further studies on cotton fibres, this time using the proteolytically cleaved Family 1 CBM from 45 the Cel7A of Trichoderma pseudokoningii S-38 and FTIR, that this protein was capable of disrupting the strong hydrogen bonds present in crystalline cellulose (Wang et al., 2008). A molecular dynamics simulation of this interaction suggested that the CBM reduces the hydrogen bonding within the cellulose by causing a rotation of the outer (CBM-bound) cellulose chain around its axis. This rotation would twist the outer cellulose chain out of the optimal position for hydrogen bonding, thus weakening the hydrogen bonds and rendering it more susceptible to release from the crystalline bundle (Wang et al., 2008). Other work involving the modeling of CBM1 binding to the surface of crystalline cellulose has suggested that these proteins bind following an induced fit model, whereby an internal tyrosine (Y13) folds out of the protein and aligns with the planar cellulose surface (Nimlos et al., 2007). This modeling is especially interesting as it suggests that a CBM binding at the site of a broken chain within a crystalline cellulose bundle is capable of lifting the reducing end of the broken chain away from the cellulose surface, thereby allowing up to four water molecules to diffuse under the outermost cellulose chain (Nimlos et al., 2007). However, a more recent paper has demonstrated that, at least with currently available enzyme cocktails, little change occurs to the nanoscopic order/organization during the course of enzymatic deconstruction (Chauve et al., 2013). This suggests that enzymatic disruption of the bulk of the nanoscopic structural features within the biomass cannot be achieved rapidly. This in turn implies that if effective nanoscopic amorphogenesis can be achieved, there could be significant improvements in enzymatic hydrolyzability, as current enzyme mixtures do not appear to be capable of penetrating into and disrupting the highly compact nanoscopic structural organization of cellulose. Overall, it is possible that the effects of the non-hydrolytic proteins described above on the various levels of molecular organization within lignocellulosic biomass are all manifestations of a more general function of these proteins. Nanoscopic disruption of the microfibrils could disrupt the microfibril-microfbril interactions at the microscopic level, which, in turn, could lead to a disruption of 46 inter-fibre associations, thus leading to macroscopic fragmentation and dispersion of the substrate. Alternatively, it is conceivable that different types of non-hydrolytic proteins could exhibit specific effects on the substrate. For example, the expansins may act only at the microscopic and macroscopic levels, by causing a loosening of adjacent microfibrils and/or cells, while Family 1 CBMs may primarily act at the nanoscopic level, by disrupting hydrogen bonding and crystallinity within individual microfibrils. To summarize, the various amorphogenesis-inducing proteins identified to date are listed in Table 1, and the disruptive effects of amorphogenesis-inducing proteins at the macroscopic, microscopic and nanoscopic levels are represented in Figure 1, detailing macroscopic fibre dispersion and disaggregation, microscopic cell wall loosening/roughening, and nanoscopic pitting/decrystallization. Table 1: Amorphogenesis-inducing proteins identified to date, along with their putative function and level of disruption. Proteins Organisms Putative Function Level of Disruption References CBMs Bactera, Fungi Fibre pitting/roughening, Nanoscopic, Gilkes et al. 1991, small particle release, Microscopic Din et al. 1991, 1994, localized reduction in crystallinity. Gao et al. 2001, Wang et al. 2008 Swollenin, Loosenin Fungi Cotton fibre swelling, Microscopic, Jäger et al. 2011, dispersion of cellulose aggregates. Macroscopic Quiroz-Castañeda et al. 2011, Saloheimo et al. 2002 Expansins Plants Loosening of plant cell walls, Microscopic, Baker et al. 2000, dispersion of filter paper Macroscopic Cosgrove 2000 Expansin-like proteins Bacteria Loosening of filter paper, Microscopic, Kerff et al. 2008, dispersion of cellulose aggregates. Macroscopic Lee et al. 2010 Fibril-forming Protein Fungi Fibril release from filter paper Microscopic Banka et al. 1998 47 Figure 1: Model of protein-induced amorphogenesis of biomass occurring at the nanoscopic (A1), microscopic (B1), and macroscopic (C1) levels. Manifestations include the nanoscopic pitting of microfibrils (A2), microscopic dispersion, swelling, weakening and loosening of cell walls (B2), and the dispersion and disaggregation of macroscopic associations of fibre cells (C2). 48 1.4 Focus on Swollenin One of the better-studied amorphogenesis-inducing proteins, and the protein that was most extensively studied in this thesis, is the fungal protein, Swollenin (Saloheimo et al., 2002). Swollenin was originally identified as a protein of interest when screening the T. reesei genome for enzymes with sequence similarity to the cell-wall loosening plant enzymes known as the expansins (Saloheimo et al., 2002). As mentioned earlier, this protein has previously been shown to cause macroscopic dispersion of Valonia and filter paper, as well as the microscopic swelling of cotton fibres and roughening of filter paper fibres (Jäger et al., 2011; Saloheimo et al., 2002). The ~475-residue protein is composed of an N-terminal 36-residue Family 1 CBM connected through a very long (~50aa) O-glycosylated linker region to an expansin-like domain (Brotman et al., 2008; Saloheimo et al., 2002). Although the crystal structure of Swollenin has not yet been solved, Swollenin has ~25% identity over ~200 amino acids with an expansin domain (Yennawar et al., 2006). This expansin domain is composed of two tightly associated domains (D1 and D2), which have a carbohydrate-binding face spanning across the two domains. In addition, a secondary conserved binding site is present on the underside of D2 (Yennawar et al., 2006). Additionally, Swollenin is known to contain a region with homology to FNIII repeats, a protein sequence known to be extremely flexible, with the ability to unfold and refold repeatedly (Hsin et al., 2011; Saloheimo et al., 2002). Thus the structure of Swollenin includes multiple binding sites connected together via a long flexible linker region and a putatively flexible FNIII-like repeat. This highly-flexible structure fusing multiple carbohydrate binding sites, coupled with the observed activities of Swollenin (weakening, loosening and opening up cellulosic substrates), along with its homology to expansins which have been suggested to disrupt hydrogen bonding (Cosgrove 2000), has led to the theory that Swollenin functions through the weakening of hydrogen bonding interactions between cellulose chains/fibrils, or between cellulose and other cell wall components (Saloheimo et al., 2002). 49 Regarding the potential targets of Swollenin in nature, this protein has greater sequence similarity with the plant β-expansins, which act predominantly on Type II cell walls (grasses and commelinoid monocots) than to the α-expansins involved in acid-induced plant cell growth (Cosgrove 2000; Saloheimo et al. 2002). This suggests that Swollenin may be more active on grasses and commelinoid monocots such as corn stover than on woody biomass. The experiments performed to date to assess the disruptive effects of Swollenin and other amorphogenesis-inducing proteins have made use of a number of different techniques, which together provide a general picture of the role of these proteins in biomass deconstruction. However, attempts to elucidate potential synergistic interactions between these amorphogenesis-inducing proteins and cellulolytic enzymes have met with limited success. These experiments are discussed below, followed by a summary of the drawbacks of the techniques used to date for quantifying amorphogenesis. 1.5 Enhancement of biomass deconstruction by amorphogenesis-inducing proteins An additional technique that has been used to observe amorphogenesis has been the indirect approach of measuring changes in the hydrolyzability of the substrate after treatment with amorphogenesis-inducing proteins. This approach has been used to demonstrate enhanced hydrolysis of a range of substrates after treatment with several different disruptive proteins. Several expansins and expansin-like proteins have been shown to enhance hydrolysis of model cellulosic substrates, as well as two cases where improved hydrolysis of a pretreated lignocellulosic material was observed (Baker et al. 2000; Chen et al. 2010; Kim et al. 2009; Lee et al. 2010; Wang et al. 2010; Zhou et al. 2011). Specifically, the bacterial expansin-like proteins HcEXLX2 from Hahella chejuensis and BsEXLX1 from Bacillus subtilis were found to promote hydrolysis of filter paper.However, this promotion was only observed at extremely low enzyme loadings (0.06 Filter Paper Units, FPU) and conversion levels (Kim et al. 2009; Lee et al. 2010). Additionally, supplementation of dilute-acid 50 pretreated poplar with β-expansin from Zea mays pollen during hydrolysis resulted in slight (<10%) improvements in hydrolysis in the range of 55-80% conversion, but not at conversion levels outside this range (Baker et al., 2000). In addition to the expansins and bacterial expansin-like proteins, several fungal proteins with sequence homology to the expansins have been shown to enhance the hydrolyzability of cellulosic and lignocellulosic biomass. For example, four recent reports on the effects of three different Swollenin proteins on hydrolysis rates suggest that these proteins are capable of acting synergistically with cellulase preparations to enhance hydrolysis (Chen et al. 2010; Jäger et al. 2011; Wang et al. 2010; Zhou et al. 2011). Wang et al. demonstrated that Swo1 from T. reesei expressed in Aspergillus oryzae was capable of promoting an 82% increase in hydrolysis of filter paper by cellulase when compared to a BSA control, albeit at very low enzyme loadings (0.06 FPU) (Wang et al., 2010). A positive effect on hydrolysis was also observed when AfSwo1, as well as recombinant Swo2 from T. pseudokoningii S-38 expressed in A. niger, were used to supplement cellulases during the hydrolysis of Avicel PH-101. However, this work did not make use of an effective protein control (Chen et al., 2010; Zhou et al., 2011). Additionally, pretreatment with Swollenin from Kluyveromyces lactis was shown to enhance hydrolysis of filter paper, α-cellulose, and Avicel PH101, but not Sigmacell 101, at a cellulase loading of 0.28 FPU (Jäger et al., 2011). The fungal Swollenin-like protein, Loosenin, has also been shown to enhance the hydrolyzability (at low conversion levels) of both cotton fibres and the natural lignocellulosic material Agave tequilana relative to protein controls (Quiroz-Castañeda et al., 2011). In addition to these expansins and expansin-related proteins, some CBMs have also been implicated in improving hydrolysis. For example, a slight positive effect on enzymatic hydrolysis was observed on filter paper and microcrystalline cellulose in the presence of CBM1 from T. reesei (Lemos et al., 2003). However, this study did not make use of a protein control. The importance of a protein 51 control for studies involving CBM-mediated disruption was demonstrated by Lee et al. (2010), who showed that addition of BSA as a protein control promoted enzymatic hydrolysis to a greater extent than did CBM3 from Clostridium thermocellum. This suggested that there is a potential weak enhancement of hydrolysis induced by non-specific effects of adding protein to the reaction. Thus a protein control must always be included to ensure that the observed effect is indeed due to specific effects of the CBM, rather than the indirect effects of simply adding inactive protein (Lee et al., 2010; Lemos et al., 2003). Finally, CBM2a was found to enhance the hydrolysis of cotton by an isolated cellulase catalytic domain (without a CBM). Again this study did not make use of a protein control and subsequent experiments using CBM2a on Avicel PH-101 were actually found to cause a decrease in the hydrolyzability of the substrate (Esteghlalian et al., 2001b). It was hypothesized that this decrease in hydrolysis was due either to incomplete protein removal after CBM2a treatment leading to blocking of cellulase binding sites by the CBM, or to residual proteinase activity after attempted removal of CBM2a from the substrate by proteinase. Overall, the studies that have tried to demonstrate the enhancement of enzymatic hydrolyzability by amorphogenesis-inducing proteins have met with limited success. Specifically, most improvements appear to occur only at very low enzyme loadings and at conversion levels which are insignificant for potential industrial sugar production platforms. The exception to this appears to be the work of Baker et al. (2000) who demonstrated a slight increase in hydrolysis yields of dilute-acid pretreated poplar between the conversion levels of 55-80%. Despite the minimal enhancement of hydrolysis at commercially relevant enzyme loadings and conversion levels, the ability of these proteins to enhance hydrolysis at low enzyme loadings does support the contribution of these amorphogenesis-52 inducing proteins to cellulose hydrolysis. However, this past work also indicates that a more thorough understanding of their mechanisms might improve conversion efficiencies at higher enzyme loadings. It is possible that previous attempts to demonstrate hydrolysis boosting effects of amorphogenesis-inducing proteins were hindered by issues with experimental design. For example, it is likely that some amorphogenesis-inducing proteins have activities which are already contained within the commercial cellulase mixtures employed for synergism experiments. This would result in the masking of potential synergistic interactions by the amorphogenesis-inducing proteins already present within the cellulase mixture. For example, Swollenin is known to be expressed at relatively low levels by T. reesei under cellulase-inducing conditions (Saloheimo et al., 2002), suggesting that the activity of Swollenin already present within the cellulase broth may mask any additional amorphogenesis effects after exogenous Swollenin addition. Conversely, it is possible that certain amorphogenesis-inducing proteins require the enzymatic activities of additional enzymes in order to enhance hydrolysis. For example, Baker et al. (2000) demonstrated that a β-expansin only exhibited synergistic interactions in the presence of the full cellulolytic mixture from T. reesei, and did not exhibit synergy with a reconstituted enzyme mixture containing only a single cellobiohydrolase and single endoglucanase. This was taken to mean that there is an additional factor present within the full cellulase mixture that is required for the efficient activity of the β-expansin (Baker et al., 2000). While these two arguments (the masking effect and the lack of required additional enzyme activity) appear contradictory, little is currently known about the interactions between amorphogenesis-inducing proteins and cellulases, and thus either or both of these scenarios could be occurring during synergism experiments. The complex interactions between amorphogenesis-inducing proteins and cellulase systems could thus be a key factor that limits the efficient exploitation of amorphogenesis-inducing enzymes. 53 In summary, assessing the synergistic interactions between amorphogenesis-inducing proteins and cellulases has proved challenging for a number of reasons. However, by developing a better understanding of the fundamental mechanisms by which these proteins disrupt cellulosic biomass and by carefully designing experiments such that the complex interactions between amorphogenesis-inducing proteins and other biomass deconstructing enzymes can be teased out, it should be possible to overcome these challenges and develop enhanced cellulolytic enzyme mixtures exploiting the amorphogenesis-inducing abilities of these proteins. To date, all previous work on amorphogenesis-inducing proteins have made use of techniques that have provided very little quantitative data on the specific effects of these disruptive proteins on the biomass. In order to advance our knowledge of these proteins, it is necessary to first understand the drawbacks of current techniques, and work toward developing and applying new techniques for quantifying the effects of amorphogenesis. Some of the drawbacks and strengths of current techniques are discussed in the following section. 1.6 Current techniques used for measuring amorphogenesis Some of the techniques applied to measuring macroscopic substrate disruption include measuring particle size, image analysis of aggregate dispersion, and measuring the extension of biomass using an extensometer. For measuring microscopic disruption, the techniques have been predominantly microscopy-based, including direct visualization of cell wall disruption (Din et al., 1991; Jäger et al., 2011; Pinto et al., 2004; Saloheimo et al., 2002; Tomme et al., 1995), or have involved the application of image analysis software to measure microscopic dispersion/fragmentation of the substrate (Jäger et al., 2011). At the nanoscopic level, disruption of microfibril surfaces has been observed by AFM (Lee et al., 2000), while crystallinity measurements have been used to quantify changes in the cellulose crystal structure after amorphogenesis (Gao et al. 2001; Jäger et al. 2011; Wang et al. 2008). Finally, 54 enhancement of substrate hydrolyzability has been used as an indirect measure of substrate disruption (Baker et al., 2000; Chen et al., 2010; Kim et al., 2009; Lee et al., 2010; Wang et al., 2010; Zhou et al., 2011). Although this range of complementary techniques has provided a relatively good idea of the general effects of amorphogenesis, there are a number of drawbacks to the techniques applied to date. Essentially, all of the observations made via microscopy are qualitative observations (semi-quantitative at best, if image analysis software is employed). Additionally, techniques such as measuring particle size and crystallinity focus on quantifying only a single possible aspect of amorphogenesis. For example, it is possible that amorphogenesis primarily occurs through effects such as delamination or fibrillation, where relatively large, intact fragments, still containing crystalline regions, are released from the bulk of the substrate. If this were the case, the disruptive proteins would cause an increase in the accessibility of the substrate, without affecting the crystallinity. This hypothesis is supported by Pinto et al. (2004) who observed an increase in cellulose accessibility without a concomitant decrease in crystallinity after CBM-induced disruption. Alternatively, or simultaneously, amorphogenesis could occur as a product of substrate swelling or loosening at the interactions between microfibrils, which would result in the overall weakening of the cell wall, leaving the crystalline cores of the microfibrils relatively untouched. Thus it is possible that any changes in crystallinity of the substrate during amorphogenesis occur only as a relatively minor secondary effect of the more general process of amorphogenesis. Attempts to quantify amorphogenesis by measuring the crystallinity of the substrate have met with conflicting and inconclusive results. For example, while Fourier Transform Infrared Spectroscopy (FTIR) measurements of the effects of CBM1 from T. pseudokoningii S-38 on cotton fibres suggest a CBM-induced reduction in crystallinity (Wang et al., 2008), the effect of the highly similar CBM1 from T. reesei on Whatman CF11 cellulose fibres, measured using X-ray diffraction (XRD), appeared not to 55 influence the crystallinity of the substrate (Pinto et al., 2004). Additionally, the bacterial CBM3a from Clostridium Cellulovorans has been shown to cause a reduction in the crystallinity of cotton fibres using both FTIR and XRD (Ciolacu et al., 2010). Finally, a recombinant Swollenin, Swo2 from T. pseudokoningii S-38 was actually found to cause an increase in the crystallinity of Avicel PH-101 (Zhou et al., 2011). While these results suggest that very specific combinations of disruptive protein and substrate result in a reduction in substrate crystallinity, it seems more likely that these contradictory results stem from issues surrounding the use of crystallinity measurements for quantifying amorphogenesis (discussed in more detail in Section 1.1.3, above). Enhancement of substrate hydrolyzability has also been suggested as a technique for measuring amorphogenesis. Several groups of researchers have demonstrated enhanced hydrolysis of cellulosic and lignocellulosic substrates after treatment with non-hydrolytic disruptive proteins (See Section 1.5). While this large body of evidence suggests that disruptive proteins are indeed capable of enhancing the hydrolyzability of both model and native cellulosic substrates, this technique uses an indirect approach for quantifying substrate amorphogenesis and could therefore lead to misinterpretation of the results. For example, it is possible that the β-expansin disruptive protein used to supplement enzymatic hydrolysis of dilute acid pretreated poplar is acting to block lignin rather than to promote amorphogenesis and increase accessibility, thereby preventing non-productive adsorption of the cellulases to the lignin. This putative lignin-blocking effect could partly explain the increased hydrolysis rate observed with β-expansin supplementation (Baker et al., 2000). Additionally, the presence of contradictory results in the literature when the same disruptive protein (CBM2a) was used to test for hydrolysis enhancement by different research groups (Discussed in Section 1.5) places further doubt on the suitability of using enhancement of substrate hydrolyzability as a tool for quantifying amorphogenesis. 56 As mentioned earlier, one factor hindering the rapid development of an in-depth understanding of the mechanisms of action of proteins such as Swollenin is the end-products that these enzymes produce. In the case of enzymes which directly cleave the carbohydrates within lignocellulosic biomass (such as hydrolases) the end products are soluble sugars which are relatively easy to quantify. However, proteins such as Swollenin do not appear to release soluble sugars, making the accurate quantification of their effects difficult, which in turn impedes efforts to perform side-by-side comparisons between various disruptive proteins, or to demonstrate how well a specific disruptive protein works on a particular biomass substrate. This is particularly important when developing new enzyme cocktails, where direct comparisons between enzymes must be performed in order to fine-tune enzyme mixtures to develop the most efficient enzyme combinations for enzymatic biomass deconstruction. Overall, the techniques applied to date for measuring protein-induced amorphogenesis have provided a general picture of the process of amorphogenesis (i.e. the opening up, swelling, and dispersion of the biomass). However, the lack of a technique capable of accurately and quantitatively describing the various manifestations of amorphogenesis has hindered the rapid development of our understanding of the roles and relative effectiveness of the various amorphogenesis-inducing proteins. It has also delayed their effective exploitation in commercial enzyme mixtures for biomass deconstruction. In the following section, a variety of different techniques which could potentially provide valuable insights into the process of amorphogenesis are described. 1.7 Techniques for measuring accessibility with potential for quantifying amorphogenesis While the various putative effects of amorphogenesis, such as delamination, fibrillation, swelling, loosening, roughening, pitting, weakening, or decrystallization of the substrate can all be thought of as distinct processes, these phenomena all share a common theme. They result in changes in the accessibility and surface morphology of the substrate. It is therefore important that any technique 57 for quantifying amorphogenesis be capable of recognizing subtle changes in these substrate characteristics. Several methods with potential for quantifying amorphogenesis have already been developed by the pulp and paper industry. These include measuring characteristics such as the water retention value (WRV), mean fibre size, Nitrogen adsorption, and Mercury porosimetry. Alternatively, techniques such as solute exclusion, differential scanning calorimetry (DSC), time-domain nuclear magnetic resonance (TD-NMR), Simons’ Staining, particle scattering analysis and protein adsorption can also be used. 1.7.1 Water retention value The centrifugal WRV is one of the most widely used techniques for quantifying fibre swelling in pulp and paper (Chandra et al., 2008b; Hopner et al., 1955). Calculating the WRV of a sample involves placing a known mass of wetted substrate on a mesh screen, and centrifuging for a set time at a designated rpm. The moisture content of the substrate is then calculated. This technique was introduced by Hopner et al. and is an indirect indication of the degree of swelling of the sample. If the sample is swollen, more polar groups will be exposed to the water, and more water will be retained (Hopner et al., 1955). However, this technique has significant limitations. These include the heavy dependence of the calculated WRVs on test conditions combined with its inaccuracy for analyzing highly swollen pulps. Additionally, the nature of the protocol introduces inherent human error, which prevents the application of this technique for quantifying minor subtle changes in the degree of swelling. This technique is therefore not likely to be capable of quantifying the subtle changes in surface morphology of the substrate induced by non-hydrolytic disruptive proteins. 58 1.7.2 Fibre quality analyzer (FQA) FQA is a technique that can be used to quantify macroscopic properties of cellulosic or lignocellulosic fibres. Briefly, dilute fibre samples suspended in water are passed through a flow cell, where images are captured and analyzed using image analysis software. The length and width of the fibres passing through can then be determined (Robertson et al., 1999). While this technique is simple and easy to use, the data generated only provide information on the macroscopic properties of the fibres. Thus, while FQA could potentially be used to detect macroscopic swelling or fragmentation of the substrate induced by non-hydrolytic disruptive proteins, it is unlikely that it would be capable of detecting such putative amorphogenesis-induced effects as pitting or roughening of the fibres. This is particularly important because pitting and roughening of the fibre surface would greatly enhance the amount of enzyme-accessible surface area (a factor known to play a significant role in determining the efficiency of enzymatic deconstruction), without necessarily altering the macroscopic fibre dimensions. 1.7.3 Nitrogen adsorption and mercury porosimetry Nitrogen adsorption and mercury porosimetry are two similar techniques involving quantification of the amount of Nitrogen or Mercury interacting with the substrate over a range of pressures. Using the Nitrogen adsorption method, dried cellulosic or lignocellulosic samples are exposed to N2; the amount of N2 adsorbed to the substrate gives an indication of the total surface area at the sub-nanometer level (N2 has a Van der Waals radius of ~0.3nm), accounting for both the internal (pores) and external surface areas (Beardmore et al., 1980; Chandra et al., 2008b; Gharpuray et al., 1983; Haselton, 1954). Mercury porosimetry on the other hand solely provides an indication of the internal surface area. This technique involves submerging a dried cellulosic or lignocellulosic sample in Mercury. The pressure is then increased, pushing the Mercury into smaller and smaller pores within the substrate. Because more pressure is required to force the Mercury into smaller pores, measuring the 59 amount of mercury adsorbed to the substrate over a range of pressures can give the pore size distribution of the substrate (Chandra et al. 2008b; Gregg and Sing 1985; Keller and Staudt 2005). One of the main drawbacks to using either of these methods is the requirement for sample drying prior to performing the measurements, which can lead to collapse of pores and hornification of the substrate (Luo and Zhu, 2011). These techniques are therefore unlikely to be useful in quantifying amorphogenesis induced by non-hydrolytic disruptive proteins, as it is probable that sample drying would mask any increase in pore volume/surface area induced by these proteins. 1.7.4 Solute exclusion A more accurate technique for quantifying the accessibility and surface morphology of the substrate is the solute exclusion method (Aggebrandt and Samuelson, 1964; Eriksson et al., 1991; Stone and Scallan, 1968). Application of this technique can be used to measure the total pore volume and pore distribution of pretreated substrates (Grethlein, 1985; Weimer and Weston, 1985). This technique involves exposing the cellulosic samples to a series of solutions containing known concentrations of a range of solutes with narrowly distributed molecular weights, such as a series of polyethylene glycol or dextran chains (Aggebrandt and Samuelson, 1964). Essentially, this technique works by quantifying the pore volume accessible to a solute of a known size; this can be measured by incubating the impregnation solution with the cellulose and determining the reduction in concentration of the solute in the solution. While the solute exclusion method has been widely used for analyzing the pore volume and distribution of cellulosic and lignocellulosic substrates, some drawbacks to this technique include the significant time investment required to generate reproducible results, as well as the inability of this technique to accurately quantify pores with irregular shapes. Finally, the major setback to using this technique for the quantification of amorphogenesis is that it is capable only of quantifying the internal surface area of the substrate, and does not give a value for the total accessible surface area. Although 60 manifestations of amorphogenesis such as swelling, loosening, roughening or pitting might result in the formation of new pores, which could then be quantified by solute exclusion, if amorphogenesis resulted in the fibrillation or delamination of the substrate, it is conceivable that the accessible pore volume would remain relatively constant, even as the total accessible surface area increased. 1.7.5 Differential scanning calorimetry and time-domain nuclear magnetic resonance Other techniques for analyzing pore volume and distribution within cellulosic and lignocellulosic materials include differential scanning calorimetry (DSC) (Nakamura et al., 1981; Nelson, 1977) and nuclear magnetic resonance (NMR) (Felby et al., 2008). These techniques take advantage of the unique properties of water observed within the hydrated micropores of the substrate. Water within the substrate can be categorized into three distinct classes: primary bound water, secondary bound water, and free/bulk water (Felby et al., 2008; Weise et al., 1996). Primary bound water is directly bound to the substrate through both capillary forces as well as hydrogen bonding to hydroxyl groups on the cellulose, hemicelluloses and lignin. Secondary bound water is composed of water molecules held in place through hydrogen bonding to the underlying primary bound water. Primary and secondary bound water is therefore found in close proximity to the substrate surface, and particularly within the micropores of the substrate, where the limited space constrains the movement of the water molecules. Finally, free/bulk water is found in the large pores, as well as in the inter-fibre and intra-fibre (luminal) spaces. The volume of water within each class therefore depends on the size and distribution of the pores within the substrate (Felby et al., 2008). Due to the constraints on movement of the primary bound water and the first 1-2 monolayers of the secondary bound water imposed by their interactions with the substrate, the water molecules within these fractions are incapable of reorienting to form the crystalline ice lattice, and therefore do not freeze. This water can be referred to as non-freezing bound water (Hartley et al., 1992; Nakamura et 61 al., 1981). Additionally, the layers of water slightly more distant from the substrate are also spatially constrained by the hydrogen bonding network propagating from the substrate surface. These constraints produce an energy barrier to reorienting into the ice lattice, which in turn depresses the melting point of the water by about 2° C (Weise et al., 1996). DSC exploits these unique characteristics of water at the water-substrate interface by quantifying the volume undergoing the phase transition from ice to liquid at a given temperature as a frozen sample is heated. Since this phase transition is an endothermic process, it requires more energy input to heat the sample compared to an inert reference material. In heat-flux DSC, the temperatures of the sample and reference material are kept exactly equal as they are heated. The increased energy input required to heat the sample compared to the reference material is measured as a function of time and temperature. Alternatively, power-compensation DSC can be used, where the energy input to the sample and reference material is kept constant, and the difference in temperature of the sample compared to the reference material is recorded. By accurately measuring the difference in energy input (heat flux DSC) or temperature (power-compensation DSC) between the sample and the reference material, these techniques can be used to quantify the volume of water undergoing the phase transition over a range of temperatures (Haines et al., 1998). Water that changes from ice to liquid at 0 °C is the free/bulk water, while water undergoing the phase transition at a lower temperature, usually between 0 °C and -2 °C, is the freezing bound water (Weise et al., 1996). Calculating the difference between the total volume of water undergoing the phase transition (i.e. the sum of the bulk/free water and the freezing bound water) and the total volume of water within the sample provides a measurement of the volume of non-freezing bound water. Since the volume of water in each of these classes is directly related to the pore structure of the substrate, use of DSC provides an accurate measurement of this substrate characteristic. 62 A second technique that can be used for quantifying the volumes of primary bound, secondary bound and free/bulk water within the substrate is time-domain nuclear magnetic resonance (TD-NMR) (Froix and Nelson, 1975). This technique can be used to analyze the spin-spin (T2) relaxation times of the hydrogen nuclei within water molecules. This relaxation time is dependent on how constrained the movements of the hydrogen nuclei are, such that the hydrogen nuclei of water molecules involved in tight binding or enclosed within small compartments have shorter relaxation times than those of the free/bulk water (Felby et al., 2008; Froix and Nelson, 1975). TD-NMR, like DSC, can therefore be used to quantify the volume of water within the primary bound, secondary bound, and free/bulk classes, which in turn is indicative of the pore structure of the substrate. Techniques such as DSC and TD-NMR are thus good candidates for being able to quantify the effects of amorphogenesis, as they appear to be able to quantify accessibility at various levels of structural organization. For example, primary bound water is indicative of the nanoscopic accessibility, while secondary bound water might represent the porosity of the sample. One issue with this technique is that it is incapable of differentiating between water constrained by cellulose (representing cellulose accessibility) and water constrained by other compounds within the biomass (i.e. lignin). Thus these techniques will likely be incapable of providing detailed information on the changes of cellulose accessibility, but will provide information on the bulk accessibility of the total surface of the biomass. Because cellulose accessibility, rather than bulk accessibility, is likely the key determinant of enzymatic hydrolyzability, DSC and TD-NMR may not accurately reflect the true hydrolyzability of a given biomass substrate. Despite this, application of DSC and TD-NMR should be considered as a promising technique for quantifying protein-induced amorphogenesis. 63 1.7.6 Particle scattering Particle scattering techniques, such as small angle X-ray scattering (SAXS) and small angle neutron scattering (SANS) have been used to quantify regions of disordered cellulose within microfibrils by analyzing the scattering density contrast between crystalline and amorphous cellulose (Cheng et al., 2011). These two techniques can be used to measure structural features at the 1-100 nm range (Cheng et al., 2011). Of these techniques, SANS has the greater potential for application to quantifying amorphogenesis, as the neutron scattering density contrast can be greatly enhanced by deuterating the substrate (Nishiyama et al., 2003), and SANS is more penetrating than SAXS, allowing for measurement of structural features buried within the biomass (Cheng et al., 2011). SANS experiments are carried out by replacing the labile hydrogen atoms within the disordered, water-accessible regions of cellulose with deuterium using vapour exchange with heavy water. Because hydrogen and deuterium scatter neutrons in fundamentally different ways, this specific deuteration of the water-accessible regions of cellulose can then be quantified (Fischer et al., 1978). The SANS technique has been used to identify small disordered regions periodically distributed along the microfibrils of higher plants (Nishiyama et al., 2003). This technique has great potential for quantifying subtle changes in the amount and distribution of water-accessible cellulose within cellulosic or lignocellulosic substrates after incubation with amorphogenesis-inducing proteins. However the usefulness of this technique will likely be limited due to the small number of facilities capable of performing SANS experiments, and the high cost of operating this equipment. 1.7.7 Simons’ stain One technique capable of quantifying changes in the overall surface morphology of the substrate is the Simon’s Stain method (Chandra et al., 2008a; Yu and Atalla, 1998). This technique makes use of a high molecular weight direct orange dye with strong affinity for cellulose, and a low molecular 64 weight direct blue dye with weaker affinity for cellulose. Upon incubation with the substrate, the blue dye penetrates into the smaller pores where it binds weakly with the cellulose. The larger orange dye is then added to the substrate, where it binds to the surface cellulose, as well as within the larger pores. Because the orange dye has a higher affinity for cellulose than the blue dye, binding of the orange dye to the substrate displaces the blue dye, leaving the blue dye only within the smaller pores of the substrate inaccessible to the larger orange dye. The ratio of the adsorbed orange dye to the blue dye provides an indication of substrate porosity, while total orange dye binding gives an idea of the accessible surface area of cellulose (Chandra et al., 2008a; Yu and Atalla, 1998). While it has been shown that the ratio of orange dye to blue dye can be used as a predictor of enzymatic hydrolysis extent (Arantes and Saddler, 2011), there are some problems associated with using the Simons’ Stain method for quantifying amorphogenesis. First, because the orange dye consists of a heterogeneous mixture of dyes of different sizes, this technique does not accurately represent the true pore size distribution of the substrate, and instead categorizes all substrate pores into just two size classes (those containing the blue dye, and those where the orange dye mixture has replaced the blue dye). However, refining the Simons’ Stain technique by fractionating the orange dye has been carried out to provide a more homogeneous dye, alleviating this concern (Esteghlalian et al., 2001a). Additionally, more recent work has employed adsorption of the orange dye alone, which is approximately the same size as cellulolytic enzymes, as a probe for cellulose accessible to cellulases, regardless of biomass porosity (Chandra et al., 2008a). This modified technique has also been shown to accurately predict enzymatic hydrolyzability (Chandra et al., 2008a). When considering the enzymatic hydrolysis of cellulose, it is important to consider the fact that cellulases recognize fine substructures within the cellulosic architecture (McLean et al., 2002). Therefore, although Simons’ Stain appears to be capable of providing a relatively accurate depiction of 65 the amount of cellulose surface area accessible to the dye, these dyes likely recognize the substrate in a less discriminatory fashion than the cellulases. It is therefore likely that measurements of cellulose accessible to cellulase based on the Simons’ Staining protocol would misrepresent the amount of cellulose actually recognized and adsorbed to by cellulases. However, one benefit of using Simons’ stain over other similar techniques (Mercury porosimetry, Nitrogen adsorbtion, Solute exchange, DSC, TD-NMR), is that Simons’ stain is specific for cellulose, and thus does not bind to lignin when used for analyzing lignocellulosic substrates (Kitamura and Kyoshi, 1971). This property allows Simons’ stain to be used to determine not just the total amount of accessible surface area of the substrate, but rather the total amount of accessible cellulose, which is a more important factor when considering enzymatic hydrolysis (Arantes and Saddler, 2011; Yu and Atalla, 1998). 1.7.8 Cellulase adsorption One way of getting around the inability of Simons’ stain to differentiate between specific subsites of cellulose recognized by cellulases is to use the cellulases themselves as accessibility probes. This typically involves incubating either a mixture of cellulases or a purified cellulase with the substrate and quantifying the total amount of protein adsorbed. While this technique gives an accurate representation of the amount of accessible surface area of the substrate, there are two key problems with using this approach. First, unless the adsorption study is carried out at 4°C (which is not representative of hydrolysis reaction conditions), the cellulases will hydrolyze the substrate, thereby changing the substrate accessibility during the course of the experiment; and second, cellulases are known to irreversibly adsorb to lignin, which prevents the accurate quantification of accessible cellulose. Lignin-binding can be prevented by first blocking the accessible sites on lignin with a protein such as bovine serum albumin (BSA), which binds to lignin but not to cellulose (Yang and Wyman, 2006). 66 Recent attempts at using cellulases to quantify accessibility have involved the production of a fluorescently-tagged cellulase to enhance the accuracy and sensitivity of protein adsorption experiments, however the problems with lignin-binding and substrate hydrolysis remain. In addition, the use of a monocomponent cellulase (such as Cel7A) for quantifying cellulose accessibility (Jeoh et al., 2007) may be misleading, as different cellulases recognize different substructures within the substrate. For example, fluorescently-tagged Cel7A has a cellulose-binding domain that primarily recognizes crystalline cellulose, so use of this probe will only quantify the accessibility of regions of crystalline cellulose (Fox et al., 2013), and will therefore not be capable of accurately quantifying changes in the accessibility of amorphous regions. It is possible that cellulase inhibitors could be used to prevent substrate hydrolysis during protein adsorption experiments at relevant temperatures (i.e. 50°C), however, while inhibitors such as hexachloropalladate have been shown to inhibit Cel7a from T. reesei, this compound is incapable of inhibiting all the enzymes present in commercial cellulase preparations (Lassig et al., 1995; Shultz et al., 1995). Thus the inhibition approach would only work if hexachloropalladate-inhibitable cellulases were used for the adsorption studies. Overall, protein adsorption studies coupled with BSA blocking (Eriksson et al., 2002; Yang and Wyman, 2006) can be used to quantify the total amount of cellulose accessible to cellulase, however problems with substrate hydrolysis during the experiment, as well as issues with using subsite-specific monocomponent cellulases (such as the crystalline-cellulose specific Cel7A) render this technique imperfect for use in quantifying protein-mediated amorphogenesis. 1.7.9 CBM adsorption To date, no technique has stood out as an ideal technique for quantifying changes in surface morphology and accessibility of cellulosic substrates promoted by amorphogenesis-inducing proteins. However, one technique that has recently been applied to tracking the amount and distribution of 67 specific polymers within biomass appears to have significant potential for developing quantitative information on changes in accessibility and surface morphology of the cellulosic fraction of biomass (Filonova et al., 2007a, 2007b; Kawakubo et al., 2010). This technique exploits the binding specificity of existing biological probes and involves the specific adsorption of CBMs to their particular carbohydrate targets within the biomass. This is a particularly interesting technique because it is rapid, cheap, sensitive, non-toxic, and can be performed under conditions representative of enzymatic hydrolysis conditions (i.e. the samples do not need to be dried and can be assayed at 50°C). CBM binding can be calculated by quantifying the depletion of the CBM from the solution, either by measuring the OD280 and using the extinction coefficient of the particular CBM employed, or by conjugating a fluorescent probe and spectrophotometrically measuring the depletion of this probe from the solution. Alternatively, it has been demonstrated that a His-tag can be conjugated to the CBM, allowing for subsequent anti-His detection for quantification of binding (McCartney et al., 2004). Adsorption of CBMs to biomass has previously been applied by Hildén et al. (2003), who demonstrated that a fluorescein isothiocyanate (FITC) tagged cellulose-binding CBM from Phanerochaete chrysosporium Cel7D did not bind mannan or xylan, and could thus be used as a probe for specifically recognizing cellulose within lignocellulosic materials such as spruce, birch and pulp fibres (Hildén et al., 2003). The adsorption of cellulose specific CBMs for quantifying the amount of accessible cellulose was later performed by other groups, who used this technique to correlate enzymatic hydrolyzability with cellulose accessibility (Hong et al., 2007; Liu et al., 2012). Expanding on the application of CBMs for quantifying the amount of accessible cellulose, this technique was also used to track the accessibility of a variety of polymers in planta. For example, McCartney et al. in 2004 conjugated His-tags to CBMs from families 2a, 6 and 29, enabling anti-his tag detection of these CBMs. This technique was used to demonstrate the in planta binding of CBM2a to 68 crystalline cellulose, CBM6 to β-1,4 xylan, and CBM29 to a variety of polymers containing β-1,4 linked glucan or mannan as the backbone (McCartney et al., 2004). More recently, Filonova et al. (2007a) demonstrated the use of fluorescently-tagged mannan-specific CBMs for quantifying the accessibility of mannan in wood tissues and pulp fibres. In this work, the authors also used a crystalline-cellulose specific CBM (CBM1 from T. reesei Cel7A) to quantify the amount of accessible crystalline cellulose. Highlighting the exceptional specificities of CBMs for their target polymers, these researchers demonstrated that each of the two mannan-specific CBMs they employed (CBM27(TmMan5) and CBM35(CjMan5C)) exhibited greater specificity for mannan than a monoclonal antibody specific for β-1,4 mannan/ β-1,4 galactomannan. In all of these CBM adsorption experiments, a non-specific protein is first added to the biomass in order to block the sites on the surface of lignin within the biomass which may non-specifically adsorb to the CBMs, thereby artificially increasing the measured amount of CBM bound to the cellulose. For example, Filonova et al. (2007a) performed a pre-incubation with ovalbumin to prevent non-specific adsorption of the CBMs to lignin. In addition to applying CBMs as molecular probes for the various cell wall polymers, the highly specific nature of CBM binding has previously been employed to quantify the amounts of accessible crystalline and accessible amorphous cellulose in softwoods subjected to a variety of pretreatment methods (Kawakubo et al., 2010; McLean et al., 2002). In this work a family 3 and a family 28 CBM from C. josui were fluorescently tagged using Cyan Fluorescent Protein (CFP) and used to quantify the amount of accessible crystalline and amorphous cellulose, respectively. In this work, a linear correlation was observed between the binding of either CBM and the saccharificiation yield during enzymatic deconstruction of the pretreated biomass. Surprisingly, both CBMs were found to preferentially bind to dislocations within the fibres. This is somewhat contrary to the expected binding pattern, as these dislocations are thought to be enriched in amorphous cellulose (Hidayat et al., 2012), so it would be expected that binding of CBM28 (with preferential binding toward amorphous cellulose) would be 69 enriched at these regions, but CBM3 would not necessarily be expected to bind preferentially to these regions, as it was thought at the time to be a crystalline-cellulose specific CBM. This apparent discrepancy can be explained by the more recent work by Fox et al., who demonstrated that a similar family 3 CBM from C. thermocellum appeared to be the most promiscuous of all CBMs studied, binding to both crystalline and amorphous cellulose (Fox et al., 2013). This suggests that CBM3 was perhaps not the best candidate for use as a probe for crystalline cellulose, and reconciles the apparent binding of a ‘crystalline-specific’ CBM to the more amorphous cellulose within fibre dislocations. Although the role of CBMs in localizing catalytic domains to their target polymers within the lignocellulosic matrix has been understood for decades, it was only recently that the potential for exploiting the specificities of CBMs as highly specific molecular probes was realized. This application of CBMs has only recently been explored, but appears to have great potential as a powerful tool in determining the abundance and localization of specific polymers within the complex, heterogeneous multi-level structure of the lignocellulosic matrix. In Section 3.2 of this work, adsorption of CBM2a from C. fimi was used to quantify the amount of accessible crystalline cellulose, while CBM44 from C. thermocellum was used to quantify the amount of accessible amorphous cellulose. This pair of probes is used to follow changes in the accessibility and surface morphology of a pure cellulosic substrate after chemical or protein-induced disruption of the cellulosic structure. An additional potential benefit of this CBM adsorption technique revolves around the ability of these probes to recognize very fine changes in cellulose accessibility occurring at the nanoscopic level. While it is relatively easy to separate nanoscopic and microscopic effects from those at the macroscopic level, the differences between nanoscopic and microscopic effects are more subtle and challenging to quantify. For example, techniques such as Simons’ Stain and cellulase adsorption will likely be incapable of differentiating between increases in accessibility at the microscopic level (i.e. increased accessibility 70 between adjacent microfibrils) and at the nanoscopic level (i.e. increased accessibility of individual cellulose chains at the level of the elementary fibril). Recent work has suggested that enzymatic hydrolysis of cellulosic substrates promotes dramatic changes in the structural organization of the cellulose at the microscopic level, while promoting negligible changes at the nanoscopic level (Chauve et al., 2013). This highlights the need to be able to differentiate between changes at the nanoscopic and microscopic levels. Additionally, this suggests that even cellulases from the hyper-cellulolytic T. reesei strain employed in the work by Chauve et al. (2013) are unable to directly break apart the nanoscopic structures found within cellulosic substrates, although it is possible that this inability could be due at least in part to the particular reaction conditions used (i.e. the particular enzymes and substrate used). It therefore seems possible that dramatic improvements in the efficiency of enzymatic deconstruction of cellulose could be achieved if disruptive proteins were capable of opening up the nanoscale assemblies within the fibrillar structure of cellulose. It should be noted that the putative ability of CBMs to disrupt biomass (discussed in Sections 1.3 and 1.5) occurs at timescales in the range of several hours or days, whereas the application of similar CBMs to quantifying cellulose accessibility (described in this Section and in more detail in Section 3.2) involve only a 60 minute adsorption step, resulting in negligible disruption of the biomass. Due to the ability of CBMs to recognize the fine substructures of cellulose, these protein domains can be used as probes for changes occurring at the nanoscopic level of the biomass. Specifically, Type B CBMs such as CBM44 used in this work are known to bind to isolated carbohydrate chains that have been separated from the bulk of the fibrillar structures (Boraston et al., 2004; Najmudin et al., 2006). Conversely, Type A CBMs such as CBM2a preferentially bind to the planar surfaces present on the faces of fibrillar structures (Boraston et al., 2004; McLean et al., 2000). Thus the amount of binding of each CBM, the ratio of the two CBMs, and the sum of their binding can be used together to gain insights into changes occurring in the biomass at both the nanoscopic and microscopic levels of 71 organization. In other words, the amount of the CBMs bound to the cellulose give an indication of the microscopic accessibility, while the ratio between the two CBMs indicates nanoscopic surface features of the cellulose. The application of CBMs to quantifying changes occurring at the nanoscopic and microscopic levels of biomass organization are discussed in much more detail in Section 3.2 of this thesis. 1.7.10 Summary of techniques Techniques such as the water retention value, FQA, solute exclusion, DSC, TD-NMR, particle scattering, Simons’ stain, cellulase adsorption and CBM adsorption can be used to measure the accessibility of lignocellulosic biomass at various levels of substrate organization. Since the end goal of amorphogenesis is to enhance accessibility thereby facilitating access of the enzymes to the carbohydrate chains, it is likely that these techniques would therefore be applicable to quantifying amorphogenesis. In order to better understand the relationship between amorphogenesis and enzymatic hydrolyzability, it would be ideal to apply a technique capable of both quantifying amorphogenesis and providing an accurate predictor of the enzymatic hydrolyzability of the biomass. For these reasons, Section 3.1 of this work focuses on selecting and developing a technique which can accurately predict the enzymatic hydrolyzability of lignocellulosic biomass, while in subsequent sections (3.2.5, 3.3.4, and 3.4.3.3) the selected technique is used to quantify the manifestations of amorphogenesis. 1.8 Thesis overview and research approach 1.8.1 Summary The limited accessibility of the cellulose within lignocellulosic biomass is likely the single most influential factor restricting efficient enzymatic hydrolysis (Arantes and Saddler, 2010, 2011). This limited accessibility is thus a significant contributor to the high enzyme loadings required for efficient 72 deconstruction of biomass into sugars for subsequent conversion to fuels and chemicals. These high enzyme loadings increase the cost of the hydrolysis step, which is known to be one of the major cost contributors in the overall biomass to fuels and chemicals process (Humbird et al., 2011; Klein-Marcuschamer et al., 2012; Stephen et al., 2012). Although most pretreatments are used to “open up” the biomass prior to enzymatic hydrolysis, these pretreatments must always be carried out at compromised conditions in order to reduce energy and chemical consumption and to prevent the degradation of fermentable sugars from occurring during the pretreatment step (discussed in Section 1.1.8). The pretreated biomass will therefore retain some structural features, and will not be completely opened up and fully accessible to the enzymes involved in deconstruction. In order to get full access to the cellulosic fraction of the biomass, the enzymes themselves must be capable of opening up and disrupting the structure of the lignocellulosic matrix. While it is well known that the hydrolytic and oxidative enzymes are capable of directly cleaving and solubilizing the carbohydrate chains, these enzymes must first be able to diffuse to their accessible carbohydrates (i.e. cellulose) within the lignocellulosic matrix. Recently, non-hydrolytic, non-oxidative proteins, known as amorphogenesis-inducing proteins, have been identified which appear to be capable of opening up the fibrillar structure of the biomass without directly breaking it down into its component sugars. These proteins could play a major role in facilitating the action of the hydrolytic and oxidative enzymes by disrupting the structure of the biomass and consequently enhancing the accessibility of the cellulose and other carbohydrates to the action of the hydrolytic and oxidative enzymes (Arantes and Saddler, 2010). By enhancing the accessibility of the carbohydrates within the biomass to the enzymes involved in biomass deconstruction, these amorphogenesis-inducing proteins should be able to enhance the overall efficiency of the enzyme mixture, thereby lowering the cost of the enzymatic hydrolysis step of the bioconversion process. 73 One of the major issues when working with amorphogenesis-inducing proteins revolves around difficulties encountered when trying to quantify their disruptive effects. This inability to quantify the effects of these proteins hinders efforts to improve their disruptive activities, and prevents accurate comparisons between potential candidate proteins for enhancing the efficiency of biomass disruption. As Lord Kelvin stated, “To measure is to know. If you cannot measure it, you cannot improve it.” For this reason, a significant portion of this work was dedicated to developing a technique involving CBM adsorption for quantifying biomass disruption promoted by amorphogenesis-inducing proteins. In addition this work focussed on developing a better understanding of the effects of a particularly interesting disruptive protein, Swollenin, using both the CBM adsorption technique, as well as a suite of other approaches. Overall, the goal of the work outlined in this thesis was to develop a technique for broad application to better understanding amorphogenesis-inducing proteins and to use this technique and others to enhance our knowledge of how Swollenin affects industrially relevant lignocellulosic biomass. 1.8.2 Research approach In order to achieve the goals outlined above, this work was divided into four Chapters. These Chapters make up Sections 3.1, 3.2, 3.3 and 3.4, and include: 1. Assessing the role of accessibility in the enzymatic hydrolysis of pretreated lignocellulosic biomass via a suite of different techniques. Selection of one technique for further development for use in quantifying the amorphogenesis step of biomass deconstruction. 2. Develop and apply this technique to accurately quantifying the effects of amorphogenesis-inducing proteins on the cellulosic fraction of biomass. 74 3. After quantifying the disruptive effect on cellulosic biomass, quantify the macroscopic disruptive effects of Swollenin on a model cellulosic fibre as well as on a range of pretreated lignocellulosic materials. 4. Investigate the microscopic/nanoscopic disruptive effects of Swollenin on an industrially-relevant pretreated lignocellulosic substrate. Assess any potential synergistic interactions with hydrolases to enhance overall sugar yields. In the first Chapter (Section 3.1), the concept that cellulose accessibility to cellulases is a key factor in determining the ease of enzymatic hydrolyzability was assessed in order to ensure that efforts to enhance cellulose accessibility through amorphogenesis would indeed be beneficial to the efficiency of enzymatic biomass deconstruction. In this section, a variety of industrially relevant pretreated lignocellulosic substrates were prepared, and a number of techniques were used to assess any correlation between cellulose accessibility and the ease of enzymatic hydrolysis of these substrates. The main goal of this part of the work was to determine what technique would be best suited for both quantifying amorphogenesis and also acting as a good indicator of enzymatic hydrolyzability. Cellulose accessibility measurements were carried out using traditional techniques such as Simons’ Stain, cellulase adsorption and fibre quality analysis, and at the same time, initial testing of the CBM probe technique was carried out. Of these techniques, the CBM adsorption assay was selected for further development as it appeared to be the best candidate for both quantifying amorphogenesis and providing an indicator of the enzymatic hydrolyzability of the substrate. In the second chapter (Section 3.2), the CBM adsorption technique was further developed where a Type A and a Type B CBM were selected as probes for preferentially recognizing crystalline and amorphous cellulose, respectively. By measuring the amount of each CBM adsorbed to the biomass, the total cellulose accessibility could be determined, along with the amount of accessible amorphous 75 cellulose and accessible crystalline cellulose. This technique was successfully used to quantitatively measure changes in microscopic/nanoscopic cellulose accessibility and surface morphology of the cellulose after treatment with Swollenin. Additionally, the observed changes in the CBM adsorption profile provided new insights into the specific effects of Swollenin on cellulosic biomass at the microscopic/nanoscopic level. After quantifying the microscopic/nanoscopic effects of Swollenin on a model cellulosic substrate, the work reported in Chapter 3 (Section 3.3) investigated the effects of Swollenin on the macroscopic properties of cellulosic and lignocellulosic biomass. Initially, the fragmentation profile of fibres during enzymatic hydrolysis was assessed. This indicated that fragmentation occurs extremely rapidly at the very early stages of enzymatic deconstruction prior to the significant solubilisation of sugars. This suggested that fragmentation could be considered as one indication of an amorphogenesis step that facilitates the subsequent enzymatic hydrolysis and depolymerisation of the cellulose. The ability of Swollenin to promote macroscopic amorphogenesis on a model cellulosic substrate, as well as on a range of pretreated lignocellulosic woody biomass, was then assessed. Swollenin did not promote fragmentation or swelling of the lignocellulosic substrates. However, when Swollenin was applied to a model cellulosic substrate, Swollenin-induced fragmentation was observed. The surface morphology of the model cellulose at the dislocations of the cellulosic fibres where Swollenin-induced fragmentation occurs was then analyzed. This was performed by conjugating the Type A and Type B CBMs to different fluorescent probes and using confocal microscopy to assess their binding profile along the length of the cellulosic fibres. The subsequent observations suggested that the dislocations of the fibres were enriched in amorphous cellulose, confirming that Swollenin targets the amorphous regions of the cellulose. This suggested that Swollenin promotes fragmentation of cellulosic fibres by targeted disruption of the amorphous cellulose within fibre dislocations. This supported the 76 results reported in Section 3.2 which showed that Swollenin targeted the amorphous regions of cellulose rather than the crystalline regions. Although Swollenin did not appear to promote macroscopic disruption of woody lignocellulosic biomass, it is likely that Swollenin might instead play a role at the more microscopic/nanoscopic level (as indicated when using model cellulosic substrates, Section 3.2.). Although previous work had employed Swollenin to enhance the hydrolysis of poplar, the protein sequence of Swollenin is more similar to the expansin families involved in loosening grass cell walls than those involved in loosening woody cell walls. For these reasons, the final Chapter of this work (Section 3.4) focussed on assessing potential microscopic/nanoscopic effects of Swollenin on an industrially relevant grassy substrate, steam pretreated corn stover. Application of the CBM adsorption technique developed in Section 3.2 to Swollenin treated corn stover revealed that Swollenin directly disrupted the more amorphous regions of the biomass. Additionally, the synergism experiments indicated that Swollenin synergized strongly with xylanases in the release of xylose. Although previous work has demonstrated that Swollenin acts to disrupt pure cellulosic substrates, the work reported here suggested that the main role of Swollenin is to enhance the disruption and solubilisation of the hemicellulosic portion of grassy biomass. This thesis work demonstrated the successful application of a two-probe CBM adsorption technique for quantifying changes in cellulose accessibility and surface morphology during the amorphogenesis step of enzymatic hydrolysis. Additionally, it was found that Swollenin, which has previously only been shown to disrupt model cellulosic substrates, plays a more significant role in the disruption of the amorphous regions of cellulose and on the hemicellulosic fraction of industrially relevant pretreated lignocellulosic biomass. 77 2. Materials and methods 2.1 Proteins and enzymes CEL7A, CEL5A, XYN10A and XYN11A were purified as described previously (Hu et al., 2011). CtCBM44 from Clostridium thermocellum was purchased from NZYTech (Lisbon, Portugal, product code: CR0049) Swollenin was produced at the VTT Technical Research Center of Finland by Merjä Pentilla, Markku Saloheimo, and Martina Andberg. Briefly, Swollenin was expressed in T. reesei under the cbh1 promoter with a C-terminal His6-tag and purified using immobilized metal ion affinity chromatography followed by anion exchange chromatography (DEAE Sepharose) following the procedure previously used for CEL61A (Karlsson et al., 2001). CBM2a from C. fimi was expressed in E. coli BL-21 using the plasmid construction described previously and purified as also described previously (Ong et al., 1993). Cells were grown at 37 °C in Terrific Broth to OD280 ~2.0, then induced with 0.2 mM Isopropyl-β-D-1-thiogalactopyranoside (IPTG) (Sigma-Aldrich) and incubated for an additional 24-32 h at 30 °C. Cells were harvested by centrifugation, and the pellet was then used immediately for purification or frozen for future use. Cells were lysed by shaking in 5 mL Bugbuster (EMD Millipore) per gram cell paste supplemented with benzonase nuclease (EMD Millipore) for 20 minutes at room temperature. Lysed cells were clarified by centrifugation, and the clarified extract was added to Avicel (15 g Avicel per L of original cell culture) (Sigma-Aldrich) and incubated for 1 h at room temperature or overnight at 4 °C. The slurry was then filtered and washed three times with 1 M Sodium Chloride in 50mM Potassium Phosphate, pH 7.0 and three times with 50mM Potassium Phosphate pH 7.0, then eluted by incubating the cellulose paste with 6 M guanidine hydrochloride for 30 min at room temperature before filtering. All subsequent steps were performed on ice to prevent precipitation. The CfCBM2a solution in 6 M guanidine hydrochloride was concentrated to 78 ~1/10 volume over an Amicon (EMD-Millipore) ultrafiltration apparatus using a 10 kDa polysulfone membrane (EMD-Millipore). The concentrated solution was then centrifuged to remove fines and precipitate, then slowly diluted ~20 times with 1 mM ice cold potassium phosphate pH 7.0 in order to refold the CBM. This solution was then concentrated again, and rediluted with 1mM ice cold potassium phosphate pH 7.0. Finally, CfCBM2a was desalted into 50 mM potassium phosphate pH 7.0. Purity was confirmed by SDS-PAGE. In order to conjugate fluorescent tags to the CBMs, each CBM was made up to 5 mg/mL in 1 mL 0.1 M sodium bicarbonate buffer, pH 8.3. A 10:1 ratio of protein to dye was used for tagging. Specifically, 0.5 mg of dye (6-((7-Amino-4-Methylcoumarin-3-Acetyl)amino)Hexanoic Acid, (AMCA-X, Invitrogen) for CfCBM2a and Oregon Green 514 (OG, Invitrogen) for CtCBM44) was dissolved in 50 µL dimethylsulfoxide (DMSO). While stirring/vortexing the protein solution (~5 mg in 1 mL), the 50 µL AMCA-X or OG solution in DMSO was added and incubated at room temperature with continuous shaking for 1 h. The dyed CBMs were then desalted into 50mM Potassium Phosphate pH 7.0. It should be noted that the binding of CtCBM44 was significantly reduced after tagging, suggesting that some of the CBMs were tagged at or near the binding cleft, inhibiting the binding of the CBM. However, it was apparent that not all CtCBM44 molecules were inhibited in their binding, as confocal microscopy was still able to detect these CBMs bound to the cellulose fibres. 2.2 Biomass preparation Representatives from softwood (lodgepole pine, douglas fir), hardwood (hybrid poplar), and agricultural residues (corn stover) were pretreated at near-optimal conditions using SO2-catalyzed steam pretreatment and ethanol-organosolv pretreatment as described previously (Arantes and Saddler, 2011; Tu et al., 2007). After pretreatment, samples were filtered and washed thoroughly, then refrigerated until use. 79 Model cellulosic substrates employed in this work included Avicel PH-101 (Sigma-Aldrich), hardwood dissolving pulp (Tembec, Montreal, QC, Canada), Cellulose III (kindly provided by the National Renewable Energy Laboratory (NREL, Golden, CO, USA), cellulose nanocrystals (CNC, kindly provided by Alan Rudie and Richard Reiner of the United States Department of Agriculture Forest Products Laboratory (Madison, WI)). Additionally, Cellulose II was derived from Avicel according to a method described previously (Mittal et al., 2011) and phosphoric acid swollen cellulose (PASC) was also derived from Avicel according to a method described previously (Zhang et al., 2006). Phosphoric acid treatments were also used to disrupt cotton fibres for the CBM-adsorption experiments in Section 3.2. Briefly, ice-cold phosphoric acid (Fisher Scientific) solutions were produced at various concentrations and 14.5 mL was added to 50 mL centrifuge tubes containing 0.2 g cotton fibres (Sigma-Aldrich, product code: C6663) pre-wetted with 0.5 mL nanopure water to give final o-phosphoric acid concentrations of 0–78% w/w. Samples were incubated for one hour on ice with occasional mixing. Ice-cold nanopure water (35 mL) was slowly added to each sample, followed by centrifugation at 10,000 g for 15 minutes. The fibres were resuspended in 50 mL nanopure water and washed a further four times with 50 mL nanopure water, followed by one wash with 50 mL 20 mM Na2CO3 and two subsequent washes in 50 mL nanopure water. The cotton fibres were then lyophilized overnight. 2.3 Determining pretreated biomass composition The chemical composition of the pretreated materials was determined according to Technical Association of the Pulp and Paper Industry (TAPPI) standard method T222 om-88. Monomeric sugars were measured by HPAEC-PAD (High Performance Anion Exchange Chromatography with Pulsed Amperometric Detection). A Dionex system with Carbopak PA1 column was used with fucose as internal standard, as previously described (Bura et al., 2002). All analyses were performed in triplicate. 80 2.4 Enzymatic hydrolysis and Swollenin treatments For all hydrolysis reactions except the synergism experiments, two commercial preparations (both from Novozymes, Bagsværd, Denmark), a cellulase cocktail (Celluclast 1.5 L; protein content 129.8 mg/mL) derived from Trichoderma reesei and a β-glucosidase preparation (Novozym 188; protein content 233 mg/mL) derived from Aspergillus niger, were used. Protein concentrations were determined using the modified ninhydrin method (Starcher, 2001). Bovine serum albumin (BSA, Sigma-Aldrich) was used as the protein standard. Determining the minimum enzyme loading required for 70% conversion was carried out as described previously (Arantes and Saddler, 2011). For these hydrolysis-to-70% conversion experiments, hydrolysis of pretreated substrates was carried out batch-wise at 2% (w/v) solids loading in sodium acetate buffer 50 mmol/L pH 4.8, supplemented with 0.02% (w/v) tetracycline (Sigma-Aldrich) and 0.015% (w/v) cyclohexamide (Sigma-Aldrich), to prevent microbial contamination. The reaction mixtures (1 mL) were mechanically shaken in an orbital shaker incubator (Combi-D24 hybridization incubator, FINEPCR®, Yang-Chung, Seoul, Korea) at 50 °C. The conditions for cellulase and β-glucosidase loadings, hydrolysis time, and solids loadings were determined according to the statistical design of experiments (Arantes and Saddler, 2011). Glucose concentration was determined using a modified glucose oxidase and horseradish peroxidase assay (Berlin et al., 2006a). Hydrolysis yields (%) of the pretreated substrates were calculated from the cellulose content as a percentage of the theoretically available cellulose in the pretreated substrate. For the hydrolysis reactions where the extent of hydrolysis was assessed for longer than 48 hours, 1mL reactions were performed as described above for the 70% conversion reactions, but with a Celluclast 1.5L loading of 20 mg/g supplemented with 5 cellobiohydrolase units (CBU) per gram pulp (dry weight). 81 The variably-hydrolyzed dissolving pulp samples used for tracking changes in fibre length and CBM adsorption profiles over the course of hydrolysis were prepared as follows. Dissolving pulp (6 g dry weight) was weighed into 250 mL Erlenmeyer flasks and hydrolyzed in 120 mL 50 mM Sodium Acetate buffer, pH 5.0, using an enzyme loading of 13 mg Celluclast 1.5L (Novozymes, Bagsværd, Denmark) per gram pulp (dry weight), supplemented with 5 CBU per gram pulp (dry weight). Flasks were stoppered and incubated in a shaking incubator at 50 °C. After the desired elapsed time, flasks were transferred to a hot water bath set to 100 °C and incubated for 15 minutes. For the 0 hour timepoint, the buffer was added to the pulp and the sample was transferred to the hot water bath prior to adding enzyme. As the temperature increased past 60 °C, enzyme was added. This was done to achieve very rapid inactivation of the enzymes in the 0h sample. Hydrolyzed pulps were then filtered, washed 3 times with water, then incubated with protease from Aspergillus melleus (Sigma, product code: P4032) overnight at 37 °C. After protease treatment, samples were filtered, washed three times with 100 mL 1 M NaCl in 50mM Potassium Phosphate pH 7.0, three times with 100mL 50mM Potassium Phosphate pH 7.0, and three times with 100 mL nanopure water. Removal of protein from the pulp was confirmed by the Ninhydrin assay, modified from Starcher (2001) for analyzing protein contents within solid substrate. Briefly, 10 mg dry weight of each hydrolyzed pulp sample was incubated in 100 µL 1 N HCl overnight at 105 °C. Ninhydrin solution (200 µL at 2%, Sigma-Aldrich) was then added, and the mixture was heated at 100 °C for 20 minutes, then allowed to cool. Ethanol (1 mL at 50% (v/v) was then added, the reaction was vortexed and centrifuged, then 300 µL supernatant (diluted 1:10 in nanopure water) was transferred to a microplate for reading. Enzyme treatment of dissolving pulp for quantification of enzyme-induced fibre fragmentation was carried out in 2 mL screw cap tubes containing 2 mg (dry weight) dissolving pulp in 1.5 mL 50 mM Sodium Acetate pH 5.0 with 50 μg enzyme per mg pulp. Samples were incubated overnight with shaking at 50 °C. Samples were heat inactivated at 100 °C for 10 minutes prior to performing FQA. 82 Swollenin-treatments for subsequent CBM analysis of accessibility and surface morphology were performed as described for the fibre fragmentation studies. Prior to CBM adsorption, the Swollenin-treated fibres with incubated overnight at 37 °C, heat-inactivated at 100 °C for 10 minutes, centrifuged, then washed thoroughly by exchanging the supernatant with fresh buffer six times. Enzymatic hydrolysis of SPCS was carried out to determine the degree of synergy of Swollenin with isolated hydrolytic components. Hydrolysis was performed using 10 µg hydrolytic enzyme plus 10 µg Swollenin or BSA per mg dry SPCS, as well as 2 µg β-glucosidase (Novo 188, Novozymes, Bagsværd, Denmark) per mg dry SPCS. Hydrolysis was carried out using 10 mg SPCS (dry weight) in 1 mL 50 mM Na-Acetate buffer, pH 5.0 (Fisher). The degree of synergy (DS) was calculated using the following equation:  =  ∑ ,   where SRcombined represents the sugar release when the Swollenin and hydrolase were incubated simultaneously with substrates, SRSwollenin represents the amount of sugar released by Swollenin alone, and SRHydrolase represents the amount of sugar released by each hydrolase alone. For the individual reactions involving either Swollenin or a hydrolase, BSA was used as a protein control to bring the protein concentration of the individual treatments up to the same final concentration of the combined (Swollenin + hydrolase) treatments. 2.5 Accessibility measurements 2.5.1 Simons’ staining Simons' Stain (SS), which was derived from a staining technique used in the pulp and paper industry to examine changes in the physical structure of pulp fibres under the microscope was adapted for evaluating the pore structure (internal surface area) of cellulosic materials (Yu and Atalla, 1998). It 83 was performed according to the modified procedure of Chandra et al. (2008a). Pontamine fast orange 6RN (direct orange; DO) and Pontamine fast sky blue 6BX (direct blue; DB) dyes were used (Pylam Products Co. Inc., Garden City, NY, USA). Fractionation of DO was performed according to Esteghlalian et al. (2001a). 2.5.2 FQA and settlability Fibre dimensions (fibre length, fibre width), both population and distribution, were determined using a high resolution Fibre Quality Analyzer (FQA) (LDA02, OpTest Equipment, Inc., Hawkesbury, ON, Canada) as described previously (Robertson et al., 1999). The number of fibres counted per sample was 20,000. The ranges of fibre length and fibre width measured in this study were 0.05 – 10.00 mm and 7 – 60 μm, respectively. The settlability was visualized by weighing out 10mg (dry weight) of hydrolyzed pulp, staining overnight in 1 mL of 0.5 mg/mL Direct Orange (Pontamine Fast Orange 6RN, lot no. 814071, Pylam Products Co. Inc., Garden City, NY) in PBS buffer at 70 °C in a shaking incubator. Samples were then centrifuged, and the liquid exchanged ten times with water to remove unbound dye. Dyed fibres were then allowed to gravity settle in upright 1.5 mL Eppendorf tubes for two days. 2.5.3 Cellulase adsorption The maximum extent of protein (cellulase and β-glucosidase) adsorption was used as an indication of the surface area of a particular substrate available for protein binding. Protein adsorption isotherms were established by varying the amounts of protein (cellulase + β-glucosidase) added to the different pretreated substrates (2 mg/mL) in sodium acetate buffer (50 mmol/L, pH 4.8). Free protein was determined by measuring the amount of protein in the supernatant after incubation at 4 °C and 150 rpm for 1 hour to reach equilibrium. Bound protein was calculated as the difference between free 84 protein and the total protein initially added to the reaction medium. The protein content was determined using the ninhydrin assay (Starcher, 2001). The experimental data was fitted to the Langmuir adsorption isotherm using the following linearized form of the equation: 1/Pads=1/PmaxKp+(1/Pmax)P, where P is the concentration of unadsorbed protein (mg of protein/mL), Pads is the concentration of adsorbed protein (mg of protein/mg of substrate), Pmax is the maximal adsorbed protein (mg of protein/mg of substrate) and Kp is the equilibrium constant (mL/mg of protein). 2.5.4 Nitrogen adsorption The surface area of pretreated biomass samples was determined by nitrogen adsorption using an 11-point BET procedure (Satterfield, 1991) using an Autosorb-1 surface area analyzer (Quantachrome instruments, FL, USA). 2.5.5 CBM adsorption For the untagged CBM adsorption experiments, biomass samples were weighed out (usually 2-5 mg dry weight) and incubated in 1 mL CBM solution containing a known concentration of a single CBM. Samples were incubated for 60 minutes in a FinepcrCombi SV12 hybridization incubator at 30 rpm to allow CBM adsorption to reach equilibrium, and then centrifuged at 16,000 g for 10 minutes in a benchtop centrifuge. The absorbance of the supernatant at 280nm was then measured on a Varian Cary 50 Bio Spectrophotometer (Agilent Technologies, Mississauga, Ontario, Canada) and used to calculate the amount of CBM remaining in solution. The extinction coefficients used to calculate CBM adsorption were 27,625 M-1 and 27,365 M-1 for CBM2a and CBM44, respectively (McLean et al., 2002; Pace et al., 1995). 85 A key limitation to this technique is that, if CBM loadings are too low, all of the CBM will be bound to the cellulose, making accurate quantification of the amount of CBM left in the supernatant impossible. Although calculating the saturating levels of CBM adsorption for every sample would be ideal for quantifying CBM adsorption to each sample, this method would involve using large amounts of CBM for every sample. As a compromise, various substrates were tested by performing CBM adsorption over a range of concentrations, from 50 µg per mg dry biomass up to 1000 µg/mg. This was done to determine what minimum CBM loading provided sufficient CBM in the supernatant over a broad range of cellulosic and lignocellulosic substrates for accurate quantification by OD280. This work revealed that a CBM loading of 250 µg CBM per mg cellulose provided sufficient residual CBM in the supernatant for accurate quantification, even on highly accessible substrates such as PASC. Thus the amount of CBM adsorbed to each sample in this work does not represent the maximum amount of CBM adsorbed, but rather the amount adsorbed at equilibrium when incubated with 250 µg CBM per mg dry biomass. 2.6 CBM adsorption for Scatchard plots Dissolving pulp (5 mg dry weight) was weighed into 2 mL screwcap tubes and incubated with CBM2a or CBM44 alone, or with a 50:50 mixture of the two CBMs. A range of concentrations were used, from 0 to 1000 µg CBM per mg cellulose. Samples were incubated for 2 hours at room temperature to ensure equilibrium was reached. The supernatants were then analyzed for protein concentration using the Ninhydrin method using known concentrations of BSA as standards. In brief, 40 µL supernatant was incubated with 100 µL 1 M HCl overnight at 100°C. Ninhydrin solution (2%, 200 µL) was then added and the samples were incubated for 20 min at 100°C. Dilute ethanol (50% v/v, 1 mL) was then added to each sample. Samples were then vortexed, and 300 µL was transferred to a microplate. Adsorption at 570 nm was then determined using a PerkinElmer Wallac Victor3 1420 multilabel counter running Wallac 1420 Manager software. 86 2.7 SEM imaging Lyophilized biomass samples were mounted on aluminum SEM stubs using double sided tape and sputter-coated with 10 nm Au/Pd (80:20 mix) then imaged on a Hitachi S-2600 VP-SEM (Tokyo, Japan). 2.8 X-ray diffraction The cellulose crystallinity index (CrI) was measured by X-ray diffraction (XRD) as described by Nishiyama et al. (2002). Briefly, pulp samples were hydrolyzed then washed, filtered and freeze-dried, before mounting onto a zero-background plate. The data was collected with a Bruker D8-Advance powder X-ray diffractometer (Bruker, Billerica, MA, USA). Bruker TOPAS version 4.2 was used to model percent crystallinity and Nishiyama’s cellulose 1β was used to model cellulose. The percentage cellulose crystallinity was calculated as: 100 x (crystalline area/total area), where the total area is equal to the sum of the crystalline area and the amorphous area. Modelling was carried out by Anita Lam, Earth and Ocean Sciences, UBC. 2.9 Confocal microscopy Confocal microscope imaging was performed using an inverted Zeiss Axiovert LSM 5 confocal microscope equipped with epifluorescence, Nomarski optics, and LSM 5 Pascal software. Image acquisition and analysis was done with the LSM 5 Pascal volume-rendering software. Images were opened in the GNU Image Manipulation Program version 2.8 (www.gimp.org). In order to produce the overlaid image, the colored pixels were extracted from both images and minimum threshold pixel intensity was set to filter out low-intensity regions. The remaining pixels were then saturated and overlaid on top of the original SEM image. 87 2.10 Monomer and oligomer analysis after Swollenin treatment 2.10.1 High performance liquid chromatography (HPLC) The supernatants from Swollenin-treated SPCS were analyzed for the presence of oligomeric and monomeric sugars. Monomeric sugars were determined by High Performance Liquid Chromatography (HPLC) on a Dionex DX-3000 HPLC system fitted with an AS3500 autosampler, a UV detector and a GP40 gradient pump. Oligomeric sugars were determined by acid hydrolyzing the supernatant to convert all oligomers into monomers. The total monomers (monomers released by Swollenin and monomers produced from acid-hydrolyzing the oligomers) in the supernatant were then quantified by HPLC. The amount of oligomers released by Swollenin was quantified by subtracting the monomers present in the unhydrolyzed supernatant from the total monomers observed in the acid-hydrolyzed supernatant. 2.10.2 Polyacrylamide carbohydrate electrophoresis (PACE) PACE was carried out by conjugating the solubilized monomers and oligomers to a fluorescent probe and separating these sugars by size on a polyacrylamide gel as described previously (Goubet et al., 2011). In brief, 500 µL of supernatants from the Swollenin and control treated SPCS were lyophilized. The dried sugars were conjugated to the fluorescent probe, 8-aminonaphthalene-1,3,6-trisulfonic acid (ANTS) (Invitrogen) by dissolving the sugars in 10 µL 0.1 M ANTS in 3/17 (v/v) acetic acid/water and 10 µL 0.1 M NaCNBH3 (Acros Organics) in DMSO and incubating overnight at 60 °C. Because this reaction is specific for the reducing end of the monomeric and oligomeric sugars found in the supernatant, a single ANTS molecule was conjugated to each monomer or oligomer, allowing for accurate separation based on electrophoretic mobility of the ANTS-conjugated sugars (Jackson et al., 1994). Tagged sugars were then lyophilized and dissolved in 100 µL 6 M Urea (BioRad) and stored at -20 °C. Polyacrylamide gels (19.5% w/w) were poured and 88 sugar samples were run for 45 minutes at 200 volts and imaged on an AlphaImager 2200 (Alpha Innotech). Densitometry was performed using AlphaImager 2200 Software, with manual peak selection. The process of ANTS derivitization of sugars is shown in Figure 2. OOHOHOHOH OHNaBH3CNANTS OHOHOHOHOH NHO3-S SO3-SO3-O3-S SO3-SO3-NH22Na+ Figure 2: Derivatization process of sugars or oligomers with ANTS in the presence of NaBH3CN. 89 3. Results and discussion 3.1 The role of accessibility in enzymatic hydrolysis 3.1.1 Background As discussed in the introduction, one of the most widely-accepted factors that influences the effective enzymatic deconstruction of lignocellulosic biomass is the accessibility of the cellulosic fraction to the enzymes involved in deconstruction. In order for enzymatic hydrolysis (or oxidative cleavage) to occur, these enzymes must first be capable of diffusing through the densely-packed lignocellulosic matrix to access the β-1,4 bonds within the cellulose chains. As also discussed previously, an ideal pretreatment process could theoretically be used to try to fractionate the cellulose from the bulk of the lignocellulosic biomass and render it more accessible to enzymatic attack. However, the high energy and chemical input required combined with the resultant formation of sugar degradation products during the pretreatment step would lead to higher costs and low overall sugar yields for the bioconversion process (Chandra et al., 2007; Donaldson et al., 1988). For these reasons, pretreatments will always be performed at compromised conditions in order to partially open up the biomass while using minimal energy and chemical inputs and preserving as much of the sugars intact as possible. As pretreatments cannot be used to open up the biomass to fully access the cellulose in a cost-effective manner, even after pretreatment a significant portion of the cellulose remains largely inaccessible to the cellulases. Visual confirmation of this can be seen in scanning electron microscopy images of optimally pretreated biomass samples, where the relatively intact fibrillar structure, comprising mostly microfibrils, can still be observed after pretreatment (Donaldson et al., 1988). While microfibrils appear to make up the dominant component of pretreated plant cell walls, other architectural features that limit the accessibility of the cellulose to cellulases are present over multiple levels of structural organization (see Section 1.1.2) These structural factors limiting the accessibility of cellulose to the enzymes range in scale by several orders of magnitude, from a few nanometers (at the 90 level of elementary fibrils) up to several millimeters (at the level of individual fibres). As described earlier, disruptive proteins appear to be capable of opening up the biomass at each of these levels of organization. However, the dramatic differences in scales provide a challenge when attempting to quantify changes in cellulose accessibility induced by these proteins. Despite the inherent challenges involved in determining cellulose accessibility, it is essential that this metric of pretreated biomass be accurately measured. This is partly because cellulose accessibility to cellulase is likely the single most important factor limiting enzymatic hydrolysis rates (Arantes and Saddler, 2010, 2011; Chandra et al., 2008a; Esteghlalian et al., 2001a; Grethlein, 1985; Jeoh et al., 2007; Sinitsyn et al., 1991; Thompson et al., 1992; Wiman et al., 2012) and also because cellulose accessibility is dependent on a variety of other substrate factors such as cellulose crystallinity, particle size, and lignin and hemicellulose content. Thus, accurate measurements of cellulose accessibility to cellulase provides an indicator of enzymatic hydrolyzability of the substrate, while taking into account a variety of other substrate factors involved in limiting enzymatic deconstruction rates. In order to determine the role of accessibility at different levels of organization in influencing enzymatic hydrolysis rates, five different techniques were used to quantify cellulose accessibility on a range of pretreated biomass: Simons’ Stain, FQA, cellulase adsorption, CBM adsorption, and Nitrogen adsorption. These techniques were chosen due to their relative simplicity and ability to quantify accessibility over a range of organizational scales. Specifically, Simons’ stain was used to measure the porosity (internal/microscopic surface area) of the biomass, while FQA was performed to measure properties related to the external/macroscopic surface area and Nitrogen adsorption was used to quantify nanoscopic accessibility. Finally, both cellulase adsorption and CBM adsorption were used to specifically measure the amount of exposed cellulose that is recognized by carbohydrate-active proteins at the microscopic level. This measurement is particularly important because the inherent fibrillar 91 organization of cellulose presents multiple ‘faces’, which have been shown to be differentially susceptible to enzymatic attack (Lehtiö et al., 2003; McLean et al., 2000; Tormo et al., 1996). It was hypothesized that defining the amount of cellulose that is actually bound by cellulases and CBMs would likely provide more accurate descriptions of hydrolyzability than other techniques which are unable to differentiate between the accessible hydrolyzable cellulose and accessible but unhydrolyzable cellulose. Overall, this range of techniques was used to determine if accessibility at specific levels of substrate organization (e.g. FQA for macroscopic accessibility, SS for microscopic porosity, CBM and cellulase adsorption for microscopic accessibility, and Nitrogen adsorption for nanoscopic accessibility) were particularly important in influencing the enzymatic hydrolyzability of the substrate. Measurements obtained using these techniques on a variety of pretreated biomass samples were correlated with the enzymatic hydrolyzability of a range of lignocellulosic samples. 3.1.2 Substrate preparation In order to prepare a range of different industrially relevant substrates, representative biomass from agricultural residues, hardwood, and softwood were used including corn stover, corn fibre, hybrid poplar, douglas fir, and beetle-killed lodgepole pine. Pretreatments employed included SO2-catalyzed steam and/or ethanol-organosolv (EO) pretreatment (Arantes and Saddler, 2011). The pretreatment conditions used as well as the composition of the biomass after pretreatment are shown in Table 2. These pretreatment conditions were chosen based on previous work that demonstrated that these compromised pretreatment conditions provided optimal sugar recovery while utilizing minimal chemical and energy inputs, resulting in improved cellulose accessibility (Arantes and Saddler, 2011). 3.1.3 Correlating accessibility with enzymatic hydrolyzability Previous work had determine the minimum enzyme loading required to achieve 70% conversion in 72 hours at 2% substrate consistency for the range of pretreated biomass substrates listed above 92 (Arantes and Saddler, 2011). A conversion level of 70% was chosen because typical hydrolysis profiles for pretreated lignocellulosic substrates generally follow a trend whereby the reaction proceeds rapidly over the first portion of hydrolysis, then begins to slow down at higher conversion levels. A representative conversion curve is shown in Figure 3. Conversion of the final 30% of the cellulose requires much longer incubation times, and often proves impossible to fully convert to glucose, leading to diminishing returns for additional hours of hydrolysis time. For these reasons, it has been proposed that enzymatic hydrolysis schemes should involve the conversion of only the readily hydrolyzable fraction of the biomass, defined here as the first 70%, without trying to use excessive reaction time or enzyme loading to hydrolyze the final 30%. Table 2: Pretreatment conditions and substrate composition after pretreatment for a range of substrates Substrate Pretreatment conditions Composition of pretreated feedstocks (% w/w) SO2a-steam pretreatment Arab Galc Glud Xyle Manf AILg Abbreviation Corn stover 190°C, 5 minutes, 3% SO2 0.8 0.2 55.1 12.0 1.9 18.9 SPCS Corn fibre 190°C, 5 minutes, 4% SO2 6.9 2.8 38.2 15.3 2.2 12.6 SPCF Douglas fir 200°C, 5 minutes, 4% SO2 BDLh BDL 50.6 0.4 1.0 47.0 SPDF Lodgepole pine 200°C, 5 minutes, 4% SO2 BDL BDL 52.4 0.6 1.0 45.9 SPLP Ethanol-organosolv pretreatment Corn fibre 170°C, 30 minutes; 65% EtOH, 0.75% H2SO4 2.1 1.6 57.9 11.5 3.0 15.7 OPCF Poplar 195°C, 40 minutes; 70% EtOH, 1.0% H2SO4 BDL BDL 77.0 6.0 2.4 16.0 OPP Lodgepole pine 170°C, 60 minutes; 65% EtOH, 1.1% H2SO4 0.1 0.1 74.8 1.6 1.8 17.3 OPLP aSulfur dioxide, bArabinan, cXylan, dGlucan, fMannan, gAcid-insoluble lignin, hBelow detectable level. 93 Figure 3: Representative conversion curve of glucan conversion over time for a typical enzymatic deconstruction reaction. Several studies have suggested that by only targeting the easily-hydrolyzable portion of the biomass a more cost-effective overall process can be achieved, requiring less enzyme, shorter residence time, and smaller reaction vessels (Gregg and Saddler, 1996; Shen and Agblevor, 2008). The minimum protein loadings required for 70% hydrolysis using a cellulase cocktail (Celluclast 1.5 L) supplemented with β-glucosidase (Novozym 188) for each of the pretreated substrates is shown in Table 3. Table 3: Amount of protein required to achieve 70% conversion after 72 hours at 2% consistency (Arantes and Saddler 2011). Pretreated Biomass OPCF OPLP OPP SPCS SPDF SPLP SPCF Protein required for 70% conversion (mg/g glucan) 18 43 48 54 61 63 23 Most samples were found to require between 40-65 mg enzyme per gram glucan, with the exceptions being the pretreated corn fibre samples, which were found to require far lower enzyme loadings (~20 mg/g glucan), regardless of pretreatment strategy employed. Although it is not clear why 94 corn fibre is more amenable to enzymatic hydrolysis, it should be noted that corn fibre plays an entirely different physiological role in nature compared to the other biomass samples analyzed here. Specifically, corn fibre is a by-product of the corn processing industry and consists of the husks of corn kernels after processing to remove the corn starch, whereas all other samples employed in this study are primarily involved in structural support (i.e. corn stalks, tree trunks). As well as having different physiological roles, corn fibre also has a distinct biomass composition, with higher arabinan and galactan content, and relatively low lignin content (Table 2). In general, the organosolv-pretreated samples were found to require marginally less enzyme for conversion than the steam-pretreated samples, with the steam-pretreated softwoods being particularly recalcitrant to enzymatic hydrolysis. Softwoods are known to have particularly condensed and recalcitrant lignin compared to other substrates (Pan et al., 2005), and while organosolv-pretreatments are capable of solubilizing a significant portion of this lignin, steam-pretreatments have been found to be less effective at removing softwood lignin. This leads to a high proportion of residual lignin in the steam-pretreated softwood biomass substrates, as has been found in previous work (Chandra et al., 2007; Galbe and Zacchi, 2007; Pan et al., 2005) and in this study (Table 2). It is likely that this high lignin content is one of the major contributors to the relative recalcitrance of the steam-pretreated substrates to enzymatic conversion, either through steric masking/blocking enzyme access to the cellulose, or through irreversible non-productive adsorption of enzymes to the lignin fraction of the biomass (Kumar et al., 2012). In softwoods, lignin is thought to play a more significant role in hindering enzymatic deconstruction compared to the minor inhibitory effects of softwood-derived hemicelluloses. However, in the case of agricultural residues and hardwoods, the hemicelluloses in these feedstocks are also known to play a major role in determining the ease of enzymatic deconstruction (Bura et al., 2009; Hu et al., 2011; Kumar and Wyman, 2009; Selig et al., 2009; Várnai et al., 2010). 95 In order to gain insights into the influence of accessibility on enzymatic hydrolyzability, a variety of techniques for quantifying accessibility was applied to the pretreated substrates. Measurements of accessibility were then correlated with the hydrolyzability of the sample, defined here as the minimum amount of protein required to attain 70% glucan conversion after 72 hours at 2% consistency. A commonly used technique for assessing cellulose accessibility and porosity is the Simons’ Staining (SS) technique. Prior to correlating SS values to substrate hydrolyzability, the SS values were first correlated with the glucan content (r ≥ 0.9)of the pretreated samples to ensure that the technique was quantifying the accessible cellulosic fraction of the biomass, rather than the overall accessibility of the total surface (cellulose, hemicellulose and lignin) of the biomass (Figure 4). Figure 4: Glucan content of pretreated biomass samples relative to Simons’ Stain dye ratio. Both the steam- and organosolv-pretreated corn fibre samples (SPCF, OPCF) were excluded from the correlation analysis (adapted from Arantes and Saddler, 2011). In general, the Orange:Blue dye ratio was found to correlate with increasing glucan content in the pretreated structural (corn stalks, hardwood, softwood) biomass samples, as has been reported previously (Chandra et al., 2008a), but not for the corn fibre samples (Figure 4). It is unclear why the OPLPOPPSPCSSPDFSPLPOPCFSPCF01020304050607080900.5 0.75 1 1.25 1.5Glucan Content (%)Simons' Stain (Orange:Blue Ratio)96 steam pretreated corn fibre exhibits an exceptionally high SS ratio, although it is possible that the larger proportion of hemicellulose in this sample (~27%, Table 2) compared to other samples (~1-18%, Table 2) may play a role. In particular, while SS has been shown to exhibit little to no binding to xylan and lignin, binding to arabinan has not been ruled out (Kitamura and Kyoshi, 1971), suggesting that the high arabinan content in the SPCF (~7%, Table 2) may contribute to the discrepancy between the predicted and observed ratio of SS on SPCF. Additionally, the observation that OPCF, which has an arabinan content of only 2%, lies directly on the trendline for SS ratio versus glucan content, lends further support for the hypothesis that the arabinan may be inflating the SS ratio. After determining that the SS dye ratio correlated well with the glucan content of the pretreated structural biomass samples, these SS values were correlated with the minimum protein loading required to achieve 70% conversion (Figure 5). Figure 5: Minimum enzyme loading plotted against Simons’ Stain dye ratio. The corn fibre samples were excluded from the correlation analysis (adapted from Arantes and Saddler, 2011). In general, a higher Orange:Blue ratio, indicative of greater porosity of the biomass, correlated with a decrease in the amount of protein required for 70% conversion (r ≥ 0.95), but this trend did not OPLPOPPSPCSSPDFSPLPOPCFSPCF0102030405060700.6 0.7 0.8 0.9 1 1.1 1.2 1.3 1.4Minimum Protein Loading for 70% Conversion (mg/g glucan)Simons' Stain (Orange:Blue Ratio)97 hold for the corn fibre samples (Figure 5). It is unclear why this trend did not hold for the corn fibre samples, but this may be due to the ‘non-structural’ physiological roles of corn fibre biomass compared to the rest of the biomass samples utilized here (discussed above). This non-structural biomass is much more readily deconstructed (as demonstrated by a far lower enzyme loading required for 70% conversion) than the structural biomass samples used here. The SS Orange:Blue ratio has been shown to represent the accessibility/porosity of the cellulosic fraction of the biomass at the microscopic level (Yu and Atalla, 1998). The strong correlation observed here for the structural biomass samples suggests that, at least for structural biomass, the microscopic porosity/accessibility of the substrate is a key factor in determining the ease of enzymatic hydrolysis. As discussed earlier, we have looked at accessibility at three levels with ‘nanoscopic’ representing the sub-cellulase size scale (i.e. <5 nm), microscopic representing scales larger than cellulase size but invisible to the naked eye (i.e. 5 nm – 100 µm), and macroscopic representing larger phenomenon visible to the naked eye (i.e. >100 µm). The results presented above demonstrate that accessibility at the microscopic level influences the ease of enzymatic deconstruction of biomass. However, these results do not reveal what potential effects macroscopic or nanoscopic accessibility might play in determining the ease of enzymatic deconstruction. In order to determine if macroscopic factors also influence enzymatic deconstruction, a Fibre Quality Analyzer (FQA) was used to determine the average particle size/fibre length of the pretreated biomass samples studied. While there was good correlation between the microscopic accessibility/porosity of the biomass samples and the minimum enzyme loading required for 70% hydrolysis, the correlation between the fibre length and minimum enzyme loading was much weaker (r = 0.8), even when the corn fibre samples were excluded from the regression analysis (Figure 6). The corn fibre samples were excluded from the regression analysis because, as with microscopic accessibility 98 measurements, these samples appeared to be outliers, possibly due to their different physiological roles in the biomass. Attempts to correlate fibre width (another macroscopic fibre property) with ease of enzymatic deconstruction also did not reveal a clear trend, confirming that these macroscopic properties (fibre length and width) do not appear to influence the ease of enzymatic hydrolyzability. Together these results suggest that internal/microscopic accessibility influences enzymatic hydrolysis to a greater extent than do external/macroscopic factors. This trend has recently been reported by Zhang et al. (2013) who performed a literature review of studies addressing the influence of particle size on enzymatic hydrolysis. This work suggested that micron-scale changes in particle size played a more dominant role in determining the ease of enzymatic deconstruction than do changes at the millimeter scale. After assessing the influence of microscopic and macroscopic accessibility on the ease of enzymatic deconstruction, the influence of nanoscopic accessibility was next determined. Figure 6: Minimum enzyme loading required for 70% hydrolysis plotted against the average fibre length. The corn fibre samples were excluded from the correlation analysis (adapted from Arantes and Saddler, 2011). In order to calculate the nanoscopic accessibility of the pretreated biomass fibres, Nitrogen adsorption experiments were performed. Briefly, samples were outgassed under vacuum then subjected to varying pressures of Nitrogen. The amount of Nitrogen adsorbed to the samples at equilibrium for OPLPOPPSPCSSPDFSPLPOPCFSPCF0102030405060700 0.2 0.4 0.6 0.8 1Minimum Protein Loading for 70% Conversion (mg/g glucan)Average Fibre Length (mm)99 each pressure was used to produce a BET plot (Brunauer et al., 1938), which was used to calculate the total specific surface area accessible to Nitrogen gas. Since Nitrogen gas has a Van der Waals radius of ~0.3 nm, the accessibility values for BET calculations, expressed in m2/g, represent the total surface area accessible to a ~0.3 nm diameter probe. In Figure 7, the BET surface area of an assortment of pretreated samples is displayed. Figure 7: Minimum enzyme loading required to achieve 70% conversion plotted against nanoscopic accessibility as measured by Nitrogen BET. The BET surface area calculated for each of the pretreated biomass samples did not reveal a clear trend between the total nanoscopic accessible surface area and the minimum amount of enzyme required to achieve 70% hydrolysis. This lack of correlation is likely explained by the fact that cellulolytic enzymes are generally one order of magnitude larger than the diameter of Nitrogen molecules (i.e. 5-6 nm diameter for cellulases (Abuja et al., 1988; Weimer and Weston, 1985). Thus the Nitrogen BET accessibility measurements will recognize regions of the biomass that are too small for cellulases to access, but large enough for the much smaller gas molecules. This suggests that accessibility at the nanoscopic level does not provide a good indicator of the enzymatic hydrolyzability of the biomass. OPLPOPPSPCSSPDFSPLP0102030405060700 10 20 30 40 50 60 70 80Minimum Protein Loading for 70% Conversion (mg/g glucan)BET Surface Area (m^2/g)100 Since accessibility at the microscopic level (as determined by Simons’ Staining) was found play a more determinant role in enzymatic hydrolyzability than those at the macroscopic or nanoscopic levels, accessibility at the microscopic level was investigated in more detail. Specifically, the maximum amount of cellulase (Celluclast 1.5L) that adsorbed to each of the pretreated substrates was determined (Figure 8). This metric has previously been found to correlate well with the enzymatic hydrolyzability of pretreated substrates (Hogan et al., 1990) and with the available cellulosic surface area (Jeoh et al., 2007). In general, higher maximum protein adsorption was found to correlate with a lower amount of enzyme required for 70% conversion. However, this trend was only observed when comparing substrates pretreated using the same pretreatment technology, with correlation coefficients greater than 0.99 within steam-pretreated samples and within ethanol-organosolv pretreated samples (Figure 8). This suggests that, at least within a subset of biomass samples pretreated using the same pretreatment technology, higher maximum protein adsorption implies that the substrate is more readily hydrolyzable. Figure 8: Minimum protein loading required for 70% conversion is plotted against the maximum protein adsorption for a range of pretreated biomass samples. Correlation coefficients were calculated for samples pretreated with either steam or organosolv pretreatment (adapted from Arantes and Saddler, 2011). OPCFOPLPOPPSPCSSPDFSPLPSPCF0102030405060700 20 40 60 80 100Minimum Protein Loading for 70% Conversion (mg/g glucan)Maximum Protein Adsorption Capacity (mg/g substrate)101 All steam-pretreated samples were found to require a higher amount of enzyme for hydrolysis for a similar maximum protein adsorption capacity when compared to the organosolv-pretreated samples. For example, both steam pretreated and organosolv pretreated lodgepole pine had maximum protein adsorption capacities of ~15-20 mg/g substrate, but the steam pretreated sample required ~60 mg protein/g glucan for effective hydrolysis, while the organosolv-pretreated sample only required around ~40 mg/g glucan. It is unclear exactly why the steam pretreated samples have higher maximum protein adsorption compared to their organosolv-pretreated counterparts. However, this may have to do with differences in the lignin content of the steam versus organsolv-pretreated samples. In general, steam pretreatment only removes a small portion of the lignin from the biomass as well as resulting in the redistribution and condensation of the lignin. This redistributed, condensed lignin is known to interact with enzymes (Chandra et al., 2007; Donaldson et al., 1988; Selig et al., 2007). In contrast, organosolv pretreatments are known to solubilize a significant portion of the lignin (Chandra et al., 2007; Pan et al., 2006). For example, SPLP has a lignin content of ~46%, while OPLP contains ~17% lignin. Thus it is possible that the higher proportion of lignin in the steam-pretreated samples may contribute to the higher maximum protein adsorption in these samples. This suggests that protein adsorption, particularly to steam pretreated samples, yields an artificially high accessibility value, as the enzymes appear to be recognizing not just the cellulosic fraction of the biomass, but also the lignaceous portion. Further evidence for this comes from the fact that the Simon stain, which has been shown not to bind to lignin (Kitamura and Kyoshi, 1971), provides a good correlation across all structural biomass samples studied, regardless of pretreatment technology employed (Figure 5 versus Figure 8). In order to alleviate the effect of non-specific binding of enzymes to lignin during accessibility measurements, blocking of non-specific binding to lignin by incubating the pretreated biomass with BSA prior to protein adsorption was performed during subsequent protein adsorption experiments (Yang and Wyman, 2006). 102 While maximum cellulase adsorption has traditionally been used as a means of determining cellulose accessibility, recent work has demonstrated that these enzymes are capable of promoting changes in the biomass structure within very short time scales (i.e. similar timescales to the equilibration of the enzyme-substrate system during adsorption) (Gao et al., 2014). Thus, using cellulase adsorption to biomass as a tool for assessing accessibility will not provide a fully representative description of the biomass surface due to alterations occurring in the biomass structure induced by the enzymes over the course of adsorption. Although performing the adsorption studies at 4 °C can be used to reduce this catalytic activity, some residual catalysis does still occur at these temperatures. Therefore, for subsequent adsorption studies, BSA blocking was used to reduce non-specific adsorption to lignin, and isolated CBMs were used as probes for the cellulose, rather than using the full catalytically-active enzymes. This technique is discussed in much more detail in Section 3.2 of this thesis, but the preliminary data is described here. 3.1.4 CBM adsorption as a tool for quantifying accessibility In order to test the ability of CBMs to describe the cellulose accessibility and enzymatic hydrolyzability of pretreated biomass, a range of steam-pretreated biomass samples was prepared, including agricultural residues (corn stover), hardwoods (poplar), and softwoods (lodgepole pine). Steam pretreatment was chosen for these samples because the majority of the commercial facilities utilizing enzymatic deconstruction of lignocellulosic biomass are utilizing this technology (Brown and Brown, 2013; Kazi et al., 2010), suggesting that steam pretreatment may be a potential leader in pretreatment technologies for bioconversion. The pretreatment conditions and biomass compositions after pretreatment are shown in Table 4. When using the CBM adsorption technique, fresh pretreated substrates were prepared. These substrates were different from those used in previous work involving the ANOVA modelling used to 103 determine the minimum enzyme loading required to achieve 70% conversion. Enzymatic hydrolysis efficiency was therefore plotted as the percent conversion after 48 hours, rather than minimum enzyme loading for 70% conversion. Table 4: Pretreatment conditions and substrate compositions for the biomass samples used in the CBM adsorption experiments. Substrate Pretreatment conditions Major sugar composition (%) Lignin (%) Abbreviation SO2a-steam pretreatment Glub Xylc Mand AILe ASLf Corn stover 190°C, 5 minutes, 3% SO2 50.7 ± 0.9 15.4 ± 0.1 0.7 ± 0.0 23.3 ± 0.3 3.4 ± 0.2 SPCS Poplar 200°C, 5 minutes, 3% SO2 58.2 ± 0.2 7.7 ± 0.1 2.0 ± 0.1 28.5 ± 0.2 2.8 ± 0.1 SPP Lodgepole pine 200°C, 5 minutes, 4% SO2 53.4 ± 1.0 0.2 ± 0.0 0.4 ± 0.1 42.8 ± 0.2 0.6 ± 0.0 SPLP aSulfur dioxide, bGlucan, cXylan, dMannan, eAcid-insoluble lignin, fAcid-soluble lignin The CBMs used as probes for the cellulose in this work included CBM2a from C. fimi and CBM44 from C. thermocellum. CBM2a is a Type A CBM which preferentially binds to crystalline regions of cellulose via a hydrophobic planar binding site (McLean et al., 2000, 2002), while CBM44 is a Type B CBM which preferentially recognizes amorphous cellulose via a cleft-shaped binding site capable of binding only isolated individual glucan chains (Najmudin et al., 2006). As described in the methods section, the values for CBM adsorption do not represent the saturating amount of CBM binding to each substrate, but rather represent the amount of CBM bound after incubating the substrates with a fixed concentration of CBM (in this case, 250mg/g substrate). For a detailed description of this technique and its applications, see Section 3.2. The adsorption of CBM2a and CBM44 to the steam-pretreated samples is shown in Figure 9. In general, CBM2a bound to a greater extent than did CBM44 to all of the samples tested. The CBM2a 104 bound most to the corn stover sample, followed by poplar then lodgepole pine, while the amount of bound CBM44 varied little between samples. The correlations between binding of the CBMs and enzymatic hydrolyzability are shown in Figure 10. Figure 9: Adsorption of CBM2a and CBM44 to steam-pretreated biomass samples. Samples were run in triplicate and error bars represent one standard deviation from the mean. Figure 10: Conversion of three steam pretreated samples plotted against the amount of bound CBM2a (squares), CBM44 (triangles) and the sum of both CBMs (diamonds). 050100150200250SPCS SPP SPLPmg CBM/g SubstrateCBM2aCBM44SPCSSPPSPLPSPCSSPPSPLPSPCSSPPSPLP0%10%20%30%40%50%60%70%0 50 100 150 200 250 300 350% Glucan Conversion After 48hmg CBMs/g SubstrateCBM2aCBM44Sum CBMs105 The hydrolyzability of the steam pretreated biomass samples was found to correlate with the amount of CBMs bound to each sample. While the binding of each individual CBM appeared to provide a relatively good indicator of enzymatic hydrolyzability (r ≥ 0.90 for CBM44 and r ≥ 0.95 for CBM2a), the best correlation was found between hydrolyzability and the sum of the binding of the two CBMs (r ≥ 0.99). These results reaffirm that the accessibility of the cellulose to cellulases at the microscopic level is likely one of the key factors determining the enzymatic hydrolyzability of pretreated biomass samples and demonstrates that CBMs can be used as accurate probes for the amount of cellulose accessible to cellulases. While the total amount of each CBM binding to the cellulose can be considered a measure of ‘microscopic accessibility’ (as defined by this work), the ratio between the amounts of each CBM bound can be considered a measure of nanoscopic surface changes. By comparing the amount of each different CBM bound to a given sample, insights into nanoscopic surface features can be gained. Specifically, while each CBM is approximately the same size as a cellulase, the difference between binding sites for CBM2a (a crystalline flat face (McLean et al., 2000, 2002)) and CBM44 (an isolated cellulose chain (Najmudin et al., 2006)) can be considered as nanoscopic features. CBM adsorption measurements are thus capable of providing not just the microscopic accessibility of the biomass (based on the total amount of the CBMs bound), but also of providing insights into nanoscopic features in the cellulosic substructures at the surface of the biomass fibres. Overall, the results demonstrated that the microscopic accessibility of pretreated lignocellulosic substrates plays a major role in determining enzymatic hydrolyzability. As discussed previously, thermochemical pretreatments will always be performed at compromised pretreatment conditions in order to open up the biomass without using excessive chemical or energy inputs, and without promoting the formation of sugar degradation products. It is therefore left up to the enzymes involved in biomass 106 deconstruction to perform the opening up and disruption of the remaining fibrillar organization of the pretreated biomass (Arantes and Saddler, 2010). Although enzymes such as hydrolases and lytic polysaccharide mono-oxygenases have been shown to erode cellulosic structures from the surface relatively slowly (Chanzy and Henrissat, 1985; Väljamäe et al., 1998), it has been proposed that non-hydrolytic, non-oxidative proteins may be capable of directly opening up the structural organization of the biomass, thereby increasing the amount of surface area accessible to the hydrolases and oxidative proteins (Reviewed in (Arantes and Saddler, 2010)). The ability of some of these amorphogenesis-inducing enzymes to open up biomass at the microscopic level suggests that this enhancement of accessibility by disruptive proteins has potential to greatly enhance the efficiency of enzymatic biomass deconstruction. 3.1.5 Conclusions Cellulose accessibility at the microscopic level (5 nm – 100 µm) was shown to play a major role in determining the ease of enzymatic deconstruction of pretreated lignocellulosic biomass. Conversely, accessibility at the macroscopic (>100 µm) or nanoscopic (<5 nm) levels did not appear to significantly influence ease of enzymatic deconstruction. Techniques such as Simons’ Stain, cellulase adsorption, and CBM adsorption, which measure the microscopic cellulose accessibility within pretreated biomass, provided measurements of accessibility that correlated better with ease of enzymatic deconstruction than techniques which measured either macroscopic (FQA) or nanoscopic (Nitrogen BET) accessibility. This suggests that structural factors at organizational scales close to the sizes of cellulolytic enzymes (~5-6 nm) play a greater role in determining enzymatic hydrolyzability than those at larger or smaller scales. The CBM adsorption experiments performed here indicated that CBM adsorption can be used to accurately predict the enzymatic hydrolyzability of a range of pretreated lignocellulosic biomass. This technique can be used to assess nanoscale-features on the cellulose substrate, as well as the overall 107 accessibility of the cellulose at scales relevant to enzymatic deconstruction. This ability, combined with the ease and simplicity of performing this assay, make CBM adsorption an ideal technique for further work involving quantification of the disruptive effects of non-hydrolytic/non-oxidative disruptive proteins. 3.2 Development and application of a new technique for quantifying amorphogenesis 3.2.1 Background One way to lower the costs and improve the efficiency of the bioconversion process is to reduce the amount of enzyme required for cellulose hydrolysis. By exploiting the ability of amorphogenesis-inducing proteins to enhance the accessibility of the biomass to the deconstructing enzymes, the overall efficiency of the enzyme mixture can be improved, thereby lowering the amount and cost of the enzymes required. However, in order to effectively apply these proteins to the deconstruction process, it is necessary to first develop a good understanding of precisely what effect these proteins are having on the biomass. Previous attempts to understand the manifestations of amorphogenesis induced by these disruptive proteins at the multiple levels of substrate organization have made use of a suite of qualitative or semi-quantitative techniques. As discussed in Section 1.3, these have included visual analysis of macroscopic aggregate dispersion (Jäger et al., 2011; Saloheimo et al., 2002), as well as particle size reduction (Chen et al., 2010; Jäger et al., 2011; Wang et al., 2010) and extensibility measurements (Baker et al., 2000; Cosgrove, 2000; Kerff et al., 2008; Lee et al., 2010; McQueen-Mason et al., 1992; Tabuchi et al., 2011) to qualify macroscopic disruption. At the microscopic level, extensive use of microscopy was applied to directly visualize cell wall disruption, and image-analysis software was used to analyze microscopic particle release/dispersion (Banka et al., 1998; Ciolacu et al., 2010; Din et al., 1991, 1994; Gilkes et al., 1993; Jäger et al., 2011; Pinto et al., 2004; Quiroz-Castañeda et al., 2011; 108 Saloheimo et al., 2002; Tomme et al., 1995). Finally, at the nanoscopic level, AFM has been used to observe direct disruption of microfibril surfaces (Lee et al., 2000), and crystallinity measurements have been used to quantify changes in the cellulose crystal structure after amorphogenesis (Chen et al., 2010; Ciolacu et al., 2010; Gao et al., 2001; Wang et al., 2008). The enhancement of substrate hydrolyzability has also been used as an indirect measure of substrate disruption (Baker et al., 2000; Lee et al., 2010; Quiroz-Castañeda et al., 2011). Overall, these attempts have been qualitative or semi-quantitative in nature and have failed to provide an accurate, reproducible metric for quantifying substrate disruption promoted by amorphogenesis-inducing proteins at the various levels of organization within the plant biomass. As shown earlier, adsorption of these CBMs to lignocellulosic biomass provided a good indicator of enzymatic hydrolyzability. Since these CBMs can preferentially bind to specific substructures of the cellulose (Fox et al., 2013; McLean et al., 2002), it is possible that CBM adsorption could be used to not only quantify general changes in microscopic cellulose accessibility, but also to distinguish subtle changes occurring at the surface of the cellulosic fibrils. The CBMs employed were the Type A CBM, CfCBM2a, which preferentially binds to crystalline cellulose via a planar hydrophobic binding face (McLean et al., 2000, 2002), and the Type B CBM, CtCBM44, which preferentially binds to amorphous cellulose via a cleft-shaped binding groove (Najmudin et al., 2006). Due to the fundamentally different architectures of the cellulose binding sites of these two CBMs (i.e. the planar face of CBM2a vs the cleft shape of CBM44), these two CBMs can be used to distinguish between crystalline bundles of cellulose, recognized by CBM2a, and individual cellulose chains recognized by CBM44. Because of the cleft-shaped structure of the binding site of CBM44, only isolated chains or those that are loosely associated with the bulk of the cellulose bundle 109 are bound by this CBM (Najmudin et al., 2006). Representative crystal structures of a Type A and a Type B CBM are shown in Figure 11, below. Figure 11: Representative crystal structures of a Type A CBM (Left) and a Type B CBM (Right). The CBMs illustrated here are the Family 1 CBM from T. reesei Cel7A (Left) (PDB code 1CBH; (Kraulis et al., 1989), and the Family 4 CBM from C. fimi endo-1,4-glucanase C (Right) (PDB code 1GU3; (Boraston et al., 2002b). Reproduced from (Guillén et al., 2010). Although CBM44 has been used to quantify amorphous cellulose, CBM44 has also been shown to bind with equal affinity to both cellulose and xyloglucans (Najmudin et al., 2006). This accommodation of xyloglucan side chains appears to be a common theme within the CBMs that recognize isolated cellulose chains (i.e. Type B cellulose-binding CBMs) (Najmudin et al., 2006). Fortunately, xyloglucans are only present in significant quantities during the growing (primary) cell wall, where they play a key role in regulating cell elongation (Hayashi, 1989; Takeda et al., 2002). Xyloglucan content has been shown to dramatically decrease in mature (secondary) cell walls, which make up the bulk of plant biomass, leaving only very minor amounts of xyloglucan in the pretreated lignocellulosic substrates employed in this work (Gomez et al., 2008; Hayashi, 1989; Vogel, 2008). Thus the Type B CBM44 is a suitable probe for quantifying the amount of amorphous cellulose present in the pretreated biomass and model cellulosic substrates used in this work, but would be a more problematic probe for quantifying amorphous cellulose in primary cell walls enriched in xyloglucan. 110 It should be noted here that CBMs have previously been referred to as being “crystalline cellulose specific” or “amorphous cellulose specific” (Filonova et al., 2007a; Jamal et al., 2004). However, it is becoming apparent that cellulose in nature exists in a spectrum of crystallinities (Langan et al., 2014), and that CBMs have a range of affinities for more- and less-ordered regions of cellulose (Fox et al., 2013). Specifically, recent XRD data has revealed that the crystalline cores of elementary fibrils are only 2-3 nm in diameter, while the fibril itself is 3-4 nm in diameter. This implies that there exists a crystalline core within the center of the elementary fibrils and that there is a progressive loss in crystallinity from the center toward the outside of the fibrils (Langan et al., 2014). Additional evidence for this ‘spectrum of crystallinities’ theory comes from experiments using phosphoric acid to reduce the crystallinity of Avicel (Hall et al., 2010). In this work, it was found that there was a linear correlation between Avicel crystallinity and the concentration of acid used to reduce the crystallinity (between 75-80% acid only). This linear correlation suggests that, at the low end of the acid concentrations, some cellulose regions that are slightly crystalline are being converted to amorphous cellulose, but more crystalline regions of the cellulose are harder to open up and thus retain their crystalline structure. This range of concentrations of acid leading to a range of Avicel crystallinities suggests that there is a spectrum of cellulose crystallinities within Avicel. In other words, cellulose does not have distinct populations of ‘crystalline’ and ‘amorphous’ cellulose, as it appears that there is a continuum of crystallinities between the most crystalline regions at the core of elementary cellulose fibrils and the most amorphous regions (Langan et al., 2014). However, in order to simplify terminology for the purposes of this work, cellulose has been categorized binarily into either `more crystalline cellulose’, which is recognized by CBM2a, or ‘less crystalline cellulose’ which is recognized by CBM44. As the binding face of CBM2a is only a few nanometers wide, this binding face recognizes associations of just 2-3 adjacent cellulose chains (McLean et al., 2000, 2002). For the purposes of this 111 work, ‘crystalline’ cellulose is thus defined as associations of at least 2-3 parallel cellulose chains. This definition of crystalline cellulose is likely less stringent than traditional definitions of crystalline cellulose. This is because the techniques traditionally used for measuring crystallinity make use of various diffraction/spectroscopy techniques which require a regular repeating structure in order to generate ‘crystalline’ diffraction spectra. For example, lateral association of 2-3 cellulose chains over a length of ~5 nm (the length of CBM2a) could theoretically be recognized as crystalline cellulose by CBM2a, whereas this region would not be recognized as crystalline cellulose using techniques such as XRD or FTIR. Additionally, while traditional diffraction techniques require multiple sheets of cellulose within a bundle in order to produce a crystalline cellulose diffraction spectrum, CBM2a requires only one sheet containing a lateral association of cellulose chains. Thus it should be noted our definition of ‘crystalline cellulose’ is quite “loose”, and likely recognizes a significant amount of ‘paracrystalline’ or ‘microcrystalline substructures’ found within the biomass which are recognized by CBM2a but which would not be recognized as crystalline using traditional diffraction techniques. Despite the relatively broad specificity of CBM2a, the fundamental difference in binding site architecture between this CBM and CBM44, which has a cleft-shaped binding site capable of recognizing individual glucan chains (Najmudin et al., 2006), was expected to provide an accurate representation of the surface morphology of the cellulose within plant cell walls. Adding support to this theory, competitive binding experiments on phosphoric acid swollen cellulose (PASC) have been used to demonstrate that CBM2a has little binding site overlap with a Type B CBM (CBM4-1) which is known to preferentially recognize the amorphous regions of cellulose while showing negligible affinity for crystalline regions (McLean et al., 2000, 2002). An additional fundamental difference between the CBM adsorption technique presented here and traditional crystallinity measurements is that these traditional techniques measure a bulk property 112 of the cellulose (i.e. a 3-dimensional parameter), whereas CBM adsorption measures a surface phenomenon (i.e. a 2-dimensional parameter). The enzymes involved in cellulose deconstruction predominantly act at the surface of cellulosic structures (particularly in the case of the non-complexed cellulase enzymes employed in this work) (Liu et al., 2011; Resch et al., 2013; Teeri, 1997; Väljamäe et al., 1998). Thus, the ability of CBM adsorption to specifically quantify cellulose properties at surfaces where these enzyme-substrate interactions are occurring, suggests that this technique could be used to gain detailed insights into the effects of cellulose surface morphology during enzymatic hydrolysis. Overall, the CBM adsorption technique was developed not so much as a tool for measuring cellulose crystallinity, but rather to provide detailed information on cellulosic structures, combining crystallinity, accessibility, and surface morphology factors into a single metric for quantifying amorphogenesis and other enzyme-mediated changes in cellulosic structures. The CBM adsorption technique described here was first assessed for its ability to differentiate between crystalline and amorphous cellulose when quantifying a range of chemically disrupted model cellulosic substrates exhibiting various degrees of amorphogenesis. The ability of CBM adsorption to predict enzymatic hydrolyzability of these substrates was then investigated. After confirming the applicability of CBM adsorption to quantifying amorphogenesis, this technique was applied to track changes in cellulose architecture occurring during enzymatic deconstruction of cellulose, as well as after treatment with the amorphogenesis inducing protein, Swollenin. 3.2.2 Measuring cellulose accessibility 3.2.2.1 Adsorption of Simons’ stain to model cellulosic substrates Prior to applying the CBM adsorption technique to quantifying amorphogenesis, the ability of the CBMs to accurately quantify the accessibility and surface morphology of a range of model cellulosic substrates was assessed. Specifically, CBM adsorption measurements were compared to accessibility 113 measurements obtained using the Simons’ Stain technique (Chandra et al., 2008a) for model celluloses, including relatively amorphous celluloses such as Cellulose II and PASC, more ordered celluloses including dissolving pulp (DSP) and Avicel PH-101, as well as highly crystalline/ordered cellulose nanocrystals. The binding of the Direct Orange (DO) Simons’ stain dye and each CBM to the range of model cellulosic substrates is shown in Figure 12. Figure 12: Adsorption of Simons’ Stain DO dye to a range of model cellulosic substrates. The least ordered substrates (PASC and Cellulose II) were found to adsorb more dye than other substrates. CNC and Cellulose III adsorbed the least dye. PASC: Phosphoric Acid Swollen Cellulose, DSP: Dissolving Pulp, CNC: Cellulose Nanocrystals. The application of the Simons’ staining technique to this range of model cellulosic substrates revealed that, as expected, a decrease in the general structural organization of the biomass resulted in an increase in the accessibility of the cellulose (as measured here by the adsorption of the Direct Orange dye (Chandra et al., 2008a)). Previous work has demonstrated that Cellulose III tends to adsorb far less enzyme than other celluloses during enzymatic hydrolysis (Chundawat et al., 2011; Igarashi et al., 2007). It is not yet known 0204060Cellulose III Cellulose II PASC DSP Avicel CNCmg Dye Bound/g Glucan114 precisely why Cellulose III adsorbs less enzyme, although the fact that the crystal lattice is distinctly different from Cellulose I and II could play a role in this phenomenon (Chundawat et al., 2011). This reorganized crystal lattice appears to present a binding face which is not recognized by cellulases or CBMs. This could be due to the lack of Cellulose III in nature, resulting in a lack of evolutionary pressure for enzymes to recognize this novel structure. The greatly reduced binding of the DO dye to Cellulose III compared to other cellulosic substrates has not previously been observed, and implies that whatever factors are involved in reducing the enzyme-binding to Cellulose III probably also influence the binding of the DO dye. Despite the low levels of adsorption of the DO dye and cellulases, Cellulose III has been shown to be rapidly hydrolyzed by cellulases (Chundawat et al., 2011; Igarashi et al., 2007), contrary to the currently accepted theory that enzyme adsorption is a good indicator of enzymatic hydrolyzability of a substrate. It has been suggested that the increased susceptibility of Cellulose III to enzymatic depolymerisation could be due to reduced energy required to abstract individual cellulose chains from the surface of the cellulosic fibrils (Beckham et al., 2011). While this could explain the increased susceptibility of Cellulose III to enzymatic deconstruction, it is still not clear how the enzymes are capable of rapidly depolymerizing this cellulose without first adsorbing to the surface (Beckham et al., 2011; Chundawat et al., 2011; Igarashi et al., 2007). This discrepancy is interesting and warrants further research, but is outside the scope of this thesis. The relationship between cellulose accessibility, as measured by DO dye adsorption, and enzymatic hydrolyzability is shown in Figure 13. In general, an increase in dye adsorption correlated with an increase in the enzymatic hydrolyzability of the cellulose (r ≥ 0.95). This trend held for all cellulose samples analyzed except for Cellulose III, which hydrolyzed better than either Avicel or Dissolving pulp 115 while adsorbing dramatically less dye. As Cellulose III has previously shown to be an outlier in the amount of cellulase it adsorbs, this sample was excluded from the trendline calculated in Figure 13. Figure 13: Percent conversion after 48 hours for a range of model cellulosic substrates plotted against adsorption of Simons’ Stain DO dye. In general, the more disordered celluloses adsorbed more dye and hydrolysed more readily. The Cellulose III data point was considered an outlier and was excluded from the correlation analysis. PASC: Phosphoric Acid Swollen Cellulose, DSP: Dissolving Pulp, CNC: Cellulose Nanocrystals. 3.2.2.2 Adsorption of CBM2a and CBM44 to dissolving pulp To try to determine if CBM2a and CBM44 were indeed recognizing distinct binding sites on the cellulosic substrates, Scatchard plots were used to calculate the maximum adsorption of the CBMs to dissolving pulp. Dissolving pulp was chosen as it is used extensively throughout this thesis as a model cellulosic substrate and because it contains both crystalline and amorphous cellulose features. To try to determine if there was overlap in the CBM2a and CBM44 binding sites, the CBMs were bound to dissolving pulp over a range of different CBM loadings. CBMs were added both individually and in a 50:50 mixture at the same total CBM loading as employed for the individual adsorption experiments (Figure 14). The equation Cellulose III01020304050607010 20 30 40 50 60 70Conversion (%)mg Dye/g cellulose 116 [] []  =  ×  −  × [] was used to represent the plots, where [LE] is the amount of bound enzyme per milligram cellulose, [E]free is the concentration of free enzyme, Emax is the maximum protein adsorption, and Ka represents the slope. Using this equation, the Emax values were calculated for CBM2a alone (35.4 µg/mg), CBM44 alone (62.1 µg/mg), and the 50:50 mixture of CBM2a and CBM44 (81.8 µg/mg) (Figure 14). It was apparent that the Emax value for the mixture of the two CBMs was higher than that of either individual CBM, implying that the two CBMs recognized distinct binding sites (potentially with some degree of overlap between the pools) on the cellulose surface. This result is consistent with the different binding site architecture between the two CBMs (Type A versus Type B). Figure 14: Scatchard plots of CBM2a and CBM44 bound to dissolving pulp. Best fit lines were calculated for the linear sections of the plots only. Emax values represent the maximum amount of CBM adsorbed at equilibrium per mg of cellulose. y = -0.1538x + 5.4452R² = 0.9911Emax = 35.40.00.51.01.52.02.515 35 55Bound CBM/Free CBMBound CBM (µg)CBM2ay = -0.0296x + 1.8372R² = 0.9679Emax = 62.10.00.20.40.60.81.01.21.40 100 200Bound CBM (µg)CBM44y = -0.0234x + 1.9148R² = 0.9898Emax = 81.80.00.20.40.60.81.01.21.41.61.80 50 100 150Bound CBM (µg)CBM2a + CBM44117 3.2.2.3 Adsorption of CBMs to model cellulosic substrates After correlating cellulose accessibility measured by DO dye adsorption with the enzymatic hydrolyzability of a range of model cellulosic substrates, the adsorption of CBM2a and CBM44 to these substrates was next assessed (Figure 15). In general, samples which bound more DO dye also bound more CBM, and the least ordered substrates (PASC and Cellulose II) were found to adsorb more CBM than other substrates. These disordered substrates also had the lowest ratio of bound CBM2a to bound CBM44, indicating a more amorphous cellulosic surface (Figure 16). The CNC had the highest ratio of CBM2a:CBM44 adsorbed. As anticipated, the more disordered samples (Cellulose II and PASC) were found to have the lowest CBM2a:CBM44 ratio, while more crystalline/ordered samples such as CNC had the highest ratio of CBM2a:CBM44 (Figure 16). The Cellulose III adsorbed negligible amounts of either CBM, and was thus excluded from the CBM2a:CBM44 ratio calculation. Figure 15: Adsorption of CBMs to a range of model cellulosic substrates. Samples were run in triplicate and error bars represent one standard deviation from the mean. PASC: Phosphoric Acid Swollen Cellulose, DSP: Dissolving Pulp, CNC: Cellulose Nanocrystals. 0204060Cellulose III Cellulose II PASC DSP Avicel CNCmg CBM Bound/g GlucanCBM2a CBM44118 Figure 16: Ratio of CBM2a to CBM44 adsorbed to a range of model cellulosic substrates. Samples were run in triplicate and error bars represent one standard deviation from the mean. PASC: Phosphoric Acid Swollen Cellulose, DSP: Dissolving Pulp, CNC: Cellulose Nanocrystals. In general, an increase in adsorption of either of the CBMs correlated well with an increase in the enzymatic hydrolyzability of the cellulose (Figure 17). However, as with DO dye adsorption, Cellulose III was found to be an outlier, adsorbing negligible amounts of each CBM while still hydrolyzing relatively well. This lack of adsorption and ease of hydrolysis of Cellulose III has been reported previously (Chundawat et al., 2011; Igarashi et al., 2007). In the case of CBM adsorption, the cellulose nanocrystals (CNC) also appeared as an outlier when plotting hydrolyzability against the amount of adsorbed CBM. Specifically both CBMs adsorbed to a greater extent to CNC than was expected, based on the best fit line produced from plotting adsorption against hydrolyzability for the other model cellulosic substrates (Figure 17). This discrepancy can likely be explained by the nanoscopic particle size of CNC. Since CNC derived from wood has particle sizes in the nanometer scale (~100-200 nm x ~10 nm (Habibi et al., 2010)), the overall surface area of this cellulose is actually quite high, leading to increased adsorption of the CBMs. 012345Cellulose II PASC DSP Avicel CNCCBM2a:CBM44 Ratio119 Figure 17: Percent conversion after 48 hours of a range of model cellulosic substrates plotted against CBM adsorption (r ≥ 0.95 for each CBM). Adsorption of CBM2a is represented by diamonds while CBM44 is represented by squares. Filled shapes represent those included in the correlation analysis, while empty shapes represent outliers. Samples were run in triplicate and error bars represent one standard deviation from the mean. Regarding CBM44 in particular, although this CBM bound the least to CNC out of all the celluloses tested, binding was still higher than was predicted based on the best fit line. Due to the highly-crystalline nature of CNC, this result suggests that CBM44 may be capable of binding to certain cellulose chains at the peripheral edges/corners of the crystalline cellulose bundles. For the case of CBM2a, CNC adsorbed more CBM2a than either Avicel or dissolving pulp, despite having the lowest hydrolysis efficiency. However, the exceptional crystallinity of CNC suggests that essentially all of the accessible surface area measured by CBM2a would be present as highly crystalline cellulose. Thus, at least for the case of exceptionally crystalline pure cellulose, at the nanoscopic level of the individual elementary fibrils which comprise CNC, the crystallinity of the cellulose may influence the hydrolyzability of the substrate (Hall et al., 2010). Cellulose IIICNCCNC0102030405060700 10 20 30 40 50 60 70Cellulose Conversion (%)mg CBM bound/g CelluloseCBM2aCBM44CBM2aCBM44120 Overall these results indicated that CBM adsorption could be used to gain valuable insights into the complex relationships between cellulose surface morphology and enzymatic hydrolyzability. Additionally, this confirmed the ability of the two-probe CBM technique to accurately differentiate between crystalline and amorphous cellulose. This indicated its applicability as a tool for quantifying amorphogenesis by measuring the overall cellulose accessibility (combined adsorption of both CBMs) as well as changes in the nanoscopic properties of the cellulose surface morphology. 3.2.3 Quantification of acid-induced cellulose disruption 3.2.3.1 CBM adsorption As protein-mediated amorphogenesis is still an evolving concept, we initially assessed the sensitivity and reproducibility of a CBM-mediated method for quantifying changes in cellulose accessibility/surface morphology. We did this by using concentrated phosphoric acid to artificially produce a range of celluloses representing varying degrees of ‘amorphogenesis’, in a manner similar to the work performed by Zhang et al. (2006). The use of harsh acid treatments was intended to provide an exaggerated range of disrupted substrates and was not intended to be representative of milder, biological treatments. Previous work on the effect of phosphoric acid on cellulosic biomass has shown that, at acid concentrations below ~75%, cellulose is swollen slightly during the acid treatment. However, at acid concentrations above 75%, a dramatic dissolution of the cellulose is observed (Zhang et al., 2006). In this work, the disruptive effect of the acid was initially qualitatively assessed using SEM (Figure 18) followed by a quantitative assessment, where the adsorption of each of the substructure-specific CBMs was determined (Figure 19). 121 Figure 18: Scanning electron microscopy of cotton fibres after treatment with a range of o-phosphoric acid concentrations. All images were taken at x1200 magnification and each image depicts a representative fibre for the indicated acid concentration. 122 Figure 19: Adsorption of crystalline cellulose-binding CBM2a and amorphous cellulose-binding CBM44 to cotton fibres treated with a range of o-phosphoric acid concentrations. Experiments were run in triplicate and error bars represent one standard deviation from the mean. In order to progressively disrupt cotton fibres to assess the ability of CBM adsorption to quantify cellulose accessibility, a range of phosphoric acid concentrations was used. After control treatment (nanopure water, 0% (w/w) acid), cotton fibres appear smooth, with few surface features. As the acid concentration was increased to near the point of cellulose dissolution (~73%-78%), evidence of substrate amorphogenesis was apparent at the surface of the cotton fibres. At 74% phosphoric acid, initial signs of splitting, roughening, fibrillation and peeling/delamination of the fibres began to appear. As the acid concentration was increased from 74% to 76%, these effects became more pronounced. At 77%, the fibre structure was almost completely destroyed, with large portions of the outer layer of the fibre appearing to peel off, revealing a rough, fibrillated underlying structure. After treatment with 78% phosphoric acid no fibre structure remained and all the cellulose appears to have been dissolved and reprecipitated into amorphous cellulosic ‘mats’. 01020304050607073 74 75 76 77 78mg CBM Bound/g Cotton% Phosphoric Acid (w/w)CBM44CBM2a123 An increase in phosphoric acid-mediated disruption generally correlated with an increase in binding of both the amorphous-binding and crystalline-binding CBMs up to a concentration of 77% acid treatment (Figure 19). This increase in the combined binding of both CBMs provided a good indication that the overall (crystalline and amorphous) cellulose accessibility had increased. It had been anticipated that the increasing disruption of the fibres would result in the amorphous-binding CBM44 being bound more than the crystalline-binding CBM2a. However, as the acid concentration was increased up to 77%, the binding of CBM2a actually increased more dramatically than that of CBM44 (Figure 19). Since CBM2a recognizes only two to three adjacent chains as being ‘crystalline’ (McLean et al., 2000, 2002), the observed relative increase in CBM2a binding over CBM44 binding as the acid concentration increased was likely due to the increased solvent exposure of small microcrystalline substructures within the acid-disrupted cotton fibres. As the acid concentration was further increased from 77% to 78%, the SEM micrographs of the cotton showed that any remaining residual fibre structure was lost and that the substrate now had the form of amorphous cellulosic ‘mats’ (Figure 18). These SEM observations complemented the CBM adsorption results, where increasing the acid concentration from 77% to 78% resulted in a large increase in the amount of adsorbed CBM44, without significantly altering the adsorption of CBM2a (Figure 19). It was apparent that the specificity of CBM adsorption was distinct enough that changes in the surface morphology of the cellulosic substrates could be readily differentiated. 3.2.3.2 Relationship between initial hydrolysis rate and CBM adsorption After determining that CBM adsorption could be used to quantify acid-induced changes in cellulose accessibility, the degree of substrate disruption (quantified by CBM adsorption) was correlated with initial hydrolysis rates. Rather than correlating amorphogenesis with the conversion extent after 48 124 hours, the initial hydrolysis rate (within the first 30 minutes) was used to get a better idea of how amorphogenesis might directly influence initial hydrolysis. Each of the phosphoric-acid disrupted cotton fibre samples was hydrolyzed for 30 minutes using a commercial cellulase mixture (30 filter paper units/g cellulose of Celluclast 1.5 L, supplemented with 15 cellobiase units/g cellulose of Beta-glucosidase (Novozym 188, Novozymes A/S, Bagsværd, Denmark)). The initial hydrolysis rate (defined here as the hydrolysis rate over the first 30 minutes of the reaction) was plotted against the adsorption of each individual CBM, as well as the average adsorption of the two CBMs (Figure 20). The adsorption of each individual CBM, and particularly their average adsorption, was found to correlate well with the enhanced enzymatic hydrolyzability of the cotton fibres (r ≥ 0.95). The apparent linear correlation between the cellulose accessibility (as measured by CBM adsorption) and the initial hydrolysis rates (over the first 30 minutes) suggests that this hydrolysis reaction, at least for the conditions tested here, proceeds via a first-order reaction, where the hydrolysis rate depends linearly on the amount of accessible cellulose. Additionally, the steeper slope of the line for CBM44 (m = 35) when compared to CBM2a (m = 25) seems to indicate that, at least for the initial stages of hydrolysis, enzymatic hydrolysis rates were influenced more by the amount of accessible amorphous cellulose than they were by the amount of accessible crystalline cellulose. While both the hydrolyzability and CBM44 adsorption increased with every incremental increase in phosphoric acid concentration, this was not the case for CBM2a. As the acid concentration was increased from 77% to 78%, the adsorption of CBM2a to the substrate was not significantly affected, even as the hydrolyzability continued to increase. This suggested that attempts to correlate changes in substrate accessibility to hydrolyzability using a specific mono-component cellulase may be problematic, as some cellulases contain CBMs specific for crystalline cellulose and might therefore underestimate the accessibility of the highly amorphous regions of the substrate. 125 Figure 20: Initial hydrolysis rate vs CBM adsorption. Initial hydrolysis rate (calculated after 30 minutes of hydrolysis) of acid-disrupted cotton fibres increased with increasing CBM adsorption. Each data point represents a cotton fibre sample treated with a different concentration of o-phosphoric acid and hydrolyzed with the same enzyme loading. Experiments were run in triplicate and error bars represent one standard deviation from the mean. Overall, these results indicated that substructure specific CBMs could be used to quantify acid-induced changes in cellulose accessibility and that increases in accessibility (as determined by CBM adsorption) could provide a good predictor of initial rates of enzymatic hydrolysis. 3.2.4 Tracking changes in cellulose morphology over the course of hydrolysis Since the initial hydrolysis rate appeared to be more strongly correlated with the amount of accessible amorphous cellulose, the relationship between cellulose surface morphology and enzymatic hydrolysis was investigated further. Specifically, dissolving pulp was hydrolyzed using a commercial cellulase mixture (Celluclast 1.5L, as described above) and changes in the surface morphology were tracked by CBM adsorption. It was predicted that the amorphous cellulose should be hydrolyzed more R² = 0.9828R² = 0.9728 R² = 0.997901002003004005006007008000 5 10 15 20 25 30 35 40Initial Hydrolysis Rate ug/mL Glucose/hourmg CBM bound/g CottonCBM2aCBM44Average of both CBMs126 rapidly than the crystalline cellulose which should lead to a decrease in the adsorption of CBM44 to the cellulose over the course of hydrolysis. When the amount of each CBM bound to the pulp over the course of a 6 hour hydrolysis was assessed (Figure 20), the CBM adsorption profiles on the variably hydrolyzed dissolving pulps showed a rapid and dramatic reduction in the amount of accessible amorphous cellulose, which was reduced by 50% within the first 15 minutes of hydrolysis. The amount of accessible amorphous cellulose remained at this level for the remainder of the hydrolysis course. The amount of accessible crystalline cellulose also declined, but at a slower rate and to a lesser extent than the decline in accessible amorphous cellulose (Figure 21). Figure 21: Adsorption of CBM2a (squares, solid line) and CBM44 (triangles, dashed line) over the course of a 6 hour hydrolysis, plotted alongside glucose release (diamonds, dotted line). A rapid drop in the amount of accessible amorphous cellulose occurred within the first 5-15 minutes of hydrolysis. These results suggest that there is a portion of highly accessible, very rapidly hydrolyzed amorphous cellulose present in dissolving pulp fibres. However, it must be noted that the total amount 051015202501020304050607080901000 50 100 150 200 250 300 350 400% Glucan ConversionRelative amount of CBM adsorbed (%)Time (minutes)CBM2aCBM44% Conversion127 of this highly accessible amorphous cellulose is very small. The ~50% reduction in accessible amorphous cellulose occurs in the first 15 minutes of hydrolysis, which corresponds to a conversion level of only 2.0% (±0.3%) (Figure 21). The continued relatively rapid hydrolysis rate after this drop in the amount of accessible amorphous cellulose implies that this decrease does not directly cause a slowdown in enzymatic hydrolysis rate, as is observed at higher conversion levels. These results were then compared to changes in crystallinity of the dissolving pulp over the course of this 6-hour hydrolysis. X-ray diffraction measurements revealed a slight but rapid increase in the overall crystallinity of the cellulose, from 66% to 69%, within the first 5 minutes of hydrolysis (Figure 22), consistent with the rapid increase in crystallinity observed by Sinitsyn et al. (1989). This increase in crystallinity is likely due to the rapid hydrolysis of an amorphous fraction of the cellulose, resulting in the residual cellulose having a higher overall crystallinity index (CrI). Figure 22: Crystallinity index as determined by XRD over the course of enzymatic hydrolysis. This work also suggests that the 2-dimensional surface phenomenon observed by CBM adsorption could also be detected by measuring the bulk (3-dimensional) properties of the cellulose. The ability of the CBM adsorption technique to accurately quantify changes in the structure and surface morphology directly at the cellulose surface is evident. Specifically, while the XRD measurements of the 656667686970710 100 200 300 400CrI(%)Hydrolysis time (minutes)128 bulk properties of the cellulose showed an overall change in crystallinity of only ~4%, the ability of the CBM adsorption technique to specifically quantify effects occurring at the cellulose surface, rather than a bulk property, resulted in a much more sensitive analysis of this surface phenomenon, revealing a 50% drop in the amount of adsorbed CBM44. This highlights the much greater sensitivity of the CBM adsorption technique for quantifying changes in cellulose properties at the liquid/solid interface, with significant implications for future studies aimed at gaining insights into relationships between cellulose surface properties and enzymatic or chemical treatments. Overall, these results demonstrate that a portion of highly accessible, very easily hydrolyzed cellulose is solubilized within the first 5-15 minutes of hydrolysis. This suggests that if the cellulose can be made highly accessible/amorphous, it can be very rapidly hydrolyzed. More broadly, the experiments discussed so far in Section 3.2 have demonstrated that the CBM adsorption technique can be used to quantify a number of different processes, including acid-induced amorphogenesis and the surface morphology of cellulose after treatment with hydrolytic cellulases. The ability of this CBM adsorption technique to quantify changes in cellulose accessibility and surface morphology after treatment with the amorphogenesis-inducing protein, Swollenin, was next assessed. 3.2.5 Quantification of Swollenin-induced changes in cellulose accessibility As Swollenin had previously been shown to disrupt mercerized cotton fibres (Jäger et al., 2011; Saloheimo et al., 2002), we next tried to apply CBM adsorption to quantify any changes in cellulose accessibility and surface morphology of mercerized cotton fibres treated with Swollenin. Although mercerization is known to cause a significant reduction in the crystallinity of cellulosic substrates, mercerized cellulose has been shown to retain some adsorptive capacity for crystalline binding CBMs 129 (Široký et al., 2012). After incubation with Swollenin, binding of both the crystalline and amorphous-binding CBMs to the mercerized cotton fibres increased (Figure 23). Figure 23: Adsorption of CBMs to Swollenin-treated cotton fibres. Swollenin promoted an increase in binding of CBM44 and, particularly, CBM2a. A BSA negative protein control was found to have no significant effect on the extent of binding of either CBM. At least three replicates were performed for each sample and error bars represent one standard deviation from the mean. Subsequent SEM micrographs of Swollenin-treated mercerized cotton fibres indicated that Swollenin treatments resulted in a smoothing of the roughened patches produced during mercerization (Figure 24). This smoothing effect was in contrast to the buffer or BSA-treated mercerized cotton fibres, which retained their roughened surface. After Swollenin treatment, the roughened patches on the surface of the fibres appeared to have been sloughed off, revealing the smooth, well ordered surface of the underlying cotton fibre. An increase in the turbidity in the supernatant after Swollenin treatment was also indicative of the release of small particles into solution. It is possible that the roughened patches at the surface of the mercerized cotton fibres contain a higher proportion of amorphous cellulose than the underlying fibre, as these protruding rough regions were more exposed to the NaOH used for mercerization. This treatment has been shown to promote the conversion of crystalline cellulose I into amorphous cellulose and crystalline cellulose II (Borysiak and 0510152025CBM2a CBM44 Sum CBMsmg CBM Bound per g CottonControlSwollenin130 Garbarczyk, 2003). The release of these roughened particles from the surface of the fibre could result in an increase in both the amount of exposed amorphous cellulose (primarily on the released particles) and the amount of exposed crystalline cellulose (primarily on the newly exposed surface of the underlying cotton fibre). However, it should be noted that the small roughened particles that are released from the surface of the cotton fibres appear to be approximately 100 nm in the shorter direction, and up to 1000 nm in the longer direction (estimated from Figure 24). Since the cellulosic cores of cotton microfibrils have diameters of only 3–5 nm and lengths of 100’s to 1000’s of nm (Sjöström, 1993; Somerville et al., 2004), it is possible that the small, roughened particles released from the surface of the cotton fibres still contain a significant amount of crystalline cellulose. Figure 24: Effect of Swollenin on mercerized cotton fibres imaged by SEM. The surface of the control fibre (Left) appears roughened due to the mercerization treatment. The rough features on the surface of the mercerized cotton fibres appear to have been sloughed off by the action of Swollenin (Right). Images are of representative fibres for the indicated treatment. Although it was not evident by which specific mechanism Swollenin promoted this “smoothing” effect, it is possible that Swollenin acts in a similar manner to the Expansin family of proteins, which have been shown to weaken plant cell walls through disruption of the hydrogen bonding network 131 between plant cell wall polymers (Cosgrove, 2000; McQueen-Mason and Cosgrove, 1994). If Swollenin disrupts hydrogen bonding via a similar mechanism to Expansins this might also explain how Swollenin appears to both disrupt the cell wall structure of Whatman filter paper No. 1 fibres and result in the swelling of cotton fibres (Jäger et al., 2011; Saloheimo et al., 2002). Taken together, the CBM adsorption profiles and SEM images suggest that Swollenin is targeting the more amorphous regions of the biomass. Specifically, Swollenin appears to loosen and release the roughened particles of cellulose from the surface of the fibre, while having little effect on the underlying well-organized smooth fibre surface. This process is illustrated in the cartoon in Figure 25.. These results indicated that substructure-specific CBMs could be used to track changes in cellulose accessibility after acid treatment and also be used to track changes in surface morphology during treatment with both hydrolytic cellulases and amorphogenesis-inducing proteins. This technique has several advantages over current alternatives as it provided a direct, quantitative method able to consolidate changes in multiple substrate characteristics. Specifically, changes in the amounts of accessible amorphous cellulose, accessible crystalline cellulose and the total (amorphous and crystalline) accessible cellulose can all be quantified. It was also apparent that this method could help better indicate the mode of action of non-hydrolytic, disruptive/amorphogenesis-inducing proteins and has potential to yield novel insights into the mechanisms of glycosyl hydrolases and other accessory enzymes involved in lignocellulose deconstruction. Additionally, this technique has the potential to facilitate comparisons of the disruptive capabilities of various non-hydrolytic proteins which might promote an increase in cellulose accessibility to the more traditional, hydrolytic components of the cellulase enzyme mixture. 132 Figure 25: Representation of quantifying specific manifestations of amorphogenesis using CBM adsorption. In the example illustrated here, the untreated cellulose (top) binds approximately equal amounts of CBM2a and CBM44. After protein-induced amorphogenesis, the possible manifestations of amorphogenesis can be separated between sloughing/splitting/delaminating/peeling effects (Left) and surface decrystallization effects (Right). If the ratio of bound CBM2a:CBM44 increases or remains unchanged during amorphogenesis, this suggests the splitting/delaminating/peeling manifestation, whereas if this ratio decreases, a manifestation of amorphogenesis involving the direct surface decrystalization is occurring. In the case of Swollenin, this ratio increased, consistent with the scenario on the left representing a sloughing/splitting/delaminating/peeling effect. Cr = Crystalline-binding, Am = Amorphous-binding. It is also likely that CBMs with specificities for certain hemicelluloses, such as xylan or mannan, could be used to track changes in the accessibility of these polymers during pretreatment, amorphogenesis and hydrolysis of softwoods, hardwoods and agricultural residues. In previous work, Filonova et al. (2007a) demonstrated the use of fluorescently-tagged mannan-specific CBMs to quantify the accessibility of mannan in wood tissues and pulp, after applying a protein-based lignin-blocking technique to prevent non-specific adsorption of the CBMs to lignin (Yang and Wyman, 2006). The use of CBM-specific antibodies, or conjugation of CBMs to distinct fluorophores, has been used to provide direct visualization of the locations of the different polymers or substructures at the substrate surface (Araki et al., 2010; Filonova et al., 2007a; Kawakubo et al., 2010). By utilizing a suite of different CBMs 133 with specificities for a range of structural features of the substrate, it will be possible to track changes in the surface morphology of biomass fibres during pretreatment, amorphogenesis, and hydrolysis in unprecedented detail. Regarding Swollenin in particular, the results suggested that Swollenin is promoting targeted disruption of the more amorphous regions of the cellulosic biomass. Thus the ability of Swollenin to target amorphous regions within cellulosic biomass was further investigated in the latter sections of the thesis. 3.2.6 Conclusions Previous attempts to try and quantify the amorphogenesis process have made use of a suite of complementary qualitative and semi-quantitative techniques. While these techniques have provided some useful information regarding the effects of amorphogenesis-inducing proteins on (ligno)cellulosic substrates, they have typically provided little insight into the mode of action of these proteins and have provided very little quantitative data. A novel technique, using substructure-specific CBMs was developed and used to quantify a variety of different phenomena, including changes in the accessibility and surface morphology of cellulose during acid-induced cellulose disruption, enzymatic hydrolysis, and Swollenin-induced amorphogenesis. This approach yielded new insights into the effects of Swollenin, suggesting that, rather than it promoting the disruption of crystalline cellulose faces, instead it targets the more amorphous regions of the biomass. Understanding how Swollenin affects the less-ordered regions of cellulosic and lignocellulosic substrates was the focus of the next part of the thesis. 3.3 Amorphogenesis and fibre fragmentation 3.3.1 Background The Swollenin-induced disruption of a model cellulosic substrate suggested that this protein may also be capable of promoting amorphogenesis on more industrially-relevant pretreated 134 lignocellulosic substrates. Previous work on steam pretreated poplar has shown that Beta-expansins, to which Swollenin is closely related, enhanced the efficiency of enzymatic hydrolysis of this lignocellulosic substrate (Baker et al., 2000). This work was of particularly interest as the vast majority of previous research that has demonstrated the effects of non-hydrolytic non-oxidative proteins have been performed on model cellulosic substrates. The ability of a Beta-expansin to enhance the enzymatic hydrolysis efficiency of steam-pretreated poplar represents the only other example to date of a non-hydrolytic non-oxidative protein promoting an enhancement of the enzymatic hydrolysis of a pretreated lignocellulosic substrate at commercially relevant conversion levels (Loosenin has also been shown to enhance the conversion of Agave tequilana bagasse, but at conversion levels of just ~7% (Quiroz-Castañeda et al., 2011)). The Beta-expansin protein family has been shown to promote the weakening/disruption of hydrogen bonding within plant biomass without releasing significant amounts of reducing sugars (Cosgrove, 2000; Tabuchi et al., 2011; Yennawar et al., 2006). Thus, it seems likely that this protein promotes amorphogenesis on lignocellulosic substrates and that this amorphogenesis enhances poplar hydrolysis. However, the previous Beta-expansin work on pretreated poplar focussed on the downstream effects of the Expansin (i.e. enhancement of hydrolysis), rather than assessing the direct disruptive effects of these proteins. In the work reported here the Swollenin was applied to steam-pretreated poplar to assess any putative macroscopic disruptive effects it might have on this industrially-relevant lignocellulosic substrate. Poplar was chosen specifically because Swollenin exhibits significant sequence similarity with the Beta-expansins, which have been shown to improve hydrolysis yields on pretreated poplar (Baker et al., 2000). In addition to steam-pretreated poplar, organosolv-pretreated lodgepole pine was also chosen as an additional woody substrate due to the abundance of softwood residues available in British 135 Columbia. Steam and organosolv pretreatments were selected because steam-pretreatment has been shown to be effective against hardwoods, while organosolv pretreatment is particularly effective on softwoods (Kurabi et al., 2005; Pan et al., 2005). As described earlier, woody lignocellulosic fibres from hardwoods and softwoods contain dislocations along the fibre length. Although these dislocations have been investigated in detail (Hidayat et al., 2012), the exact makeup of the biomass within these regions is still unclear. While endoglucanases have been shown to preferentially bind to fibre dislocations (Thygesen et al., 2011) and Type B CBMs (which both typically recognize amorphous cellulose) (Ding et al., 2006; Filonova et al., 2007b; Kawakubo et al., 2010), they have also been shown to bind certain Type A CBMs (which preferentially bind crystalline cellulose) (Filonova et al., 2007a; Hildén et al., 2003; Thygesen et al., 2011), and to exhibit birefringence (an indicator of crystalline structure) (Thygesen et al., 2011), suggesting the presence of at least some ordered/crystalline cellulose within the dislocations (Reviewed in (Hidayat et al., 2012). While the nature of the cellulose within these dislocations is still not fully understood, it is generally accepted that these regions exhibit less order than the surrounding fibre sections. The CBM adsorption profiles and SEM micrographs of Swollenin-treated cotton observed in Section 3.2 of this work suggested that Swollenin was targeting the less-ordered regions of the cellulose. Additionally, the fact that both amorphogenesis and fibre fragmentation are thought to occur at the very early stages of enzymatic attack, prior to the significant release of sugars, also implicates amorphogenesis-inducing proteins in fibre fragmentation. For these reasons, the ability of Swollenin to disrupt and fragment lignocellulosic and cellulosic fibres at fibre dislocations was next assessed. Since any disruption at fibre dislocations would appear at the ‘whole-fibre’ scale, this putative disruptive effect can be classified here as a macroscopic disruption. In addition to testing the ability of Swollenin to 136 promote fragmentation at fibre dislocations, fibre width measurements were also carried out to determine if Swollenin was capable of promoting fibre swelling on lignocellulosic substrates. Although Swollenin is so named because of its ability to swell cellulosic fibres, the only instance of fibre swelling induced by Swollenin has been demonstrated on mercerized cotton fibres. These predominantly cellulosic fibres are markedly different from the lignocellulosic substrates that were used in the work reported here. In addition to lacking lignin, cotton-derived cellulosic fibres are also enriched in pectins, which are not found at high levels in potential biorefinery feedstocks such as woody biomass or most agricultural residues (Sjöström, 1993; Vaughn and Turley, 1999). It is possible that the previously documented swelling of cotton fibres induced by Swollenin is only possible due to the lack of lignin holding together adjacent cellulose microfibrils. Prior to assessing this putative Swollenin-induced fibre fragmentation, the changes in macroscopic fibre properties over the course of enzymatic deconstruction were first quantified, so that any effects of Swollenin could be put into perspective relative to the effects of the whole enzyme mixture. Specifically, fibre fragmentation and fibre swelling over a range of pretreated lignocellulosic substrates were assessed during hydrolysis. This was done, in part, to assess what macroscopic effects Swollenin might be involved in during enzymatic deconstruction (i.e. fragmentation and swelling), but also because the macroscopic mode of action of enzymatic hydrolysis on pretreated lignocellulosic biomass is still not yet fully understood. There is still considerable debate regarding the actual mechanisms by which a “cellulase mixture” deconstructs the cellulosic component of pretreated biomass substrates, particularly at the macroscopic fibre level of plant cellular structure (Chandra et al., 2007). Some workers have suggested that the initial mode of enzymatic attack takes place on the outer layer of the cellulose surface where the constituent fibres are peeled along their length, layer-by-layer, in an “onion peeling” fashion (Gama and Mota, 1997; Lee et al., 1983; Penttilä et al., 2010; Rousselle et 137 al., 2002; Wang et al., 2006). This peeling mechanism has also been termed a “shaving” or “planing” mode of action. Alternatively it has been suggested that cellulose deconstruction is a two-step process where the cellulose rich fibres are initially fragmented or disaggregated into shorter fibres, resulting in a greater overall surface area for the enzymes to subsequently attack (Clarke et al., 2011; Lee and Fan, 1980; Park et al., 2007; Saqib and Whitney, 2006). This “cutting” mode of deconstruction has also been termed a “fragmentation” mode of action. Studies using model cellulosic substrates (Gama et al., 1997; Peters et al., 1991) have suggested that the dominant mode of deconstruction was strongly influenced by the fibre dimensions of the initial substrate, with the larger particles first fragmented/cut into smaller particles (“cutting”) while the smaller particles seemed to be hydrolyzed by a “peeling/erosion” type of mechanism. By assessing the macroscopic changes in fibre properties over the course of hydrolysis, we hoped to shed some light on the mechanisms underlying enzymatic hydrolysis of whole lignocellulosic fibres. After quantifying the macroscopic effects occurring during the enzymatic deconstruction of several lignocellulosic substrates, the ability of Swollenin to swell and fragment these substrates was assessed. In addition to studying lignocellulosic substrates after Swollenin treatment, dissolving pulp was also included as a pure cellulose control, as it consists of long cellulose fibres derived from lignocellulosic biomass and it also contains dislocations that should be ideal for revealing fibre fragmentation activity. Additionally, this model cellulosic substrate was included in order to reveal any potential effects that Swollenin may promote on a cellulosic substrate which might be masked on lignocellulosic substrates. For example, it is possible that Swollenin may be capable of disrupting and swelling pure cellulosic substrates, but the lignin and/or hemicellulose found within lignocellulosic substrates may limit Swollenin-induced amorphogenesis, thus masking any effects promoted by Swollenin on the cellulosic fraction of biomass. We therefor assessed both the macroscopic changes in fibre properties for a model as well as pretreated lignocellulosic substrates. 138 Finally, in order to try to shed some light on the nature of the cellulose present within fibre dislocations, the CBM probe technique described in Section 3.2 was modified such that each CBM was conjugated to a different fluorescent tag. These fluorescent CBMs were then imaged using confocal microscopy to determine their binding profile with respect to the dislocations within the dissolving pulp fibres. It should be noted that defining whether a CBM binds crystalline cellulose or amorphous cellulose is complicated by the fact that the binding of all cellulose-binding CBMs fall somewhere on a continuous scale between highly preferential for crystalline cellulose and highly preferential for amorphous cellulose (Fox et al., 2013; McLean et al., 2002). Additionally, the crystallinities of different cellulosic substrates are surprisingly difficult to measure accurately and reproducibly (Park et al., 2010). These problems are compounded by issues with defining what constitutes crystalline cellulose. For example, if three cellulose chains are associated lengthwise, does this constitute a crystalline region? In Section 3.2 it was shown that conversion of relatively crystalline Avicel to highly amorphous PASC resulted in a dramatic increase in the adsorption of the supposedly crystalline-binding CBM, CBM2a. This likely arose from the greatly enhanced accessibility of the bulk cellulose, but also possibly from the exposure or creation of many small microcrystalline regions (of the Cellulose II form) after precipitation of the solubilized cellulose during the formation of the PASC (e.g. 3-4 cellulose chains associated together). The presence of these microcrystalline regions could explain the observation by Thygesen et al. (2011) that dislocations in filter paper fibres are targeted by amorphous-binding endoglucanase, but also appear to still contain crystalline cellulose. For the purposes of this work, crystalline cellulose is defined as cellulose preferentially recognized by CfCBM2a, and amorphous cellulose as that preferentially recognized by CtCBM44. The potential of Swollenin was next investigated to see if it would provide any Insights into its ability to promote macroscopic disruption of cellulosic and lignocellulosic substrates. A new technique 139 for imaging cellulose surface morphology involving the adsorption of fluorescently-tagged CBMs was also used to directly visualize potential targets of Swollenin action on cellulosic fibres. 3.3.2 Quantifying macroscopic fibre properties during enzymatic deconstruction Prior to assessing the effects of Swollenin on the macroscopic properties of lignocellulosic fibres, changes in fibre properties were first assessed during the course of enzymatic hydrolysis. Steam pretreated poplar was prepared at two different pretreatment temperatures (180°C and 200°C) in order to provide similar substrates with different fibre lengths. Changes in pretreatment severity are known to influence fibre length (Pan et al., 2007). In addition to the steam-pretreated poplar substrates, organosolv pretreated lodgepole pine was used as a representative softwood substrate. These substrates were hydrolyzed for 72 hours with 15mg enzyme/g glucan, as this was close to the minimum enzyme loading that could achieve 70% glucan conversion within 48 hours (an industrially relevant conversion scenario) (Figure 26) (Arantes and Saddler, 2011). Fibre lengths and widths were monitored over the course of hydrolysis. Enzymatic hydrolysis of these substrates revealed that initial fibre length did not appear to influence the enzymatic hydrolyzability of the biomass (as was also observed in Figure 6 in Section 3.1.3 of this thesis) (Figure 26). 3.3.2.1 Fibre length The analysis of fibre lengths over the course of hydrolysis showed that for all three substrates, the fibre length decreased rapidly (within the first 6 hours), then levelled out (Figure 27). Based on previous work, it is likely that the fragmentation occurring at these early hydrolysis timepoints is occurring at the dislocations found along the lengths of the fibres (Reviewed in (Hidayat et al., 2012). This rapid reduction in fibre length occurs prior to the slowdown in enzymatic hydrolysis that occurs toward the end of hydrolysis, usually when conversion levels reach ~70-80% for the substrates tested (Arantes and Saddler, 2011). This implies that, although fibre fragmentation is one of the early 140 macroscopic changes occurring during enzymatic hydrolysis, continuous fragmentation of the fibres is not required to maintain efficient deconstruction throughout the course of hydrolysis. This in turn suggests that, after initial fragmentation, enzymatic hydrolysis occurs primarily in the fibre width dimension. Figure 26: Enzymatic hydrolysis profiles of pretreated poplar and lodgepole pine. No correlation was observed between initial fibre length and enzymatic hydrolyzability. SPP180: Steam Pretreated Poplar pretreated at 180°C, SPP200: Steam Pretreated Poplar pretreated at 200°C, OPLP: Organosolv Pretreated Lodepole Pine. Figure 27: Average fibre lengths of fibres from two pretreated hardwood and one pretreated softwood substrates over the course of enzymatic hydrolysis. SPP180: Steam Pretreated Poplar pretreated at 180°C, SPP200: Steam Pretreated Poplar pretreated at 200°C, OPLP: Organosolv Pretreated Lodepole Pine. 02550751000 12 24 36 48 60 72Glucan Conversion (%)Hydrolysis Time (h)SPP180SPP200OPLP0.00.20.40.60.81.01.21.41.61.82.00 12 24 36 48 60 72Fibre Length (mm)Hydrolysis Time (h)SPP180SPP200Fibre Lengths: OPLP >> SPP180 > SPP200 141 3.3.2.2 Fibre width Changes in fibre width over the course of hydrolysis for the three pretreated lignocellulosic substrates were measured using the fibre quality analyser (FQA). Although no clear trends in the average width of the fibres could be seen during the course of hydrolysis, it was apparent that enzymatic hydrolysis promoted a dramatic increase in the variability of the fibre width (represented as standard deviations from the average fibre width) (Figure 28). This suggested that, although the average fibre width itself did not change significantly during hydrolysis, the number of fibres with widths much larger than average, as well as the number of fibres with widths much smaller than average, increased dramatically during hydrolysis. The increase in the standard deviation associated with fibre width appeared to become more pronounced after the first 6 hours of hydrolysis, around the same time that the initial fibre fragmentation process was tapering off. Together, these results suggest that enzymatic deconstruction of the lignocellulosic substrates tested here occurs through an initial reduction in fibre length, followed by enzymatic activity promoting both fibre swelling and reductions in fibre width. These two apparently contradictory effects can be readily explained if the enzymatic deconstruction proceeds through a mechanism involving the initial swelling/peeling of the surface of the fibres, followed by rapid hydrolysis of this disrupted/swollen fibre surface. This mode of enzymatic hydrolysis would result in both an increase in fibre width, as the fibres are initially swollen and peeled, followed by a decrease in width as these swollen/disrupted regions are rapidly hydrolyzed. These phenomena would likely be occurring simultaneously during the enzymatic deconstruction process, thereby producing the dramatic increase in standard deviations observed after the first 6 hours of hydrolysis. Interestingly, it appears as though the swelling mechanism precedes the peeling/shaving mechanism, as evidenced by the initial increase in average fibre width over the first 1-6 hours of hydrolysis (Figure 28). After this initial increase, the average fibre width decreases back to levels similar 142 to the widths observed prior to hydrolysis, suggesting that the peeling/shaving mechanism ‘catches up’ to the initial swelling effect. Figure 28: Average fibre width (midpoint of each bar) and standard deviation of the width (height of each bar) for SPP180, SPP200 and OPLP over the course of hydrolysis. SPP180: Steam Pretreated Poplar pretreated at 180°C, SPP200: Steam Pretreated Poplar pretreated at 200°C, OPLP: Organosolv Pretreated Lodepole Pine. 1520253035400 12 24 36 48 60 72 84Hydrolysis time (h)Fibre width (µm)SPP18020222426283032340 12 24 36 48 60 72 84SPP 200Hydrolysis time (h)Fibre Width (µm)1520253035400 12 24 36 48 60 72 84Fibre width (µm)Hydrolysis time (h)OPLP143 In order to look for visual confirmation of this swelling/peeling process, SEM microscopy was performed on the steam pretreated poplar (SPP200) samples before and after enzymatic hydrolysis. Prior to hydrolysis, when the standard deviations on the average fibre widths are small, the surface of the fibres appear smooth and well ordered (Figure 29, A,B). However, after partial hydrolysis, when the standard deviations on the fibre widths are much larger (Figure 28), a dramatic roughening and peeling of the fibre surfaces could be observed (Figure 29, C,D). The features observed on the fibre surfaces match what would be expected if enzymatic hydrolysis was occurring via a peeling/swelling of the surface of the fibres followed by rapid hydrolysis of the peeled/swollen regions. Figure 29: Representative SEM micrographs of SPP200 samples treated with either buffer (A,B) or cellulase (C,D). Fibre peeling can be observed at both low (C) and high (D) magnifications. Cellulase Control A B C D 144 3.3.3 Quantifying Swollenin-induced macroscopic manifestations of amorphogenesis on lignocellulosic biomass After revealing the macroscopic processes at play during the enzymatic deconstruction of lignocellulosic biomass, Swollenin was assessed for its potential involvement in these processes. Specifically, the putative ability of Swollenin to promote fibre fragmentation and swelling was evaluated. Initially, Swollenin was tested for its ability to fragment and swell the poplar and lodgepole pine substrates as well as the dissolving pulp which was used as a relatively pure cellulosic control. As described earlier, it was thought that relatively amorphous regions at fibre dislocations could serve as targets for the disruptive abilities of Swollenin. However, at least for the pretreated lignocellulosic samples tested here, Swollenin was not able to promote fibre swelling (Figure 30), nor was it capable of inducing fibre fragmentation (Figure 31), implying that complete disruption of fibre dislocations was not occurring. Figure 30: Average fibre width of pretreated hardwood, softwood and dissolving pulp substrates after treatment with either a BSA control or Swollenin at 10 µg protein/mg substrate. All samples were run in triplicate and error bars represent one standard deviation from the mean. In order to determine if the inability of Swollenin to fragment these lignocellulosic fibres might be due to the presence of non-cellulosic polymers (i.e. hemicellulose and/or lignin), a hardwood 0510152025303540OPLP SPP180 SPP200 DSPAverage Fibre Width (µm)BSASwollenin145 dissolving pulp, representative of almost pure cellulose, was used. Application of Swollenin to this model cellulosic substrate showed that Swollenin was in fact capable of promoting fibre fragmentation, albeit only in the absence of the bulk of the lignin and hemicellulose (dissolving pulp contains ~4% hemicellulose and 0.25 to 0.5% lignin) (Figure 31). This suggested that the hemicellulosic and/or lignaceous portion of the biomass was more recalcitrant to Swollenin-induced disruption. Despite the ability of Swollenin to disrupt the cellulosic portion of the biomass within the dislocations, the hemicellulosic and/or lignaceous compounds present within lignocellulosic biomass restricted the complete Swollenin-induced separation of fibres at their dislocations. Figure 31: Average fibre length of various pretreated lignocellulosic pulps and dissolving pulp after treatment with either buffer or Swollenin at 10µg protein/mg substrate. DSP: Dissolving Pulp, SPCS: Steam Pretreated Corn Stover, OPCS: Organosolv Pretreated Corn Stover, SPP: Steam Pretreated Poplar, OPP: Organosolv Pretreated Poplar, SPLP: Steam Pretreated Lodgepole Pine, OPLP: Organosolv Pretreated Lodgepole Pine. All samples were run in triplicate and error bars represent one standard deviation from the mean. 3.3.4 Quantifying Swollenin-induced macroscopic manifestations of amorphogenesis on a model cellulosic substrate Since Swollenin alone was able to promote fragmentation of dissolving pulp, this ability was compared to the fragmentation ability of a number of purified hydrolytic enzymes and a negative 0.00.10.20.30.40.50.60.70.8OPLP SPP180 SPP200 DSPAverage Fibre Length (mm)BSASwollenin146 protein control, BSA (Figure 32). The purpose of this work was to determine if Swollenin was capable of acting synergistically with hydrolytic enzymes in promoting fibre fragmentation. Treatment of dissolving pulp fibres with either buffer alone, BSA, a family 10 xylanase (Xyn10) or Cel7A (a cellobiohydrolase) did not promote any fragmentation of the fibres. However, both an endoglucanase (Cel5A) and Swollenin reduced the average fibre length of the dissolving pulp fibres. The observation here that endoglucanases are the most adept at promoting fibre fragmentation is consistent with previous work demonstrating the efficient fragmentation of fibres by endoglucanases (Lecourt et al., 2010). Figure 32: Average fibre length of dissolving pulp fibres treated overnight at 50°C with 10µg purified enzyme per mg substrate. BSA: Bovine Serum Albumin, Xyn: Xylanase GH10, CBH: Cel7A, EG: Cel5A, Swo: Swollenin. Interestingly, although Swollenin was not capable of promoting fragmentation to the same degree as endoglucanases, it was able to promote significant fragmentation, especially when compared to Cel7A. This demonstrates that, despite the highly active nature of Cel7A for depolymerizing cellulose chains, this cellulose depolymerizing activity did not result in fibre fragmentation. Contrarily, Swollenin does not cleave cellulose and yet is capable of promoting fibre fragmentation. Because Swollenin 0.00.10.20.30.40.50.6Average Fibre Length (mm)147 exhibits macroscopic disruptive effects more similar to those of endoglucanases (which are thought to act on amorphous cellulose) than to cellobiohydrolases (which predominantly attack more crystalline regions of the cellulose), this again suggested that Swollenin targets the more amorphous regions of the cellulosic biomass. Swollenin did not exhibit obvious synergism when combined with Cel7A or Cel5A when looking at dissolving pulp fragmentation (Figure 32), however it is unclear if varying certain reaction conditions, such as enzyme loading, might lead to synergistic interactions. Overall, these fragmentation results confirmed Swollenin’s complementary role in achieving effective enzymatic hydrolysis. It may be of even greater importance in high-consistency hydrolysis, where fibre fragmentation and hydrolysis slurry liquefaction are important preliminary steps in efficient high-solids hydrolysis (Kristensen et al., 2009). The Swollenin-induced fragmentation of dissolving pulp prompted further work into this phenomenon. The enzymatic hydrolysis and fragmentation curves for the dissolving pulp indicated that, during enzymatic deconstruction, the dissolving pulp was fragmented even more rapidly than the pretreated lignocellulosic substrates (Figure 33). For example, while it took up to three hours for the average fibre length of the lignocellulosic substrates to be reduced by 50%, this was achieved in just five minutes when hydrolyzing dissolving pulp. This extremely rapid fragmentation relative to the lignocellulosic substrates suggests that, at similarly low conversion levels, complete enzymatic deconstruction of the dislocations within the fibres is achieved much more easily in the absence of hemicellulose and lignin. This supports the data suggesting that Swollenin is capable of promoting fragmentation of dissolving pulp, but incapable of fragmenting lignocellulosic substrates. 148 Figure 33: Sugar release and average fibre length during enzymatic hydrolysis of dissolving pulp using 15mg enzyme/g glucan. All reactions were performed in triplicate and error bars represent one standard deviation from the mean. The rapid fragmentation of the dissolving pulp was also indicated by the increase in the ‘settlability’ of the pulp. Here the term settlability is used to describe the ability of the pulp to settle under gravity. For example, in Figure 34, dissolving pulp was hydrolyzed for varying lengths of time then equal masses were weighed out, stained with direct orange dye, washed, and then allowed to settle overnight in buffer. Comparing these figures, it can be seen that a reduction in fibre length corresponds directly to an increase in the settlability of the pulp. Interestingly, in both Figure 33 and Figure 34, a distinct ‘two-step’ profile can be seen for the reduction in fibre length and the increase in settlability. Specifically, an initial very rapid reduction in fibre length and increase in settlability occurs within the first five minutes of hydrolysis. This initial rapid change is followed by a gradual reduction over the course of the next hour. However, between 60 and 90 minutes there appears to be a second dramatic reduction in fibre length and increase in settlability of the fibres. Together, these results suggests that there may be two distinct categories of fibre dislocations, including major dislocations which are hydrolyzed and fragmented within the first 5 minutes of hydrolysis, and minor dislocations, which are 00.10.20.30.40.50.60246810120 60 120 180 240 300 360 420Average Fibre Length (mm)mg/mL GlucoseHydrolysis Time (minutes)Sugar ReleaseFiber Length149 more recalcitrant to fragmentation and, under the reaction conditions employed here, took 60-90 minutes of cellulase treatment before fragmentation occurred. Figure 34: ‘Settlability’ of dissolving pulp fibres over the course of hydrolysis at 15mg enzyme/g glucan. A dramatic increase in settlability is observed within the first 5 minutes, followed by a subsequent step-up in settlability between 60 and 90 minutes. Fibres were stained with Direct Orange dye and washed thoroughly prior to settling overnight in order to enhance the visibility of the fibres. The nature of the cellulose within fibre dislocations, as well as the putative presence of distinct populations of major and minor fibre dislocations was next investigated. In order to gain insights into the properties of fibre dislocations, the CBM adsorption technique developed in Section 3.2 of this thesis was modified to allow for direct visualization of cellulose surface morphology. This was carried out by conjugating distinct fluorescent tags to both CBM2a and CBM44, which preferentially bind to crystalline and amorphous cellulose, respectively. Specifically, the amine-reactive fluorescent dyes 6-((7-Amino-4-Methylcoumarin-3-Acetyl)amino)Hexanoic Acid (AMCA-X) and Oregon Green-514 (OG) were conjugated to CBM2a and CBM44, respectively. These dyes emit in either the blue region (442nm for AMCA-X) or the green region (526 nm for Oregon Green-514), resulting in the emission of blue light from the CBM2a-AMCA-X conjugate and the emission of green light from the CBM44-OG conjugate. These conjugated CBMs should therefore, upon excitation of the fluorophores, result in the emission of blue light from the predominantly crystalline regions of the cellulose and the emission of green light from the 0 5 15 30 60 90 180 120 360 Hydrolysis Time (Minutes) 150 predominantly amorphous regions of the cellulose. After simultaneous adsorption of these two fluorescently tagged CBMs to dissolving pulp fibres, the fibres were thoroughly washed, then imaged using confocal microscopy (Figure 35). Figure 35: A representative dissolving pulp fibre imaged without excitation light (A), with excitation of CBM2a-AMCA-X (B), and with excitation of CBM44-OG (C). The colored pixels from B and C were extracted, combined, and overlaid onto the original fibre image to produce figure D. The colored pixels were then enhanced (E). In general, CBM2a (blue) localized to well-ordered regions of the fibre surface while CBM44 (green) was predominantly bound within fibre dislocations. A full description of the enhancement process is provided in the preceding paragraphs. 151 Figure 35A represents a dissolving pulp fibre in the absence of excitation light, while Figure 35B and Figure 35C represent the images obtained after excitation of either CBM2a-AMCA-X (blue) or CBM44-OG (green), respectively. The colored pixels in each of these images were then extracted and overlaid on the original fibre image to produce the composite image in Figure 35D. In order to better visualize the binding patterns on the surface of the dissolving pulp fibres, a minimum color intensity threshold was selected for the blue and green colors to eliminate low-intensity (faint) regions of each color. The remaining bright pixels were then color-saturated and overlaid back onto the fibre image to produce the color-enhanced image shown in Figure 35E. From this figure it can clearly be seen that the binding of CBM2a-AMCA-X predominantly binds to the relatively well-ordered regions of the fibre surface, while binding of CBM44-OG is enriched at the fibre dislocations. It is important to note here that the patterns observed for each CBM was dependent on the gain settings during image acquisition. For example, if gain settings were maximized, then both CBMs appeared to bind to the entire surface of the fibre. It was only by lowering the gain settings that changes in the distributions of the CBMs on the fibre surface became apparent. Thus it must be emphasised that both CBMs do bind to the entire surface of the fibre. However, the binding density (amount bound per unit area) of each CBM exhibited distinct patterns. In other words, both CBMs bound throughout the surface of the fibre, but CBM44 had a higher binding density at dislocations, while CBM2a had a higher binding density along the more organized and well-structured surfaces of the fibres. The CBM adsorption patterns relating to fibre dislocations are consistent with previous data, which indicate a general trend of CBM adsorption to the dislocation sites within the fibre. Specifically, previously-studied Type B CBMs, including members from families CBM4, CBM6 and CBM28, were all found to preferentially bind to fibre dislocations (Ding et al., 2006; Filonova et al., 2007a; Kawakubo et al., 2010). This correlates with the preferential adsorption of CBM44 to the dislocation sites observed here. Additionally, previous work on certain bacterial Type A CBMs showed that these CBMs do not 152 appear to preferentially bind to dislocations. Specifically, two family 3 CBMs from the cellulosome complexes of C. josui and C. thermocellum did not exhibit preferential binding to dislocations (Ding et al., 2006; Kawakubo et al., 2010). These Type A CBMs are similar in size to the CtCBM2a used in the current work and have recently been shown to exhibit a similar binding order parameters to CtCBM2a (Fox et al., 2013). The binding order parameter was recently suggested by Fox et al. (2013) as a metric for quantifying the preference of CBMs for crystalline or amorphous regions of cellulose, with a higher binding order representing a higher selectivity for crystalline regions. The results reported in our work are similar to those reported by Kawakubo et al. (2010), who found that a Type B CBM28 preferentially bound to dislocations, while the Type A CBM3 from C. josui did not. However, Fox et al. (2013) demonstrated that the related CBM3A from C. thermocellum CipA was the most promiscuous CBM studied, suggesting that CBM2a from C. thermocellum employed in the work presented here may be a better choice as a probe for crystalline cellulose than is the CBM3 from C. josui employed previously (Kawakubo et al., 2010). In addition to using CBMs as probes, an amorphous-binding endoglucanase has also been used to demonstrate the abundance of amorphous cellulose within dislocations (Thygesen et al., 2011). Overall, the results presented here regarding CBM adsorption to fibre dislocations are consistent with previous results and suggest that fibre dislocations are enriched in amorphous cellulose compared to the smoother fibre sections. The confocal images in Figure 35 appear to show one major dislocation (at the right of the fibre), and one minor dislocation (towards the left of the fibre). The dramatic differences in the severity of these two dislocations lends weight to the idea that the step-wise reduction in fibre length (Figures 33 and 34) is due to the presence of distinct populations of major, easily-hydrolyzable dislocations and minor, more recalcitrant dislocations. In other words, it is possible that the initial very rapid reduction in fibre length may be due to hydrolysis within major dislocations, while the second fibre length reduction step may be due to the somewhat slower hydrolysis of the more intact minor dislocations. 153 Our earlier work showed that Swollenin appears to target the more amorphous regions of cellulosic biomass (Section 3.2.5). Further evidence for this comes from the ability of Swollenin to promote fibre fragmentation in a manner similar to the amorphous-targeting endoglucanases, contrary to the crystalline-targeting cellobiohydrolases. Together with the CBM adsorption binding profiles on the dissolving pulp fibres, this suggests that Swollenin is promoting fibre fragmentation by targeting the amorphous cellulose found within fibre dislocations. The fibre length after Swollenin treatment did not decrease any further after an additional 3-day incubation. This suggested that Swollenin may only be capable of promoting fragmentation at major/severe dislocations which are susceptible to disruption/separation and that other minor dislocations are not susceptible to separation by Swollenin, regardless of the length of incubation. Previous Swollenin work has suggested that the mechanism of disruption involves the weakening of hydrogen bonding, particularly within amorphous regions of cellulose or even hemicellulose (Jäger et al., 2011; Saloheimo et al., 2002). It therefore appears likely that the mechanism by which Swollenin is capable of promoting fibre fragmentation involves the weakening of hydrogen bonds, thereby allowing slippage between adjacent chains within fibre dislocations. This weakening of hydrogen bonding and chain slippage is apparently sufficient to cause fragmentation within certain severe dislocations, but not within the less severe dislocations. When Swollenin was incubated with a range of pretreated lignocellulosic substrates, including steam pretreated poplar and organosolv pretreated lodgepole pine, no reduction in fibre length was observed (Figure 31). This suggested that other factors in the biomass prevent Swollenin-induced fibre fragmentation. Although Swollenin was not capable of promoting fibre fragmentation of the woody lignocellulosic substrates, it is still possible that Swollenin can play an active role in promoting the amorphogenesis of lignocellulosic substrates. Poplar was chosen as a substrate for Swollenin due to the 154 reported ability of Beta-expansins to enhance hydrolysis yields from this woody lignocellulosic substrate (Baker et al., 2000). However, previous work on expansins indicated that Beta-expansins appear to specialize in disrupting grass cell walls (Cosgrove, 2000). It is likely that Swollenin, which is more closely related to the Beta-expansins than the Alpha-expansins, may promote amorphogenesis more readily on grass cell walls than on those derived from woody material. Additionally, the effects assessed here were all macroscopic effects of disruption (fibre fragmentation and swelling) of woody biomass. As Swollenin might promote amorphogenesis at more microscopic/nanoscopic levels its effect on a pretreated lignocellulosic grassy substrate (in this case steam-pretreated corn stover), was next assessed. 3.3.5 Conclusions This part of the work showed that the enzymatic deconstruction of pretreated wood occurs via an initial rapid fibre fragmentation, followed by a subsequent swelling/peeling mode of enzymatic action reducing fibre width. Swollenin was assessed for its ability to influence these macroscopic fibre properties during enzymatic deconstruction. Although Swollenin did not promote fragmentation or swelling of the pretreated wood substrates, Swollenin-induced fragmentation of a pure cellulosic substrate, dissolving pulp, was achieved. This suggested that compounds within the biomass such as lignin and/or hemicellulose likely play a role in restricting Swollenin-induced fibre fragmentation. Fibre length reduction profiles over the course of hydrolysis suggested that dissolving pulp fibres may contain two distinct classes of dislocations. Confocal microscopy coupled with tagged CBMs was used to determine that these dislocations are enriched in amorphous cellulose. The ability of Swollenin to promote fibre fragmentation of a relatively small proportion of potential fragmentation sites suggests that Swollenin is only capable of promoting fibre fragmentation at major/severe dislocations, where there is an abundance of amorphous cellulose. It seems likely that Swollenin is capable of weakening the 155 hydrogen bonds within relatively amorphous regions of cellulose, which allows slippage between adjacent chains within the fibre dislocations, thereby promoting fibre fragmentation. 3.4 The effects of Swollenin on an industrially-relevant biomass substrate, steam pretreated corn stover 3.4.1 Background The accessibility of cellulose is known to play a major role in influencing the efficiency of enzymatic hydrolysis (Arantes and Saddler, 2011; Jeoh et al., 2007; Rollin et al., 2011; Wiman et al., 2012). The work presented earlier in this thesis, involving the adsorption of substructure-specific CBMs, showed that Swollenin can enhance cellulose accessibility by targeting the less-ordered regions of cellulosic biomass, rather than by directly disrupting the crystalline cellulose faces. However, this earlier work also indicated that the Swollenin-induced macroscopic amorphogenesis of woody biomass substrates is limited by the presence of hemicellulose and/or lignin. In the following section, the ability of Swollenin to promote macroscopic and microscopic amorphogenesis of a grassy lignocellulosic substrate was determined and the putative synergistic interactions between Swollenin and hydrolases during the breakdown of this grassy lignocellulosic material were also assessed. Although the work reported earlier showed that Swollenin did not promote macroscopic disruption of pretreated wood substrates it is still possible that Swollenin can actively disrupt lignocellulosic grassy substrates at the microscopic or nanoscopic levels. As described previously, Swollenin has greater sequence similarity to the Beta-Expansins, which have been shown to act predominantly on Type II plant cell walls (those of the grasses and other commelinoid monocots), than to the Alpha Expansins, which have been found to act predominantly on Type I plant cell walls (those of the dicots and non-commelinoid monocots) (Cosgrove, 2000). This suggested that Swollenin may play a more active role in disrupting grass cell walls than those of woody plants. This also pointed to the 156 possibility that proteins such as the Expansins and Swollenin may play specific roles only within particular types of lignocellulosic biomass. The apparent finely-tuned specificities of these amorphogenesis-inducing proteins imply that highly-efficient combinations of protein and biomass could have been previously overlooked in prior research. Thus, we next assessed the ability of Swollenin to promote amorphogenesis at more microscopic/nanoscopic levels on a pretreated lignocellulosic grassy substrate (in this case steam-pretreated corn stover, SPCS) which is readily available throughout much of North America (Graham et al., 2007; Kadam and McMillan, 2003). In order to better understand the effects of Swollenin on SPCS, a number of techniques were applied. Specifically, the supernatants of Swollenin-treated SPCS were assessed to determine if Swollenin was capable of solubilizing any portion of this biomass. Additionally, the effects of Swollenin on the accessibility and surface morphology of this substrate was determined using the CBM adsorption technique described in Section 3.2.5. Finally, Swollenin was tested for its ability to synergize with isolated cellulase and hemicellulase enzymes in an attempt to enhance the efficiency of hydrolysis of SPCS via enhanced amorphogenesis and improved cellulose accessibility. The work was primarily performed to determine if Swollenin was capable of disrupting a pretreated lignocellulosic grassy (Type II) biomass and to investigate what portion of the biomass is being targeted by Swollenin for disruption. 3.4.2 Macroscopic effects of Swollenin on steam pretreated corn stover Earlier, Swollenin was shown to promote fibre fragmentation of dissolving pulp by targeted disruption of the disordered cellulose within fibre dislocations. However, fragmentation of other woody biomass fibres appeared to be hindered by the presence of lignin and/or hemicellulose within these substrates. It was hypothesized that Swollenin may act in a more similar manner to the Beta-expansins, which preferentially disrupt Type II (grassy) cell walls. Steam pretreated corn stover was subsequently selected to assess the possible influence of Swollenin addition to a grassy biomass with good potential 157 as a biorefinery feedstock (Kadam et al., 2008). Swollenin was applied to both steam pretreated and organosolv pretreated corn stover (see Section 1.1.8 for a brief description of these processes), and the average fibre length of control and Swollenin-treated fibres were then determined (Figure 36). It was apparent that Swollenin did not promote a reduction in the fibre length of corn stover regardless of the type of pretreatment. It is possible that, as with the woody lignocellulosic substrates assessed in Section 3.3.3, Swollenin is incapable of promoting macroscopic fibre fragmentation in the presence of lignin/hemicellulose. An alternative explanation for this lack of fibre fragmentation could be that fibre dislocations are not present in corn stover. However, work by Samaniuk et al. (2011) revealed a dramatic reduction in fibre length upon enzymatic hydrolysis with mixing, implying that fibre dislocations do indeed exist within this substrate. However, further work into the presence of these features within corn stover fibres would be required before this can be stated conclusively. Although the addition of Swollenin did not promote the macroscopic amorphogenesis of corn stover fibres the potential microscopic/nanoscopic disruptive effects of this protein were next evaluated. Figure 36: Fibre length of steam-pretreated (SPCS) and organosolv pretreated (OPCS) corn stover after treatment with either a buffer control or Swollenin. 0.00.10.20.3SPCS OPCSAverage Fibre Length (mm)BufferSwollenin158 3.4.3 Microscopic effects of Swollenin on steam pretreated corn stover 3.4.3.1 Small particle release As Swollenin had previously been shown to slough-off roughened patches from the surface of model cellulosic fibres (see Section 3.2.5), it was thought likely that this effect might also be observed on corn stover. However, Swollenin was instead found to promote a reduction in the turbidity of SPCS supernatants, as indicated by the absorbance of the supernatant at 600nm, suggesting that Swollenin might aid in the solubilisation of very small particles present within the SPCS mixtures (Figure 37). The previously observed lack of hydrolytic activity of Swollenin on cellulose and very low activity on birchwood xylan (Saloheimo et al., 2002) indicated that this solubilization is likely occurring via the release of pre-existing oligomers within the biomass, rather than through the direct cleavage of biomass polymers. Since Swollenin appeared to be capable of solubilizing a portion of the SPCS biomass, the identity of solubilized oligomers and monomers present in the Swollenin-treated samples was next assessed. Figure 37: Turbidity of the supernatants of Swollenin and control-treated SPCS. Swollenin appears to decrease the turbidity of the supernatants. The slight increase in turbidity in the presence of the BSA protein control likely corresponds to protein aggregation, also observed in the sample containing just BSA and buffer in the absence of SPCS. BSA: Bovine Serum Albumin, SPCS: Steam Pretreated Corn Stover. 00.10.20.30.40.5Buffer +SPCSBSA +SPCSSwollenin+ SPCSBuffer +BSABuffer +SwolleninTurbidity (OD600)159 3.4.3.2 Quantification of oligomer release by Swollenin 3.4.3.2.1 High performance liquid chromatography analysis In order to determine if solubilization was occurring, the supernatants of Swollenin-treated SPCS samples were analyzed by high-performance liquid chromatography (HPLC) to assay for the release of monomeric or oligomeric sugars into the supernatant. It was apparent that Swollenin was capable of releasing significant amounts of oligomers, as well as low amounts of glucose and xylose, from the SPCS substrate (Figure 38). The oligomers were predominantly composed of approximately equal amounts of glucose and xylose (Figure 38A), but also appeared to contain trace amounts of mannose, arabinose and galactose (Figure 38B). This implied that Swollenin might also play a role in hemicellulose solubilization as well as cellulose disruption. Although it could be assumed that the solubilized xylose originated from the hemicellulosic portion of the biomass, it was unclear what fraction of the solubilized glucose-containing oligomers originated from the cellulose and what fraction may have originated from the hemicellulosic fraction. The partial solubilization of the hemicellulosic fraction of SPCS by Swollenin was particularly apparent when expressed as a percent solubilization of the hemicelluloses (Figure 38C). Further work indicated that Swollenin activity was temperature dependant with incubation at 20, 35, and 50 °C releasing slightly increasing amounts of sugar from the SPCS (Figure 39). A temperature increase from 20 °C to 50 °C resulted in slightly more than a 2-fold increase in sugar release. After determining that oligomers were being released from the SPCS after Swollenin treatment, these oligomers were further assessed using polyacrylamide carbohydrate electrophoresis (Goubet et al., 2011) in order to determine the size-distribution of these released oligomers. 160 Figure 38: Sugars detected after incubation of SPCS with either Swollenin or a BSA control. (A) Oligomeric and monomeric glucose and xylose release. (B) Amount of oligomeric arabinose, galactose and mannose detected after incubation with Swollenin. These sugars were not detected in monomeric form, and oligomers were undetectable in the BSA-treated samples. (C) Percent release of cellulosic and hemicellulosic sugars in a soluble form after incubation with Swollenin relative to the original glucan and total hemicellulose content of the pretreated substrate. Values are relative to a BSA control. All reactions were run in triplicate and error bars represent one standard deviation from the mean. Figure 39: Amount of monomeric and oligomeric glucose and xylose detected after incubation of Swollenin with SPCS between 20 °C and 50 °C. Total sugar represents the sum of all oligomeric and monomeric glucose and xylose. Values are relative to a BSA control, reactions were run in triplicate, and error bars represent one standard deviation from the mean. 02040608010012010 20 30 40 50 60Sugar Concentration (µg/mL)Temperature (°C)Total SugarOligomeric GlucoseOligomeric XyloseMonomeric XyloseMonomeric Glucose0102030405060708090100Sugar Concentration (µg/mL)BSA ControlSwollenin0123456% Release0123456Sugar Concentration (µg/mL)A B C 161 3.4.3.2.2 Polyacrylamide carbohydrate electrophoresis analysis As previous work had shown that the size and composition of soluble oligomers could be quantified by polyacrylamide carbohydrate electrophoresis (PACE) (Goubet et al., 2011; Jackson et al., 1994), the supernatants from SPCS treated with either Swollenin or a protein control (BSA) were compared to a series of sugar standards, including glucose, xylose, xylobiose, xylotriose, xylotetraose, and xylohexaose. The monomers and oligomers were conjugated to a polyanionic dye and separated by size on a polyacrylamide gel prior to imaging (Jackson et al., 1994). The gel profiles of the solubilized sugars are shown in Figure 40A, and were assessed through semi-quantitative densitometry analysis (Figure 40B). The percent increase in the release of the various sugars was subsequently estimated (Figure 40C). Swollenin treatment resulted in a pronounced increase in the amount of glucose, xylose, and xylotriose, released as well as an increase in cellobiose, xylobiose and xylotetraose. Swollenin treatment also caused a marked increase in the amount of higher molecular weight compounds that were detected in the supernatant. These higher molecular weight (HMW) compounds likely represent longer-chain (> 4), substituted xylan fractions. The lack of distinct bands in this HMW region suggested that these longer xylan chains were probably substituted by arabinose, glucuronic acid and acetyl groups, as these substituents are present within the glucuronoarabinoxylan of commelinid monocots such as maize. Swollenin treatment also resulted in the release of a low molecular weight compound that was absent in the control-treated sample. Although we postulated that this band might result from the release of acetyl groups, subsequent HPLC analysis of the Swollenin and control-treated supernatants did not show any increase in the amount of free acetic acid detected in the Swollenin-treated supernatants. 162 Figure 40: PACE analysis of released carbohydrates. (A) Carbohydrates released after incubation of Swollenin and a BSA control with SPCS. Standards are labeled; HMW represents the higher molecular weight fraction. (B) Densitometry analysis and peak selection. (C) Quantification of the relative increase in sugars released by Swollenin compared to the control. Values represent the percent increase in peak area for each sugar. Previous work had shown that Swollenin did not have significant hydrolytic activity on cellulose or on purified birchwood xylan (Jäger et al., 2011; Saloheimo et al., 2002). In the work reported here Swollenin alone was able to solubilize low levels of both the cellulosic and hemicellulosic components present in the SPCS. This suggested that the release of oligomers from SPCS observed here was due to the Swollenin-induced release of pre-existing oligomers non-covalently bound within the substrate. It is possible that this solubilization was due to the non-hydrolytic weakening of hydrogen bonding between the xylo- and cello-oligomers and the cellulose and/or xylan within the substrate in a manner similar to that suggested for the related Expansin family of proteins (McQueen-Mason and Cosgrove, 1994). Although the exact mechanism of action is unresolved, previous work suggested that the flexible FNIII-like repeats within Expansin-like region of Swollenin in may be involved in facilitating cellulose disruption (Hsin et al., 2011; Saloheimo et al., 2002). Glu Xyl Xyl2 Xyl3 Xyl4 Xyl6 HMW A Standards Swollenin Control B C0%100%200%300%400%163 The ability of Swollenin to solubilize oligomeric sugars from SPCS indicated that it may play an important role in the metabolism of T. reesei. Previous work has shown that Swollenin is constitutively expressed at low levels, unlike the more tightly repressed major cellulase enzymes (Margolles-clark et al., 1997; Saloheimo et al., 2002). As Swollenin treatment of SPCS results in the release of xylobiose and other xylo-oligomers which are known inducers of the major cellulases and xylanases (Aro et al., 2005; Margolles-clark et al., 1997), another possible function of Swollenin may be to release of soluble inducers upon contact with a lignocellulosic substrate, thereby leading to increased genetic transcription and subsequent up-regulation of the enzymes required for biomass degradation. 3.4.3.3 CBM adsorption to Swollenin-treated SPCS After assessing the solubilizing effects of Swollenin on SPCS, the residual solids were analyzed via the CBM adsorption method. In Section 3.2.5 of this work, a marked increase in cellulose accessibility was observed when Swollenin was applied to a model cellulosic substrate. The two-probe CBM adsorption technique revealed that this increase in cellulose accessibility was predominantly achieved through an increase in the accessibility of the more ordered regions of the cellulose. This, combined with SEM imaging, suggested that Swollenin was splitting/peeling/sloughing off roughened patches of the cellulose, revealing the better-ordered underlying regions of the cellulose (See Figures 23-25 in Section 3.2.5). When this CBM adsorption technique was applied to quantifying the manifestations of Swollenin treatment on SPCS, adsorption of CBM2a (representing crystalline cellulose) was found to increase slightly, while adsorption of CBM44 (representing amorphous cellulose) was found to decrease markedly (Figure 41). These results are consistent with the solubilization data suggesting that the less-ordered regions of the biomass are being disrupted and solubilized by Swollenin. This solubilization 164 would reduce binding to these disordered regions by CBM44 while simultaneously revealing new CBM2a binding sites in the underlying more-ordered regions of the biomass. Figure 41: Change in CBM adsorption to SPCS after Swollenin treatment. Absolute change in CBM binding is shown on the left, while percent change is shown on the right. Samples were run in triplicate and error bars represent one standard deviation from the mean. 3.4.4 Synergism between Swollenin and hydrolytic cellulase and xylanase monocomponents To further elucidate the role that Swollenin might play in SPCS deconstruction, several purified cellulase and xylanase enzymes were incubated with Swollenin and their possible synergistic interactions were assessed. β-glucosidase was also added to these combinations to decrease possible end-product inhibition that might arise from cellobiose accumulation (Murphy et al., 2013). When Swollenin was incubated with either an exoglucanase (Cel7A), an endoglucanase (Cel5A), or two endoxylanases (Xyn10A and Xyn11A) isolated from T. reesei as described previously (Hu et al., 2011), little or no synergism was observed in releasing glucose from the SPCS (Figure 42A). However, it was apparent that there was a strong synergistic interaction between Swollenin and Cel5A, Xyn10A, and Xyn11A toward releasing xylose from SPCS (Figure 42B). When the degree of synergism between Swollenin and the various hydrolytic enzymes were compared (Table 5), the most pronounced 020406080100120140CBM2a CBM44ug CBM Bound/mg BiomassControlSwolleninCBM2a CBM44-25-20-15-10-50510% Change in CBM Adsorption165 synergistic interaction occurred between Swollenin and Xyn11A where a more than 300% increase in xylose release was obtained when compared to the xylose released when the two proteins were added separately. It was apparent that Swollenin exhibited strong synergistic interactions with endoxylanases. Figure 42: The amount of glucose (A) and xylose (B) released from SPCS after incubation of various purified hydrolytic enzymes with either a BSA control or Swollenin. Reactions were run in triplicate and error bars represent one standard deviation from the mean. Overall, the most influential effect of Swollenin addition in the work reported here was the significant enhancement of xylan hydrolysis in comparison to cellulose hydrolysis. One possible explanation for this more marked effect on the hemicellulosic component is that Swollenin is able to promote amorphogenesis within the relatively loosely ordered xylan structure, but less able to disrupt the more highly-ordered structures of the cellulose. Related work has shown how significant a role xylan “masking” can play in hindering cellulose hydrolysis, and how beneficial the addition of xylanase can be 0100200300400500600700μg/mL Glucose050100150200250300350μg/mL XyloseB A 166 in enhancing fibre swelling and overall cellulose accessibility (Hu et al., 2011; Ishizawa et al., 2007; Liao et al., 2005; Zhang et al., 2011). Table 5: The degree of synergism (DS) between various purified hydrolytic enzymes and Swollenin on SPCS. Samples were run in triplicate and σDS represents one standard deviation. Glucose Xylose DS σDS DS σDS Cel7A + Swollenin 1.09 0.17 1.11 0.25 Cel5A + Swollenin 0.77 0.08 1.75 0.21 Xyn10A + Swollenin 1.15 0.10 2.75 0.17 Xyn11A + Swollenin 0.91 0.10 3.34 0.34 Non-hydrolytic, non-oxidative proteins such as Swollenin have been shown to promote the disruption, or amorphogenesis, of cellulosic substrates during the enzymatic deconstruction of biomass (Arantes and Saddler, 2010). Previous work has shown that Swollenin can aid in the disaggregation of model cellulosic substrates (Jäger et al., 2011; Saloheimo et al., 2002). The work reported here suggests that, when more industrially relevant substrates such as SPCS are used, the greater influence of Swollenin addition to a cellulase mixture may be in enhancing accessibility by disrupting and opening up the less ordered/hemicellulosic regions of the biomass, revealing the underlying more-ordered cellulosic portion. This reaffirms the idea that Swollenin promotes amorphogenesis through a targeted disruption of the less-ordered regions within cellulosic or lignocellulosic substrates. Evidence for Swollenin targeting less ordered regions comes from the work in Section 3.2 demonstrating an increase in the amount of accessible crystalline cellulose after Swollenin treatment promoted the sloughing of roughened cellulose particles from the surface of cellulosic fibres. Additionally, in Section 3.3 Swollenin was shown to target the disordered cellulose within fibre dislocations in dissolving pulp fibres. Finally, in this Chapter Swollenin was shown to promote the targeted disruption of the hemicellulosic fraction of SPCS, resulting in a slight increase in the amount of 167 accessible crystalline cellulose, while reducing the amount of accessible amorphous cellulose (likely via solubilization). Overall, these results strongly support Swollenin’s role in promoting amorphogenesis within the less-ordered regions of cellulosic and lignocellulosic biomass. While Swollenin was found to promote fragmentation of a model cellulosic substrate, dissolving pulp (Section 3.3.4), it did not promote amorphogenesis of woody lignocellulosic substrates. However, when Swollenin was applied to a grassy substrate, SPCS, the amorphogenesis-inducing capacity of Swollenin was evidenced by the solubilization of the portion of the biomass enriched in hemicellulosic sugars, as well as in enhancing the accessibility of the crystalline regions of the cellulose, while synergizing strongly with xylanases in the release of xylose. This substrate-specific activity of Swollenin suggests that it may act in a similar manner to the Beta-expansins, to which Swollenin is related, which act predominantly on Type II plant cell walls (those of the grasses and other commelinoid monocots) (Cosgrove, 2000). This implies that Swollenin acts predominantly on Type II cell walls and various amorphogenesis-inducing proteins, including the Expansins and Swollenin, may have finely-tuned substrate specificities. The substrate-specificity observed here is consistent with previous work which showed that the degree of synergism observed between various biomass-degrading enzymes varies, depending on the biomass substrate (Hu et al., 2013, 2014). This substrate-specificity could explain the fields relatively slow progress in developing a better in-depth understanding of the effects of these amorphogenesis-inducing proteins on lignocellulosic biomass. Although this finely-tuned specificity could potentially limit the fruitful application of such proteins in commercial enzyme mixtures, this work also suggests that proteins such as Swollenin and Expansins may be capable of promoting efficient amorphogenesis when paired to particularly receptive biomass substrates. Significant research effort has already been dedicated to producing robust 168 commercial cellulase mixtures capable of efficiently degrading all sources of biomass. It is possible that a more targeted approach to developing more effective enzyme cocktails will be to produce substrate-specific enzyme mixtures tailored for efficient deconstruction of particular biomass sources. The ability of specific amorphogenesis-inducing proteins to promote disruption on certain types of biomass could thus be efficiently exploited by incorporating these proteins into commercial mixtures tailored toward the specific biomass type that these amorphogenesis-inducing proteins act upon. Future work into exactly what combinations prove most effective will be required before this aspect of amorphogenesis can be effectively exploited. 3.4.5 Conclusions Swollenin was able to solubilize the less ordered regions of SPCS, with a particularly disruptive effect on the hemicellulosic fraction of this biomass. This disruptive/solubilizing effect was shown to promote a slight increase in the accessibility of the crystalline cellulose, while simultaneously promoting a reduction in the amount of accessible amorphous cellulose, likely via solubilization of this portion of the biomass. When various protein combinations were assessed Swollenin showed strong synergism with the xylan-degrading enzymes Xylanase GH10 and Xylanase GH11 in releasing xylose from SPCS. The previously-documented inability of Swollenin to directly hydrolyze pure cellulose or xylan, combined with the Swollenin-induced oligomer release and alterations in accessibility observed here, suggests that Swollenin weakens and disrupts the substrate by promoting amorphogenesis. This disrupting activity primarily facilitated the enzymatic hydrolysis of xylan, likely enhancing enzyme access to the cellulose component of pretreated corn stover. The work within this section of the thesis suggests that Swollenin specifically acts upon Type II (grassy) cell walls and that the primary influence of Swollenin treatment of SPCS is the targeted disruption of the less-ordered regions of this biomass, resulting in enhanced xylanase activity and improved accessibility to the crystalline cellulose regions. 169 4. Conclusions and future work 4.1 Conclusions The main focus of the thesis work was to investigate the non-hydrolytic, non-oxidative disruption of cellulosic biomass, with the goal of determining if amorphogenesis-inducing proteins could be used to enhance the accessibility and reactivity of biomass to the other enzymes involved in cellulose hydrolysis. By developing a better understanding of the interactions between these disruptive proteins and the various components of lignocellulosic substrates, it was expected that improved efficiencies in enzymatic deconstruction of biomass for conversion to fuels and chemicals could be achieved. The accessibility of the cellulosic portion of the biomass was found to play a key role in determining the enzymatic hydrolyzability, supporting the suggestion that the rate limiting step of enzymatic hydrolysis is not the actual depolymerisation of the carbohydrate chains, but rather the rate at which the enzymes can gain access to the chains embedded within the biomass. One of the key challenges when working with amorphogenesis-inducing proteins is being able to qualitatively or quantitatively define the end products of amorphogenesis. While hydrolytic enzymes release more readily-quantifiable soluble sugars from the cellulosic substrate, the amorphogenesis mechanism has often been defined as a delamination, splitting, peeling, swelling, or decrystallizing of the substrate which is far more challenging to qualify or quantify than the detection of soluble sugars. Previous researchers have tried to adapt various techniques to quantify these effects, such as using the water retention value, fibre quality analyzer, solute exclusion, mercury porosimetry and nitrogen adsorption. However, most of these techniques are crude, providing qualitative information at best and do not provide the quality of data necessary for quantifying amorphogenesis. Others techniques might provide more reproducible data but measure substrate features at the wrong scale (i.e. at the whole fibre scale using the fiber quality analyser (FQA), or at the sub-nanometer scale using Nitrogen 170 adsorption). In the work within the thesis we used the Simons’ Stain technique to provide a “base level” of cellulose accessibility. However, to try to quantify the finer changes occurring at the microfibril/cellulose surface scale an even more sensitive and selective technique was required. For this reason, the CBM adsorption technique was developed and refined to try to quantify the accessible crystalline cellulose and accessible amorphous cellulose. This technique in combination with methods such as Simon’s stain was used to better quantify the amorphogenesis process. A number of amorphogenesis-inducing proteins such as Swollenin, Loosenin and the Expansins were initially assessed for their potential disruptive abilities. Swollenin, which was made available to us through our ongoing collaboration with researchers at VTT Technical Research Center of Finland, proved to be the most promising amorphogenesis-inducing protein and was applied to a range of cellulosic materials to assess whether or not this protein was truly promoting amorphogenesis. When the CBM adsorption technique was used to quantify the disruptive effects, Swollenin was shown to target the less ordered regions of the cellulose, promoting its disruption and dispersion. Subsequent work involving fluorescent tagging of the CBMs and imaging revealed that Swollenin was targeting the disordered cellulose present within fiber dislocations. Although this initial work showed that Swollenin played a role in promoting the amorphogenesis of relatively pure cellulose we wanted to determine if it also played a role in the deconstruction of more industrially-relevant lignocellulosic substrates. Swollenin was applied to corn stover treated via the pseudo-pulping steam pretreatment process to determine how this protein would interact with an industrially relevant lignocellulosic biomass. As was observed on the pure cellulosic substrate, Swollenin promoted the dispersion and disruption of the less ordered regions of the corn stover, resulting in the solubilization of hemicellulosic sugars as well as a portion of the disordered cellulose. When synergistic interactions between Swollenin and separate cellulases and xylanases were assessed, surprisingly strong synergism was observed in the 171 latter case. The degree of synergism between Swollenin and xylanases was far higher than that between Swollenin and cellulases and greater than that previously reported between amorphogenesis-inducing proteins and hydrolases during the deconstruction of lignocellulosic biomass (Baker et al., 2000). In summary, we developed a new technique to quantify cellulose amorphogenesis induced by non-hydrolytic/non-oxidative proteins and this technique, in combination with other methods, indicated that the amorphogenesis-inducing protein Swollenin targets and disrupts the less ordered regions of cellulosic and lignocellulosic substrates, showing strong synergistic interactions with xylanases. 4.2 Future work 4.2.1 Screening for effective combinations of biomass, amorphogenesis-inducing proteins, and hydrolytic/oxidative enzymes The apparent specificity of Swollenin to disrupt corn stover should be investigated in more detail. Specifically, screening for interactions between amorphogenesis-inducing proteins such as Swollenin, Expansins, Loosenin, and others on industrially relevant feedstocks including grassy biomass such as wheat straw, corn stover, and sugarcane bagasse, as well as woody biomass including both hardwoods and softwoods should be assessed. Previous work has demonstrated that the Alpha-Expansins tend to act more effectively on woody substrates, while the Beta-Expansins are more effective on grassy biomass (Cosgrove, 2000). This substrate specificity should be assessed in more detail, and the potential biomass specificity of the various amorphogenesis-inducing proteins should be determined. Quantification of disruption should be carried out at the microscopic level using techniques such as the CBM-adsorption technique presented here, as well as measuring potential oligomer release. The CBM adsorption technique could be developed into a high-throughput microplate-based technique by performing simultaneous incubation of fluorescently tagged CBMs. Synergistic interactions between these amorphogenesis-inducing proteins and various hydrolytic and oxidative enzymes on this range of relevant biomass sources should prove interesting. This work would hopefully reveal trends as 172 well as suggesting effective combinations of deconstructing enzymes, amorphogenesis-inducing proteins and specific biomass feedstocks that could be exploited to enhance the overall efficiency of cellulose hydrolysis. 4.2.2 Broader application of tagged CBMs as molecular probes The diverse binding specificities of CBMs suggests that these proteins could be used to provide detailed maps of the surface morphology and surface composition of lignocellulosic fibres during a variety of thermochemical or enzymatic processes. For example, CBMs specific for certain hemicelluloses could be used to track the potential redistribution of these polymers during pretreatment processes or they could be used to quantify the reduction in hemicellulose at the fibre surface after treatment with hemicellulases. Within the scope of enzymatic biomass deconstruction, CBMs specific for particular hemicelluloses could be used to track the amount of each hemicellulose at the fibre surface during enzymatic deconstruction, thereby providing insights into the variable recalcitrance of particular hemicelluloses to enzymatic hydrolysis. By providing detailed “maps” of fibre surface morphology and surface composition, the use of CBMs as molecular probes could be used to develop a more detailed understanding of the redistribution and solubilization of the various carbohydrate polymers within lignocellulosic biomass during thermochemical and enzymatic processes. This information could then be used to tailor enzyme cocktails towards either targeted modification (for example, increasing fibrillation while trying to maintain strength) or deconstruction of particularly recalcitrant carbohydrates within the biomass. 4.2.3 Pulp and paper applications The ability of proteins such as Swollenin to open up and disrupt lignocellulosic substrates without solubilizing a significant portion of the biomass could be desirable for pulp and paper applications. Although the work presented in this thesis demonstrated that Swollenin did not appear to 173 significantly alter the macroscopic fibre properties of pretreated woody substrates, a large amount of previous work has demonstrated that Swollenin can in fact promote the swelling and dispersion of pure cellulosic fibres. Since the bulk of pulp and paper products are processed to significantly reduce the lignin content, it is likely that amorphogenesis-inducing proteins could promote significant swelling/disruption on these purer cellulosic fibres. To date, limited research has been performed to try to better understand the effects of amorphogenesis-inducing proteins on the applicability of cellulosic fibre streams for downstream pulp and paper applications. However, it seems likely that the swelling ability of a number of these proteins could enhance the absorbency of cellulosic fibre products, with potential implications for products such as tissue paper and other absorbents. Thus the ability of amorphogenesis-inducing proteins to enhance the suitability of certain cellulosic pulps for particular downstream applications should be investigated. 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Thesis/Dissertation
eng
Forestry
Vancouver : University of British Columbia Library
Attribution-NonCommercial-NoDerivs 2.5 Canada
Graduate
The role of amorphogenesis in the enzymatic deconstruction of lignocellulosic biomass
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