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The role of adsorbed enzymes in determining the hydrolysis kinetics of pretreated lignocellulosic biomass Mok, Yiu Ki 2015

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 The role of adsorbed enzymes in determining the hydrolysis kinetics of pretreated lignocellulosic biomass  by  YIU KI MOK  B.Sc., The University of British Columbia, 2012    A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS OF THE DEGREE OF   MASTER OF SCIENCE  in  THE FACULTY OF GRADUATE AND POSTDOCTORAL STUDIES (Forestry)    THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver)     April 2015    © Yiu Ki Mok, 2015   ii  Abstract The cost-effective production of sugars from biomass continues to remain challenging, partly due to the relatively high enzyme/protein loading required to effectively hydrolyze pretreated lignocellulosic substrates. Previous works have shown conflicting observations regarding the correlation between enzyme adsorption and the hydrolytic performance of an enzyme mixture. Unfortunately, it has proven difficult to accurately determine the roles of adsorbed enzymes during the hydrolysis of lignocellulosic substrates, in part because of the interference that protein determination methods encounter from the release of sugars and other biomass derived materials, the lack of a hydrolysis strategy for hydrolysis with only adsorbed enzymes and the use of “model” substrates in many studies.    To better understand the role that adsorbed enzymes play in cellulose deconstruction, it is important that we are able to accurately quantify protein distribution and enzyme performance. Various protein quantification assays were initially assessed for their ability to accurately and reproducibly quantify protein/enzymes during typical biomass hydrolysis conditions. However, the ninhydrin assay, which was the most promising assay due to its specificity for protein and compatibility with most compounds derived from lignocellulosic samples, still suffered from the incompatibility with sugar degradation products, long hydrolysis times and potentially wide-ranging standard deviations. To overcome these limitations, an accurate and rapid modified ninhydrin assay was developed which employed a sodium borohydride treatment to eliminate sugar interference followed by acid hydrolysis at 130oC, reducing the overall reaction time to 4 hours. iii    Utilizing the modified ninhydrin assay, the role of adsorbed enzymes in determining the rate and extent of hydrolysis of several different pretreated biomass substrates was then assessed. Once the distribution of enzymes reached equilibrium, after 60 minutes, those enzymes that were adsorbed or free in solution were separated by centrifugation and subsequently assessed for their ability to hydrolyze various cellulosic substrates at different enzyme loadings. It was apparent that the adsorbed enzymes were critically important as the removal of those enzymes in solution resulted in no significant decrease in the rate and extent of hydrolysis. By using the adsorbed enzymes, enzyme loadings could be reduced by up to 53% while resulting in similar hydrolysis yields.    iv  Preface All of the work presented henceforth was conducted by Yiu Ki Mok in the Forest Products Biotechnology/Bioenergy Laboratories at the University of British Columbia, Point Grey campus.  v  Table of contents Abstract ............................................................................................................................................ii Preface............................................................................................................................................. iv Table of contents .............................................................................................................................. v List of tables .................................................................................................................................. viii List of figures .................................................................................................................................. ix List of units and abbreviations ....................................................................................................... xii Acknowledgements ........................................................................................................................ xv Dedication ..................................................................................................................................... xvi 1. Background .......................................................................................................................... 1 1.1. Bioconversion of lignocellulosic biomass........................................................................ 4 1.1.1. Pretreatment .............................................................................................................. 5 1.1.2. Enzymatic hydrolysis ................................................................................................ 8 1.2. Factors affecting enzyme adsorption.............................................................................. 10 1.2.1. Enzyme factors........................................................................................................ 11 1.2.1.1. Influence of enzyme structure on adsorption ...................................................... 11 1.2.1.2. Influence of synergistic enzyme interactions on enzyme adsorption .................. 13 1.2.2. Substrate factors ......................................................................................................... 15 1.2.2.1. Influence of lignin on enzyme adsorption ........................................................... 16 1.2.2.2. Influence of hemicellulose on enzyme adsorption .............................................. 18 1.2.2.3. Influence of cellulose on enzyme adsorption ...................................................... 21 1.2.3. Physical factors ........................................................................................................... 24 1.2.3.1. Influence of temperature on enzyme adsorption ................................................. 24 1.2.3.2. Influence of pH on enzyme adsorption ............................................................... 25 1.3. Relationship between enzyme adsorption and hydrolysis performance ........................ 25 1.4. Challenges of accurate quantification of enzyme adsorption and performance............. 29 1.5. Assays for quantifying protein adsorption ..................................................................... 31 1.5.1. Gel-based assays ..................................................................................................... 31 1.5.2. Immunoassays ......................................................................................................... 33 vi  1.5.3. Protein labelling assays ........................................................................................... 34 1.5.4. Activity dependent assays ....................................................................................... 36 1.5.5. Total protein assays................................................................................................. 37 1.6. Thesis objectives ............................................................................................................ 43 2. Materials and methods ....................................................................................................... 45 2.1. Preparation of pretreated lignocellulosic biomass ......................................................... 45 2.2. Compositional analysis of pretreated biomasses ............................................................ 45 2.3. Cellulase enzymes .......................................................................................................... 46 2.4. Enzymatic hydrolysis ..................................................................................................... 46 2.4.1. Two-stage hydrolysis .............................................................................................. 47 2.4.2. Multi-stage hydrolysis ............................................................................................ 48 2.5. Development of a modified ninhydrin assay.................................................................. 51 2.5.1. Traditional acidic ninhydrin assay .......................................................................... 51 2.5.2. Evaluating possible glucose interference of the traditional ninhydrin assay .......... 51 2.5.3. Elimination of glucose interference with sodium borohydride............................... 52 2.5.4. Influence of sodium borohydride on protein measurements................................... 53 2.5.5. Reduction of time required for complete protein hydrolysis .................................. 53 2.5.6. The development of a modified ninhydrin assay to quantify protein under lignocellulosic biomass hydrolysis conditions........................................................ 54 2.5.7. Compatibility of the modified ninhydrin assay with compounds encountered during lignocellulosic biomass hydrolysis ......................................................................... 55 2.5.8. Elucidation of possible interfering effects of individual lignocellulosic components   ................................................................................................................................. 56 2.5.9. Comparison of Current Protein Assays and the Modified Ninhydrin Assay .......... 56 3. Results and discussion ....................................................................................................... 58 3.1. Development of a NaBH4 coupled ninhydrin based assay for the quantification of protein/enzymes during the enzymatic hydrolysis of pretreated lignocellulosic biomass 58 3.1.1. Background ............................................................................................................. 58 3.1.2. Evaluation of glucose interference with the ninhydrin assay ................................. 61 3.1.3. Overcoming sugar interference ............................................................................... 63 3.1.4. Compatibility of sodium borohydride with protein ................................................ 66 vii  3.1.5. Reduction of hydrolysis time to achieve complete protein hydrolysis ................... 69 3.1.6. Compatibility with lignocellulosic biomass hydrolysis conditions ........................ 72 3.1.7. Comparison of the modified ninhydrin assay versus current total protein quantification assays ............................................................................................... 75 3.1.8. Conclusions ............................................................................................................. 77 3.2. The relative roles that adsorbed and free enzymes play in determining the rate and extent of hydrolysis of lignocellulosic biomass and the potential of a two-stage hydrolysis strategy to reduce enzyme loadings .................................................................................. 78 3.2.1. Background ............................................................................................................. 78 3.2.2. Time course of enzymatic hydrolysis of DsP ......................................................... 80 3.2.3. Two-stage hydrolysis strategy ................................................................................ 81 3.2.4. Selective removal of free enzymes during the hydrolysis of DsP .......................... 82 3.2.5. Effect of the selective removal of free enzymes on the enzymatic hydrolysis of lignocellulosic biomass ........................................................................................... 84 3.2.6. Effect of selective free enzyme removal at different enzyme loadings on enzymatic hydrolysis ................................................................................................................ 89 3.2.7. Effect of the complete removal of free enzymes on the enzymatic hydrolysis of pure cellulosic and lignocellulosic substrates ......................................................... 92 3.2.8. The potential to reduce enzyme loadings by using a two-stage hydrolysis strategy ..   ................................................................................................................................. 94 3.2.9. Conclusions ............................................................................................................. 98 4. Final conclusions and future work ................................................................................... 100 4.1. Conclusions .................................................................................................................. 100 4.2. Future work .................................................................................................................. 101 4.2.1. The influence of initial enzyme distribution in determining the rate and extent of hydrolysis at high solids loading........................................................................... 101 4.2.2. Potential recycling of the free enzymes for further rounds of hydrolysis ............. 102 4.2.3. Further assessment of the role of adsorbed enzymes on different cellulose morphologies and pretreated lignocellulosic substrates ....................................... 103 References ................................................................................................................................... 104   viii  List of tables Table 1. Summary of the major pretreatment processes used to enhance the fractionation and enzymatic hydrolysis of biomass .................................................................................................... 6 Table 2. Initial distribution of cellulase enzymes across different model and lignocellulosic substrates and total conversion obtained after hydrolysis............................................................. 27 Table 3. Compatibility of different protein assays with lignocellulosic biomass hydrolysis conditions ...................................................................................................................................... 42 Table 4. Pretreatment conditions for various biomass substrates used in this study. ................... 45 Table 5. Concentration of monomeric sugars present in commercial enzyme preparations. ....... 68 Table 6. Compositional analysis of WSFs from steam pretreated substrates. .............................. 72 Table 7. Steam pretreatment conditions and chemical composition of pretreated lignocellulosic substrates ....................................................................................................................................... 85 Table 8. Comparison of enzyme performance of a complete enzyme mixture and adsorbed enzymes after 72 hours of hydrolysis ........................................................................................... 91 Table 9. Total protein recovery after separation of adsorbed and free enzymes after 1 hour of hydrolysis at 4oC. .......................................................................................................................... 92 Table 10. The productivity and performance of a complete enzyme mixture, Cellic CTec3 as compared to adsorbed enzymes after the 72 hour of hydrolysis of pure cellulosic and lignocellulosic substrates .............................................................................................................. 94   ix  List of figures Figure 1. Proposed mechanism for cellulose amorphogenesis/depolymerization by cellulases (Arantes and Saddler, 2010). .......................................................................................................... 9 Figure 2. The complexity of lignocellulosic biomass (Rubin, 2008) ........................................... 15 Figure 3. 2D gel electrophoresis of a commercial cellulase preparation, Cellic CTec3, from Novozymes. .................................................................................................................................. 30 Figure 4. Two-stage hydrolysis strategy to separate free and adsorbed enzymes after 1 hour of hydrolysis with the complete enzyme mixture of Cellic CTec3. All hydrolysis experiments were conducted at 2% w/v solids loading.............................................................................................. 48 Figure 5. Multi-stage hydrolysis strategy to retain only adsorbed enzymes for an additional round of hydrolysis from a commercial enzyme preparation, Cellic CTec3, after 1 hour of initial hydrolysis with the complete enzyme mixture. All hydrolysis experiments were conducted at 2% w/v................................................................................................................................................. 50 Figure 6. Effect of increasing glucose concentration on the measured protein concentration of BSA by the traditional ninhydrin assay. ....................................................................................... 63 Figure 7. (a) Influence of different NaBH4 treatment conditions on the absorbance response of ninhydrin in the presence of 20g/kg glucose (b) Interference of sugars with and without NaBH4 treatment at a ratio of 1:3 NaBH4:sugar (w/w) for 60 minutes. .................................................... 65 Figure 8. Comparison of the protein concentration determined after BSA (20g/kg glucose) was diluted in 50mM sodium acetate buffer, using either the traditional ninhydrin assay or the traditional ninhydrin assay after NaBH4 treatment. ...................................................................... 67 x  Figure 9. Comparison of total protein concentrations of different commercial enzyme preparations quantified using the traditional ninhydrin assay with and without NaBH4 treatment........................................................................................................................................................ 69 Figure 10. Comparison of the time required to achieve complete hydrolysis of different proteins/enzyme preparations at 100oC and 130oC. (a) bovine serum albumin (b) Cellic CTec2, a commercial cellulase preparation (c) HTec, a commercial xylanase preparation. ....................... 70 Figure 11. The effect of different hydrolysis conditions on the total protein concentrations of different enzyme preparations. Enzyme preparations were subjected to NaBH4 treatment and hydrolyzed at 100oC for 24 hours or 130oC for 2 hours with 6M HCl before quantification by the ninhydrin assay. ............................................................................................................................ 71 Figure 12. Quantification of total protein concentration of Cellic CTec3 diluted in the WSFs from steam pretreatment by the modified ninhydrin assay. .......................................................... 75 Figure 13. Quantification of total protein concentration of Cellic CTec3 diluted in different WSFs obtained after steam pretreatment using various protein assays. ....................................... 76 Figure 14. Time course of hydrolysis of DsP after the selective removal of different proportions of free enzymes after 1 hour of hydrolysis at a solids loading of 2% w/v, 50oC with an enzyme loading of 30mg Cellic CTec3/g glucan. DsP: dissolving pulp. ................................................... 81 Figure 15. Time course of hydrolysis after the selective removal of 75% of free enzymes in solution after 1 hour of hydrolysis. (a) SPP200 (7.5mg/g Cellic CTec3/g glucan). (b) SPP190 (11.5mg/g Cellic CTec3/g glucan). (c) LPP200 (60mg/g Cellic CTec3/g glucan). SPP200: steam pretreated poplar at 200oC, 5min, 3% SO2; SPP190: steam pretreated poplar at 190oC, 5 min, 3% SO2; LPP200: steam pretreated lodgepole pine at 200oC, 5 min, 4% SO2. .................................. 88 xi  Figure 16. Cellulose hydrolysis of a range of lignocellulosic biomass at different enzyme loadings after 72 hour using a commercial enzyme mixture as compared to two-stage hydrolysis. DsP: dissolving pulp; SPP190: steam pretreated poplar at 190oC, 5 min, 3% SO2; SPP200: steam pretreated poplar at 200oC, 5 min, 3% SO2; LPP200: steam pretreated lodgepole pine at 200oC, 5 min, 4% SO2.................................................................................................................................. 89 Figure 17. Relationship between enzyme loading and total glucan conversion of a complete enzyme mixture (♦) as compared to the adsorbed and residual free enzymes (■) after separation. (a) DsP. (b) SPP200. (c) SPP190. (d) LPP200. DsP: dissolving pulp; SPP200: steam pretreated poplar at 200oC, 5min, 3% SO2; SPP190: steam pretreated poplar at 190oC, 5 min, 3% SO2; LPP200: steam pretreated lodgepole pine at 200oC, 5 min, 4% SO2. ........................................... 96 Figure 18. Relative reduction in enzyme loading to achieve 70% glucan conversion when using a complete enzyme mixture as compared to the adsorbed and residual free enzymes after separation. DsP: dissolving pulp; SPP200: steam pretreated poplar at 200oC, 5min, 3% SO2; SPP190: steam pretreated poplar at 190oC, 5 min, 3% SO2; LPP200: steam pretreated lodgepole pine at 200oC, 5 min, 4% SO2. ...................................................................................................... 97    xii  List of units and abbreviations A  absorbance AFEX  ammonia fibre expansion AIL  acid insoluble lignin am  molar extinction coefficient Ara  arabinan ARP  ammonia recycle percolation BCA  bicinchoninic acid b.d.l  below detection limit βG  β-glucosidase BSA  bovine serum albumin c  concentration of protein  CBM  carbohydrate binding modules CBH  cellobiohydrolases CD  catalytic domain cm  centimetre CNC  cellulose nanocrystals DOC  depends on concentration of selected compound DP  degree of polymerization DsP  dissolving pulp DW  dry weight EG  endoglucanases ELISA  enzyme-linked immunosorbent assay FPA  filter paper assay FPU  filter paper unit xiii  g  gram Gal  galactan Glu  glucan H2SO4  sulfuric acid HCl  hydrochloric acid HMF  5-hydroxymethylfurfural HPLC  high performance liquid chromatography hr  hour IUPAC International Union of Pure and Applied Chemistry kDa  kilo-Dalton kg  kilogram l   length of the path length in centimeters L  litre LCC  lignin–carbohydrate complexes LPMO  lytic polysaccharide monooxygenase LPP200 steam pretreated lodgepole pine at 200oC, 5 min, 4% SO2 IEF  isoelectric focusing mAb  monoclonal anitbody Man  mannan mg  milligram min  minute mL  millilitre mM  millimole per litre M  mole per litre NaBH4  sodium borohydride xiv  NaOH  sodium hydroxide n.d.  not determined nm  nanometer NREL  National Renewable Energy Laboratory PASC  phosphoric acid swollen cellulose Phe  phenolics pI  isoelectric point RMP  refiner mechanical pulps SDS-PAGE sodium dodecyl sulfate polyacrylamide gel electrophoresis SEM  scanning electron microscopy SO2  sulfur dioxide SPCS  steam pretreated corn stover SPLP  steam pretreated lodgepole pine SPP  steam pretreated poplar SPP190 steam pretreated poplar at 190oC, 5 min, 3% SO2 SPP200 steam pretreated poplar at 200oC, 5 min, 3% SO2 T  temperature TAPPI  Technical Association of the Pulp and Paper Industry UV  ultraviolet v/v  volume/volume WSF  water soluble fraction w/v  weight/volume w/w  weight/weight Xyl  xylan   xv  Acknowledgements I would like to express my deep gratitude to my supervisor, Dr. Jack Saddler, for his mentorship, guidance, encouragement throughout my MSc studies. I am grateful for the chance to work with you and have learnt many invaluable lessons along the way. I would also like to thank my committee members Dr. Heather Trajano and Dr. Valdeir Arantes as well as Dr. Jinguang Hu for their coaching and countless discussions regarding my work. Special thanks also goes to previous and current members of the Forest Products Biotechnology/Bioenergy (FPB) group for the peer tutoring and friendship, which made every day in the lab a fun and memorable experience for the past 3 years. It is amazing how fast time passes by! Last but not least, I would like to thank my family for their unwavering support throughout my study.    xvi  Dedication This thesis is dedicated to the memory of my beloved grandmother, Ms. Leung Kwai Ying, who passed away before its completion.  1  1. Background With increasing concerns about global sustainability, the transition from today’s fossil based economy towards a more sustainable bio-based economy has received significant interest across the world (Menon and Rao, 2012; Bozell and Petersen, 2010; Ragauskas et al., 2006). A key factor for the successful transition towards a bio-based economy is the development of biorefineries that allow for the efficient processing of lignocellulosic biomass to establish “sugar-platforms” that will form the basis of a sustainable biofuels and biochemicals sector. However, the cost-effective production of sugars from lignocellulosic biomass is challenging due to the significant amount of enzymes needed to achieve efficient saccharification of cellulose and hemicellulose to monomeric sugars (Klein-Marcuschamer et al., 2012; Stephen, Mabee and Saddler, 2012). In order to reduce enzyme dosages, different strategies have been pursued, such as improving enzyme catalytic activity/thermostability, enhancing the synergism among cellulase enzymes and applying various accessory enzymes/disrupting proteins to cellulase mixtures (Hu, Arantes and Saddler, 2011; Wilson, 2009; Zhang et al., 2006). Although reductions in enzyme dosages have been achieved to varying degrees of success with these strategies, additional efforts will still be required to further lower the enzyme costs if these processes are ever to become commercially successful (Hong et al., 2013; Klein-Marcuschamer et al., 2012; Stephen, Mabee and Saddler, 2012). Several researchers have suggested that it is the lack of accessibility of cellulase enzymes to the cellulosic substrate which most limits the efficiency of enzymatic hydrolysis (Gourlay, Arantes and Saddler, 2012; Arantes and Saddler, 2011; Chandra et al., 2008; Hong, Ye and Zhang, 2007; Grethlein, 1985). One interesting aspect of this work was that the removal of any unabsorbed “free enzymes” had limited influence on the overall hydrolysis 2  performance of the “cellulase mixtures” on pure cellulosic substrates, implying that the important enzymes were the ones that had been adsorbed to the substrate (Yu et al., 2013; Lee and Fan, 1983; Mandels, Kostick and Parizek, 1971). Therefore, one of the main goals of the work described in this thesis was to try to better understand the role and function of the adsorbed and the “free” enzymes as one strategy for reducing the required amount of enzyme needed to achieve effective cellulose hydrolysis. Cellulose hydrolysis is a complex process, where multiple enzyme components must interact with complex insoluble substrates (Yang et al., 2011; Zhang and Lynd, 2004; Lynd et al., 2002). Although significant research efforts have been invested to try to better assess the dynamics of enzyme-substrate interactions and its influence on enzymatic hydrolysis, many of these studies have hydrolyzed model cellulosic substrates, such as Avicel, cotton and filter paper using either purified enzyme components or older generations of cellulase preparations (Pakarinen et al., 2014; Gao et al., 2013; Varnai, Siika-Aho and Viikari, 2013; Van Dyk and Pletschke, 2012; Wang et al., 2012; Arantes and Saddler, 2011; Kristensen, Felby and Jorgensen, 2009; Lynd et al., 2002; Mansfield, Mooney and Saddler, 1999; Klyosov, 1990). As will be discussed in more detail later, the adsorption and subsequent hydrolysis of lignocellulosic biomass by “cellulase enzymes” is strongly influenced by the nature of the individual enzymes, the enzyme mixture and the physicochemical nature of the cellulosic substrate (Van Dyk and Pletschke, 2012; Yang et al., 2011; Mansfield, Mooney and Saddler, 1999). These “model” enzymes and substrate conditions are unlikely to represent industrially relevant conditions where lignocellulosic substrates containing cellulose, hemicellulose and lignin as well as complex enzyme preparations containing many cellulases and accessory enzymes will be used. Thus, 3  much more representative work is required if we are to better understand how enzyme adsorption influences the rate and yield of lignocellulosic hydrolysis. To further complicate the challenges of studying enzyme adsorption, there is a lack of an effective and accurate method to quantify enzymes/protein under typical lignocellulosic biomass hydrolysis conditions (Adney et al., 2012; McMillan et al., 2011; Adney et al., 1996). Virtually all of the current protein assay methods display limitations when performed under realistic conditions, in part due to the complex natures of both the enzyme mixtures and the biomass substrates. Current commercial enzyme preparations are highly complex mixtures, which contain multiple cellulase components as well as various accessory enzymes/disrupting proteins, such as hemicellulases, swollenin and lytic polysaccharide monooxygenases (LPMOs) (Gupta and Lee, 2013). The different amino acid compositions and the physical properties of the enzymes within a “cellulase cocktail” can further influence the measurement of total protein as most assays rely on different chemical mechanisms, which will be discussed later in the thesis (González-González et al., 2011; Kipper, Valjamae and Johansson, 2005; Owusu-Apenten and FOODnetBASE, 2002; Fountoulakis, Juranville and Manneberg, 1992; Legler et al., 1985; Chou and Goldstein, 1960). In addition, other potential interfering compounds derived both from pretreatment and enzymatic hydrolysis, such as monosaccharides, oligosaccharides, phenolic compounds, and sugar degradation products may also influence traditional protein assays (Ximenes, Kim and Ladisch, 2013; Chylenski et al., 2012; Kim et al., 2011; Ximenes et al., 2011; Ximenes et al., 2010; Zhang and Lynd, 2004; Nieves et al., 1997). As a result, significant differences (>90%) have been reported when various protein quantification methods have been used to determine enzyme distribution and performance during the hydrolysis of lignocellulosic substrates (Adney et al., 1996). One of the main goals of the thesis was to assess the potential to 4  further reduce the enzyme loadings required to achieve effective lignocellulose hydrolysis by better understanding how adsorbed enzymes might influence the rate and extent of hydrolysis of various biomass substrates. To try to better elucidate the roles of adsorbed enzymes, a more accurate way of assessing enzyme adsorption and performance had to be developed. Of the various assays that had previously been used in the literature to quantify cellulases, the ninhydrin assay appeared to be the most promising. However, there were several drawbacks with this assay, such as long hydrolysis times, potentially high standard deviations and interference by sugar degradation products (Haven and Jorgensen, 2013; Starcher, 2001; Schilling, Burchill and Clayton, 1963). Thus, an initial objective of the work was to improve the compatibility of the ninhydrin assay with lignocellulosic biomass processing conditions. After developing the modified ninhydrin assay, the roles and functions of adsorbed enzymes were next assessed over a range of industrially relevant lignocellulosic substrates. As mentioned above, many of the previous studies which had investigated the relationship between enzyme adsorption and the hydrolytic performance of complete enzyme mixtures did not represent current industrially relevant conditions. It was hoped that, by assessing the adsorption and kinetics of a state-of-the-art Novozymes commercial “cellulase cocktail” Cellic CTec3 during the time course of hydrolysis on a range of pretreated biomass substrates, this might provide us with invaluable information to allow a reduction in the enzyme loading required to achieve effective lignocellulose hydrolysis. 1.1. Bioconversion of lignocellulosic biomass As mentioned above, the development of biorefineries will require the establishment of an inexpensive and efficient biomass to sugar platform. Currently, the bioconversion of 5  lignocellulosic biomass to sugars consists of multiple processes. In general, the biomass is first pretreated followed by the enzymatic hydrolysis of the cellulosic component to produce sugars, which are then converted to fuels and chemicals through biological or chemical routes (Kumar, Singh and Singh, 2008; Galbe and Zacchi, 2007; Chandra et al., 2007). In the following sections, various pretreatment technologies and the enzymatic hydrolysis process are briefly reviewed.  1.1.1. Pretreatment Due to the recalcitrant nature of lignocellulosic biomass, a pretreatment step is often required to modify its physical and chemical properties to increase the accessibility of cellulose to cellulases (Chandra et al., 2007; Galbe and Zacchi, 2007). It is recognized that the pretreatment step is one of the most important process steps in bionconversion as the structure and composition of the substrate can have a significant influence on downstream processes, including hydrolysis and fermentation (Arantes and Saddler, 2011; Chandra et al., 2007; Mansfield, Mooney and Saddler, 1999). In general, pretreatment techniques can be categorized into four different categories: physical, biological, chemical and physicochemical pretreatments.  Physical pretreatment methods are characterized by the comminution of lignocellulosic biomass feedstocks to smaller particles, which can be facilitated using a combination of chipping, grinding, and/or milling (Kumar, Singh and Singh, 2008; Galbe and Zacchi, 2007; Mais et al., 2002). By reducing the size of the substrate, both reduction in cellulose crystallinity and increase in the specific surface area of cellulose are achieved, resulting in a substrate that is more amenable to enzymatic hydrolysis by cellulases (Chandra et al., 2007; Galbe and Zacchi, 2007). However, this process is highly energy demanding and the pretreated biomass is still highly recalcitrant to enzymatic hydrolysis. In contrast, biological methods employ the use of 6  different types of rot fungi, such as brown rot, soft rot and white rot fungi, to degrade/open up lignocellulosic materials (Galbe and Zacchi, 2007; Martinez et al., 2005; Hammel et al., 2002). Although biological processes are performed at milder conditions, they suffer from long processing times (multiple weeks/months) and the potential loss of sugars for metabolic and growth purposes (Chandra et al., 2007). Chemical pretreatments typically involve the application of chemical reagents to lignocellulosic biomass with the primary goal of increasing the accessibility of cellulose to cellulases by solubilizing hemicellulose and lignin and to a lesser degree, decreasing the degree of polymerization and crystallinity of the cellulose (Chandra et al., 2007; Galbe and Zacchi, 2007). However, large amounts of chemicals are typically required to achieve an effective treatment in processes analogous to pulping. Alternatively, a combination of physical and chemical treatments (physicochemical pretreatments) can be used to facilitate the chemical pretreatment process. Commonly employed physicochemical pretreatments include ammonia fibre explosion (AFEX), ammonia recycle percolation (ARP), liquid hot-water treatment and steam explosion (Meng and Ragauskas, 2014; Chandra et al., 2007). The different pretreatment strategies and their major of actions are summarized in Table 1.  Table 1. Summary of the major pretreatment processes used to enhance the fractionation and enzymatic hydrolysis of biomass Pretreatment Examplesa Major mode of action Biological Brown/white rot fungi Degradation of hemicellulose and lignin Chemical Alkali; Dilute acid; Ionic liquid; Organosolv Decrease DP and crystallinity Solubilisation of hemicellulose and lignin Physical Ball milling; Hammer milling Reduction of particle size and crystallinity Physicochemical AFEX; ARP; liquid hot water; steam explosion Disruption of lignin–carbohydrate complexes (LCCs) Decrease DP and crystallinity Solubilisation of hemicellulose aAmmonia fibre explosion: AFEX; Ammonia recycle percolation: ARP; 7  Among the different potential pretreatment methods, steam pretreatment, with the addition of a chemical catalyst, has been widely studied as it has been shown to be able to produce relatively easily hydrolyzable substrates from a range of different potential lignocellulosic feedstocks (Arantes and Saddler, 2011; Chandra et al., 2007). This process typically involves impregnating biomass with an acid-based catalyst, such as H2SO4 or SO2 followed by treatment with high-pressure saturated steam for a short period of time and subsequent release of pressure (Galbe and Zacchi, 2007). As a result, hemicellulose is primarily solubilized while retaining the majority of the cellulose, enhancing the ease of enzymatic hydrolysis by improving the accessibility of cellulose to cellulases (Chandra et al., 2007; Galbe and Zacchi, 2007).  Depending on the pretreatment conditions and biomass employed, substrate compositions containing varying amounts of cellulose, hemicellulose and lignin are produced (Bura, Chandra and Saddler, 2009; Chandra et al., 2007; Ewanick, Bura and Saddler, 2007; Bura et al., 2003). At lower severity treatments, hemicellulose is preserved in an insoluble form, which increases the overall sugar recovery but, the cellulosic component is typically more recalcitrant to hydrolyze due to limited accessibility of the resulting substrate. In contrast, although high severity treatments result in easily-digestible cellulosic substrates, it also suffers from a significant loss of hemicellulose sugars as well as the formation of inhibitory products that restrict downstream biological processes (Öhgren et al., 2007; Palmqvist et al., 1996). As a result, most studies typically employ compromised conditions that maximize sugar recovery while producing a cellulose-rich substrate for subsequent enzymatic hydrolysis (Arantes and Saddler, 2011; Kumar et al., 2010; Öhgren et al., 2007; Bura et al., 2003). As enzyme adsorption is necessary prior to catalytic action, it has been shown that the physicochemical characteristics of the substrates after 8  pretreatment strongly influences enzyme adsorption and desorption profiles as well as the performance of subsequent hydrolysis (Van Dyk and Pletschke, 2012; Mansfield, Mooney and Saddler, 1999). 1.1.2. Enzymatic hydrolysis In nature, cellulases can be produced by both fungi and bacteria. Depending on the source organism, cellulases may be produced as part of a complexed cellulase system (cellulosome) or non-complexed cellulase system (free enzymes) (Lynd et al., 2002). Some anaerobic bacteria and fungi can produce cellulosomes, which are large extra-cellular enzyme complexes that extend from the cell wall (Lynd et al., 2002). In contrast, most aerobic fungi and bacteria typically secrete a battery of non-complexed cellulase enzymes that allow for the efficient hydrolysis of cellulose. Cellulases from aerobic fungi, particularly the non-complexed cellulase system from the filamentous fungus Trichoderma reesei, have been the focus of considerable research (Lynd et al., 2002). According to the Carbohydrate Active Enzymes Database (http://www.cazy.org/CAZY), the enzymes that degrade the glycosidic bond between two or more carbohydrates or between a carbohydrate and a non-carbohydrate moiety are classified as glycosyl hydrolases. The cleavage of glycosidic bonds occurs via an acid/base catalysis using either a retaining or inverting mechanism (Davies and Henrissat, 1995). The retaining mechanism consists of a two-step substitution reaction and maintains the stereochemistry of the anomeric carbon in the initial substrate whereas the inverting mechanism is a one-step substitution, resulting in the inversion of the anomeric carbon’s stereochemistry (Davies and Henrissat, 1995). 9  As mentioned earlier, efficient cellulose hydrolysis requires multiple cellulase mono-components to work together in a synergistic manner. In total, T. reesei produces two cellobiohydrolases (CBH I, CBH II), five endoglucanases (EG I-V) and two β-glucosidases (BGL I-II) (Lynd et al., 2002). The general cellulose hydrolysis process can be divided into two major phases (Figure 1) (Arantes and Saddler, 2010; Lynd et al., 2002).   Figure 1. Proposed mechanism for cellulose amorphogenesis/depolymerization by cellulases (Arantes and Saddler, 2010). The primary phase, which is slow and hypothesized to be the rate-limiting step of the hydrolysis, consists of at least 2 steps to hydrolyze the insoluble cellulose to soluble cello-oligosaccharides. Briefly, a hypothesized amorphogenesis step first occurs to loosen the crystalline cellulose matrix and increase the accessibility of the cellulose chains to the cellulase 10  enzymes (Arantes and Saddler, 2010). The hydrolysis of the loosened/swelled cellulose chain to soluble cello-oligosaccharides is then facilitated by the synergistic actions of endoglucanases and cellobiohydrolases (Lynd et al., 2002). Specifically, endoglucanases participate in the random cleavage of internal amorphous regions of cellulose, generating new chain ends which cellobiohydrolases can processively act upon to produce soluble-oligosaccharides. Following the formation of soluble-oligosaccharides, a rapid secondary phase of hydrolysis then occurs and the glucose is quickly produced by the combination of cellulases and β-glucosidase (Arantes and Saddler, 2010; Lynd et al., 2002).  In addition to these canonical cellulolytic enzymes, current commercial enzyme mixtures often include other accessory enzymes, such as hemicellulolytic enzymes and lytic polysaccharide monooxygenases (LPMOs), which have been shown to greatly facilitate the hydrolytic potential of cellulase enzymes over a range of cellulosic/lignocellulosic substrates (Gupta and Lee, 2013; Hu, Arantes and Saddler, 2011; Harris et al., 2010; Kumar and Wyman, 2009; Selig, Weiss and Ji, 2008; Garcia-Aparicio et al., 2007). However, with the development of increasingly complex enzyme mixtures, the elucidation of the relationship between the hydrolytic performance of cellulases and enzyme adsorption will likely be further complicated by potential interactions among various enzymes. The challenge of accurately monitoring the adsorption and desorption of multiple enzyme components is discussed below. 1.2. Factors affecting enzyme adsorption As previously mentioned, the enzymatic hydrolysis of cellulose is a heterogeneous process that involves multiple soluble enzyme components acting on an insoluble substrate (Lynd et al., 2002). Thus, the diffusion and adsorption of cellulases to the solid substrate is a key 11  prerequisite prior to their catalytic action. In general, the composition/source of enzymes, structural properties of the substrates and hydrolysis conditions can have a significant impact on enzyme adsorption/desorption profiles and the overall hydrolysis. 1.2.1. Enzyme factors As described previously, the “cellulase cocktail” contains different types of enzymes working synergistically to try to achieve an efficient cellulose hydrolysis process. In addition, the individual enzyme components usually include multi-domain structures (Gilkes et al., 1991). It has been shown that the interactions between different domains as well as among individual enzyme components do influence the enzyme adsorption/desorption profiles (Payne et al., 2013; Boraston et al., 2004; Davies and Henrissat, 1995; Linder et al., 1995; Reinikainen et al., 1992; Ståhlberg, Johansson and Pettersson, 1991; Tomme et al., 1988; Van Tilbeurgh et al., 1986). 1.2.1.1. Influence of enzyme structure on adsorption Most non-complex cellulase enzymes are modular proteins consisting of a catalytic domain and carbohydrate binding module (CBM) that is connected by a linker protein (Gilkes et al., 1991). Among the different domains, the CBM has been reported to play a significant role in facilitating the formation of enzyme-substrate complexes (Boraston et al., 2004; Linder et al., 1995; Ståhlberg, Johansson and Pettersson, 1991; Tomme et al., 1988; Van Tilbeurgh et al., 1986). It has been shown that CBMs can: (1) increase the enzyme concentration on the surface of the substrate (proximity function); (2) determine substrate specificity (targeting function); and (3) might potentially non-hydrolytically disrupt crystalline substrates (disruptive function) (Arantes and Saddler, 2010; Boraston et al., 2004). For fungal CBMs, the interaction between CBMs and cellulose is usually facilitated by a flat or platform-like hydrophobic surface arising 12  from conserved aromatic amino acids (Boraston et al., 2004). Previous studies have shown that mutagenesis of the conserved aromatic amino acids significantly influences both the adsorption and activity of some cellulase enzymes (Linder et al., 1995; Reinikainen et al., 1992). Other studies have also investigated the effect of the removal of the CBM from cellulases and reported that CBM-lacking enzymes have reduced affinity and hydrolytic activity towards insoluble crystalline substrates, such as Avicel (Ståhlberg, Johansson and Pettersson, 1991; Tomme et al., 1988; Van Tilbeurgh et al., 1986). Based on these studies, it is clear that CBMs play an important role in enzyme-substrate interactions and the mode of hydrolysis of insoluble cellulosic substrates. In addition to the CBM, the catalytic domain (CD) also appears to influence cellulase adsorption. Depending on their hydrolytic action, different cellulase enzymes can have a pocket (or crater), cleft (or groove) or tunnel structure (Davies and Henrissat, 1995). For example, β-glucosidases have a pocket structure that is adapted to substrates containing a large number of available chain ends, such as cellobiose whereas endoglucanases have a cleft shape active site that facilitates its random binding to amorphous cellulose (Davies and Henrissat, 1995). In contrast, cellobiohydrolases have a tunnel shape that allows for processive action on the cellulose chain, with CBH I and CBH II having a preference for the reducing end and non-reducing end of cellulose respectively (Davies and Henrissat, 1995). This difference in active site topology has also been shown to affect the binding of cellulases to lignin as the catalytic domain of EG II was observed to adsorb more efficiently on lignin than the catalytic domain of CBH I (Palonen et al., 2004). This phenomenon was proposed by the authors to be due to a more open active site in endoglucanases as compared to the tunnel-shaped active site of cellobiohydrolases, which results 13  in increased hydrophobic interactions with lignin. These and other studies suggest that cellulase adsorption and hydrolysis can be further influenced by the structure of the catalytic domain. In contrast to the CBM and catalytic domain, the linker region has generally been regarded as having little function beyond domain connection and the prevention of proteolysis. However, in a recent study by Payne et al. (2013), the adsorption to bacterial cellulose of T. reesei Cel7A CBM, and CBM with an attached linker, was measured. The authors found that the CBM-linker protein enhanced the binding affinity of the CBM, suggesting that the linker may aid in enzyme adsorption during enzymatic hydrolysis. 1.2.1.2. Influence of synergistic enzyme interactions on enzyme adsorption As previously mentioned, the hydrolysis of cellulose requires the synergistic actions of a combination of cellulase enzymes where endoglucanases randomly cleave internal amorphous regions of cellulose to generate new chain ends for subsequent production of cellobiose by cellobiohydrolases and glucose by β-glucosidases (Lynd et al., 2002). As the initial step of hydrolysis occurs on an insoluble substrate, the synergistic interactions between enzymes also affect enzyme-substrate interactions.  Previous studies have demonstrated that adsorption of one cellulase enzyme can be affected in the presence of another cellulase enzyme (Maurer, Bedbrook and Radke, 2012; Igarashi et al., 2011; Eriksson, Karlsson and Tjerneld, 2002; Medve et al., 1998; Medve, Stahlberg and Tjerneld, 1994; Ryu, Kim and Mandels, 1984). For example, the adsorption of Cel7A and Cel6A on Avicel was quantified individually and in an equimolar mixture by Medve et al. (1998). These authors reported that the adsorption of Cel7A was reduced when Cel6A was added, likely due to competition for common binding sites by both enzymes. However, other 14  workers have proposed an alternative model, namely the cellulose “surface erosion model”, to explain the phenomenon (Igarashi et al., 2011; Eriksson, Karlsson and Tjerneld, 2002). In this model, it was suggested that the processive movement of Cel7A can be potentially blocked by obstacles along a cellulose chain. This blockage can be removed by the addition of other cellulase monocomponents, such as Cel6A, resulting in the release of Cel7A into solution. Unfortunately, due to the many variables involved in the experimental design, including the use of different substrates, enzyme dosages, enzyme combinations and a lack of an accurate quantification method to monitor enzyme adsorption, comparisons across different studies has remained difficult and it remains unclear which model best describes the interactions between cellulase enzymes during hydrolysis. However, it should be noted that both models support the mechanism that enzyme adsorption is affected by synergistic interactions between cellulase enzymes.  As commercial cellulase preparations continue to become increasingly complex with the addition of accessory enzymes, increased synergistic interactions between cellulases and accessory enzymes can also be anticipated (Hu et al., 2014; Hu et al., 2013; Hu, Arantes and Saddler, 2011; Bura, Chandra and Saddler, 2009; Ohgren et al., 2007). Since a major function of accessory enzymes is believed to be aiding in increasing the accessibility of cellulase enzymes to the cellulose, an increase in cellulase adsorption might be anticipated with the addition of accessory enzymes. However, a recent study showed that the addition of xylanase resulted in the release of Cel7A enzymes during the hydrolysis of steam pretreated corn stover (Hu et al., 2013). Thus, it is apparent that the role of accessory enzymes in terms of influencing enzyme adsorption to the substrate is complex.   15  1.2.2. Substrate factors In addition to enzyme factors, the substrate properties of lignocellulosic substrates also play an important role in influencing the adsorption of cellulase enzymes and subsequent enzymatic hydrolysis (Van Dyk and Pletschke, 2012; Mansfield, Mooney and Saddler, 1999). Lignocellulosic biomass is a complex material composed of three different primary components, cellulose, hemicellulose and lignin (Figure 2) (Sjöström, 1993). Depending on the source of the biomass and type of pretreatment employed, the composition, distribution and concentration of these components can vary significantly (Bura, Chandra and Saddler, 2009; Chandra et al., 2007; Ewanick, Bura and Saddler, 2007; Galbe and Zacchi, 2007; Martinez et al., 2005; Bura et al., 2003; Hammel et al., 2002). All of these substrate properties are known to strongly influence enzyme-substrate interactions.           Figure 2. The complexity of lignocellulosic biomass (Rubin, 2008) 16  1.2.2.1. Influence of lignin on enzyme adsorption Lignin, which is the second most abundant biopolymer after cellulose, is composed of aromatic polymers derived from three phenyl propane precursors, guaiacyl (G), syringyl (S), and p-hydroxyphenyl (H) units (Boerjan, Ralph and Baucher, 2003). Depending on the type and source of the lignocellulosic biomass, the composition and type of lignin can vary greatly. In general, softwoods have the highest lignin content (25-30%) followed by hardwoods (20-25%) and herbaceous biomass (10-25%) (Fan, Lee and Gharpuray, 1982). The type of lignin found can also vary as G units are predominantly found in softwoods. In contrast, G and S units are present in hardwoods whereas all three phenyl propane units are found in herbaceous biomass (Boerjan, Ralph and Baucher, 2003).  In plants, the primary functions of lignin are to impart structural integrity to the cell wall, to enable the transport of water and solutes through the vascular system and to protect against chemical and biological degradation (Boerjan, Ralph and Baucher, 2003). Although lignin is advantageous for the growth and development of plants, it is generally recognized that the residual lignin after pretreatment limits cellulose accessibility and unproductively binds cellulases (Kumar et al., 2012; Del Rio, Chandra and Saddler, 2011; Nakagame, Chandra and Saddler, 2010; Berlin et al., 2006; Mooney et al., 1998; Sewalt, Glasser and Beauchemin, 1997; Eriksson et al., 1991). Previous studies have demonstrated that lignin may act as a physical barrier that restricts the cellulase enzyme access/adsorption to the cellulose component (Arantes and Saddler, 2011; Mooney et al., 1998). This is in part due to the restriction of fibre swelling by lignin, which limits the penetration of cellulase to the individual cellulose fibres (Arantes and Saddler, 2011; 17  Mooney et al., 1998). In particular, it is recognized that softwoods are more recalcitrant than hardwoods and agricultural residues due to the higher amount of lignin and cross-linking (Nakagame, Chandra and Saddler, 2010). Additionally, the depolymerization and repolymerization of lignin during pretreatment may also result in the deposition of lignin droplets on the cellulose surface, which can further restrict the adsorption of cellulases by reducing the number of binding sites on cellulose (Donohoe et al., 2008; Selig et al., 2007). In a previous study by Kumar et al. (2012), the effects of delignification on steam-pretreated Douglas-fir were investigated. It was observed that the delignification of the biomass resulted in increased cellulose accessibility and improved hydrolysis yields, highlighting the role of lignin in limiting enzymatic hydrolysis by acting as a physical barrier that restricts enzyme adsorption to cellulose. Besides restricting the accessibility to cellulose, lignin can further affect hydrolysis efficiency by unproductively binding cellulase enzymes. Cellulase-lignin interactions are highly dependent on the type of pretreatment/post-treatment used as the characteristics of the residual lignin can vary significantly between different treatments (Del Rio, Chandra and Saddler, 2011; Berlin et al., 2006; Sewalt, Glasser and Beauchemin, 1997). In general, hydrophobic interactions are recognized as the primary forces driving protein adsorption and it is believed to be one of the major factors governing enzyme-lignin interactions (Palonen et al., 2004; Haynes and Norde, 1994). Specifically, CBMs have been shown to play a significant role in the unproductive binding of lignin by cellulases as their adsorption towards cellulose is facilitated by hydrophobic interactions arising from conserved aromatic amino acids (Boraston et al., 2004; Palonen et al., 2004). Palonen et al. (2004) previously investigated the adsorption of native Trichoderma reesei CBH I and EG II and their catalytic domain alone to isolated softwood lignins. The authors 18  found that the catalytic domain of both enzymes adsorbed less to lignin compared to the intact enzyme, suggesting that the adsorption to lignin was primarily facilitated by the CBM. Recent work also reported that an increase in the hydrophobicity of the binding face of the CBM from T. reesei Cel7A resulted in an increased affinity for lignin (Rahikainen et al., 2013).  In addition to hydrophobic interactions, electrostatic and hydrogen bonding interactions have also been reported to influence cellulase-lignin interactions (Del Rio, Chandra and Saddler, 2011; Nakagame et al., 2011; Nakagame, Chandra and Saddler, 2010; Berlin et al., 2006; Sewalt, Glasser and Beauchemin, 1997). For example, the addition of sulfonic groups to Organosolv-pretreated lodgepole pine was demonstrated by Del Rio et al. (2011) to increase the amount of free enzymes in solution and the efficiency of enzymatic hydrolysis. The effect of sulfonation on adsorption was attributed by the authors to decreased cellulase–lignin interactions resulting from increased electrostatic repulsion. An increase in the phenolic group content of lignin was also observed to increase the unproductive binding of cellulases and decrease the sugar yields obtained from subsequent hydrolysis, indicating the role of hydrogen bonding interactions between cellulases and lignin (Sewalt, Glasser and Beauchemin, 1997). Therefore, in addition to its role in restricting cellulose accessibility, lignin can also negatively influence enzyme adsorption and enzymatic hydrolysis through unproductive binding mechanisms via hydrophobic, hydrogen bonding and electrostatic interactions.  1.2.2.2. Influence of hemicellulose on enzyme adsorption Hemicellulose varies depending on the biomass feedstock but is predominantly a heterogeneous, branched polymer composed of D-xylose, D-mannose, D-galactose, D-glucose, L-arabinose, L-rhamnose, 4-O-methyl-D-glucuronic acid and galacturonic acid (Kuhad, Singh 19  and Eriksson, 1997). Hemicelluloses are further characterized by a lower degree of polymerization (<200) in comparison to cellulose and can be found in an acetylated form (Kuhad, Singh and Eriksson, 1997). The type and source of the lignocellulosic biomass has a significant influence on the composition and type of hemicellulose present (Sjöström, 1993). For example, galactoglucomannan and arabinoxylans are the predominant hemicelluloses found in softwood, representing about 20% and 5-10% of the dry weight of the biomass while hardwood and herbaceous biomass contains 4-O-methylglucoronoxylans (15-30% of the dry weight of the biomass) and glucomannans (2-5% of the dry weight of the biomass) as their major hemicelluloses (Sjöström, 1993). Similar to lignin, hemicellulose has been proposed to act as a physical barrier that limits the accessibility of cellulase enzymes to cellulose, in part due to its presence on the outer surface of cellulose fibers and inter-fibrillar spaces (Meng and Ragauskas, 2014; Hu, Arantes and Saddler, 2011). Additionally, acetyl groups in xylan have also been shown to limit enzyme adsorption by increasing the diameter or changing the hydrophobicity of cellulose as poor enzymatic hydrolyzability on acetic acid treated pulps was observed (Pan, Gilkes and Saddler, 2006). Therefore, the removal of hemicellulose is thought to improve the ease of cellulose hydrolysis (Várnai, Siika-aho and Viikari, 2010; Bura, Chandra and Saddler, 2009; Jeoh et al., 2007). In earlier work by Jeoh et al. (2007), the effect of xylan removal on the hydrolysis and cellulose accessibility of dilute acid pretreated corn stover was investigated. The authors found that hydrolysis efficiency was improved with xylan removal, which enhanced the accessibility of cellulose to cellulases. Similarly, improvements in hydrolysis yields of pretreated lignocellulosic substrates have also been reported in the presence of xylanases (Hu et al., 2013; Hu, Arantes and 20  Saddler, 2011; Kumar and Wyman, 2009). These studies thus suggest that hemicellulose can restrict enzyme adsorption by acting as a physical barrier, which subsequently limits enzymatic hydrolysis. During pretreatment, hemicelluloses may also undergo degradation to form oligosaccharides, monosaccharides and sugar-degradation products, which have been shown to influence the hydrolysis and adsorption of cellulase enzymes (Kumar et al., 2013; Sannigrahi et al., 2011; Shi et al., 2011). For example, it was demonstrated that the addition of xylooligomers derived from birchwood xylan significantly reduced the adsorption of a commercial cellulase preparation, Spezyme CP, on Avicel (Shi et al., 2011). The reasons behind these observations are still unclear, but synthetically derived xylobiose and xylooligomers have been shown to strongly inhibit enzymatic hydrolysis of pure cellulosic and lignocellulosic substrates (Baumann, Borch and Westh, 2011; Qing and Wyman, 2011; Qing, Yang and Wyman, 2010). Other work has also reported that acid based pretreatments, which primarily remove hemicellulose, can lead to the production of pseudo-lignin or char (Kumar et al., 2013; Sannigrahi et al., 2011). Pseudo-lignin is composed of insoluble compounds resulting from carbohydrate degradation and characterized by spherical droplets that reside on the cellulose surface as visualized by scanning electron microscopy (SEM) (Sannigrahi et al., 2011). It has been proposed that the presence of pseudo-lignin can reduce the number of surface binding sites on cellulose for subsequent adsorption (Kumar et al., 2013; Sannigrahi et al., 2011). In related work, the addition of pure xylose derived psuedo-lignin was also shown to reduce the amount of free Cel7A in solution, suggesting that psuedo-lignin may also bind protein unproductively, reducing hydrolysis efficiency (Kumar et al., 2013; Hu, Jung and Ragauskas, 2012) 21  1.2.2.3. Influence of cellulose on enzyme adsorption Cellulose is a linear polysaccharide composed of D-glucopyranose units linked by β-1,4-glycosidic bonds and accounts for approximately 35 to 50% of the dry weight of lignocellulosic biomass (Lynd et al., 2002). It is produced from individual chains of glucose subunits that are rotated 180o relative to each other along the main axis, resulting in the formation of a flat ribbon that consists of repeating cellobiose units (Somerville, 2006). It has been reported that each cellulose chain has a degree of polymerization ranging from 500 to 14,000-15,000 (Brett, 2000). Cellulose microfibrils are then formed as 36 parallel linear glucan chains, aggregated through intra and intermolecular hydrogen bonding as well as van der Waals forces, to form crystalline structures (Taylor, 2008; Somerville, 2006). As the direct adsorption to cellulose is required for enzymatic hydrolysis, the different properties of cellulose, type of allomorph, crystallinity, degree of polymerization and accessibility, will all affect cellulase adsorption. Cellulose can exist in major allomorphs, cellulose I, II, III and IV (Ishikawa, Okano and Sugiyama, 1997). Among the different allomorphs, cellulose I is found natively in nature and can be subdivided into cellulose Iα and Iβ depending on the crystalline structure (Gao et al., 2013; Igarashi, Wada and Samejima, 2007; Igarashi et al., 2006). However, in higher plants, the dominant form of cellulose is found as cellulose Iβ. Igarashi et al. (2006) previously investigated the adsorption of Cel7A from Trichoderma viride on cellulose Iα and Iβ. The authors found that the maximum cellulase adsorption capacity on cellulose Iβ was 1.5x times higher than cellulose Iα. Other works that converted cellulose I to cellulose III also showed a decrease in enzyme adsorption with increased sugar production (Gao et al., 2013; Igarashi, Wada and Samejima, 2007). These results thus suggest that the type of cellulose allomorph can affect both enzyme adsorption and cellulose hydrolysis. 22  Another property of cellulose that may affect enzyme-substrate interactions is the degree of crystallinity (Kumar and Wyman, 2013; Hall et al., 2010; Jeoh et al., 2007). This is in part due to the tightness of the crystalline array found in cellulose microfibrils, which has been reported to be sufficient to prevent even the entry of small molecules, such as water (Arantes and Saddler, 2010; Lynd et al., 2002). Previous work by Jeoh et al. (2007) showed that the adsorption of Cel7A from Trichoderma reesei increased with decreasing crystallinity. Recent work by Hall et al. (2010) also observed that the amount of Cel7A adsorbed to Avicel decreases linearly at crystallinity values above 35-45% while a constant level of adsorption was observed at lower crystallinity values. However, other studies reported no correlation between crystallinity and cellulase adsorption (Kumar and Wyman, 2013). Similarly, mixed results have been reported in the literature when determining the influence of cellulase crystallinity on hydrolysis (Mansfield, Mooney and Saddler, 1999). This difference in observations between studies may be attributed to the use of different characterization techniques, which can have a significant impact on the quantification of crystallinity values (Park et al., 2010). Furthermore, physical treatments used to alter the crystallinity of a substrate may also influence other substrate properties, such as specific surface area (Chandra et al., 2007; Mansfield, Mooney and Saddler, 1999). As a result, it has remained difficult to assess the possible influence of crystallinity on enzyme adsorption and hydrolysis. In addition to the type of cellulose allomorph and crystallinity, the accessibility of cellulose may also play an influential role in enzyme adsorption and has been suggested to be the major contributor in determining the rate and extent of overall hydrolysis (Arantes and Saddler, 2010). Cellulose accessibility can be measured using a variety of methods, including Simon’s stain, water retention values, direct enzyme adsorption, and carbohydrate binding module (CBM) 23  adsorption (Gourlay, Arantes and Saddler, 2012; Chandra et al., 2008; Hong, Ye and Zhang, 2007; Grethlein, 1985). Previous work has reported that there is a linear correlation between the initial hydrolyzability of a pretreated substrate and its accessibility to a molecule with a nominal diameter of 51 Å, which is considered to be representative of the size of a typical cellulase enzyme (Grethlein, 1985). Similarly, Gourlay et al. (2012) demonstrated that the initial enzymatic hydrolysis rates of cotton fibres followed the same profile as the amount of accessible amorphous and crystalline cellulose, as determined by the adsorption of CBM2a and CBM44. Other work by Arantes and Saddler (2011) has also shown that there is a strong linear correlation between the minimum protein loading to achieve efficient hydrolysis of organosolv and steam pretreated biomass and the available surface area of cellulose as measured by Simons’ stain and the maximum of protein adsorbed. As direct physical contact of cellulase enzymes and cellulose must occur prior to enzymatic hydrolysis, the accessibility of cellulose is thus an important factor that influences both enzyme adsorption and hydrolysis efficiency. In contrast to the cellulose properties described above, there are a limited number of studies which have investigated the impact of cellulose chain length on adsorption and hydrolysis (Yang et al., 2011). Kaplan et al. (1970) reported that the adsorption of cellulase on cotton decreased after outdoor weathering due to a decrease in degree of polymerization (DP) and ring opening of cellulose. However, other cellulose properties that are known to influence enzyme adsorption were not investigated. Unfortunately, as DP can be closely associated with changes in crystallinity and accessible surface area, the effects of DP on enzyme adsorption and hydrolysis remain unclear (Chandra et al., 2007; Mansfield, Mooney and Saddler, 1999).  24  1.2.3. Physical factors During the enzymatic hydrolysis of lignocellulosic biomass, operating conditions are often maintained at a pH of 4.8-5.0 and temperature of 50oC, which are regarded as the optimal conditions for hydrolysis by Trichoderma reesei cellulases. It is anticipated that changes in these parameters may affect both enzyme activity and adsorption.  1.2.3.1. Influence of temperature on enzyme adsorption In general, adsorption studies are performed at 4oC to minimize hydrolysis of the substrate (Pareek, Gillgren and Jonsson, 2013; Medve et al., 1998). However, as hydrolysis temperatures of 50oC are often employed, the effect of temperature on enzyme adsorption has been subsequently studied on pure cellulosic substrates, purified lignin and lignocellulosic substrates in many studies (Shang et al., 2014; Pareek, Gillgren and Jonsson, 2013; Zheng et al., 2013; Tu, Pan and Saddler, 2009; Kim, Jang and Jeong, 1998; Kyriacou, Neufeld and MacKenzie, 1988). It has been typically reported that the adsorption of cellulases on lignin and lignocellulosic substrates increases with increasing temperature (Shang et al., 2014; Pareek, Gillgren and Jonsson, 2013; Zheng et al., 2013; Tu, Pan and Saddler, 2009). For example, when studying the adsorption of a commercial cellulase mixture, Spezyme CP, on aqueous ammonia treated corncobs, Shang et al. (2014) reported that temperatures between 4-37oC favored desorption whereas higher temperatures favored adsorption. This phenomenon was proposed by the authors to be in part due to the increased exposure of hydrophobic regions on the protein at higher temperatures, which resulted in an increased absorption capacity.    25  1.2.3.2. Influence of pH on enzyme adsorption In addition to temperature, the pH of hydrolysis is also typically maintained between 4.8 and 5.0 (Lan, Lou and Zhu, 2013). Under these pH values, most cellulase enzymes are positively charged whereas lignocellulosic fibres are negatively charged, which facilitates enzyme adsorption (Nakagame et al., 2011). In previous works, it has been observed that pH can affect the charges of both substrate surface and cellulases (Lan, Lou and Zhu, 2013; Nakagame et al., 2011). Therefore, an increase in pH would increase the negative charge on the surface of lignocellulosic substrate and cellulases, favouring the desorption of enzymes (Lan, Lou and Zhu, 2013; Rahikainen et al., 2013; Du et al., 2012). Based on these previous studies, it is apparent that physical factors can further influence enzyme adsorption and desorption. 1.3.  Relationship between enzyme adsorption and hydrolysis performance As mentioned above, the composition/source of enzymes, the structural properties of lignocellulosic biomass substrates and hydrolysis conditions can have a significant impact on the adsorption profiles of cellulase enzymes. It would thus be anticipated that the adsorption of cellulases will vary depending on the pretreatment and hydrolysis conditions employed.  Interestingly, a review of the literature suggests that a relatively large proportion of the added enzymes remain free in solution regardless of the substrates, pretreatment conditions or enzyme mixtures used (Table 2). For example, many studies reported that more than 50% of the added enzymes remain in solution for pure cellulosic substrates, such as Avicel and filter paper (Gao et al., 2013; Kristensen, Felby and Jorgensen, 2009; Kumar and Wyman, 2008; Tu, Chandra and Saddler, 2007; Lu et al., 2002). Similarly, approximately 65-72% of the added enzymes were observed to be free in solution across two independent studies using different 26  pretreated corn stover and enzyme preparations (Pribowo, Arantes and Saddler, 2012; Zhu, Sathitsuksanoh and Zhang, 2009). 27  Table 2. Initial distribution of cellulase enzymes across different model and lignocellulosic substrates and total conversion obtained after hydrolysis. Substrate Pretreatment Enzyme mixture Free Enzymes (%) Adsorbed Enzymes (%) Total conversion (%) Reference Avicel  Spezyme-CP GC-220 59-63 49-67 37-41 33-51 n.d.b (Kumar and Wyman, 2008) Avicel Douglas fir  SO2 catalyzed steam Celluclast 1.5 L Novozym 188 70 50 30 50 80 100 (Tu, Chandra and Saddler, 2007) Avicel Douglas fir  SO2 catalyzed steam Celluclast 1.5 L Novozym 188 60-80 40-70 20-40 30-60 80 100 (Lu et al., 2002) Avicel  CBH I CBH II EG I ~65 ~65 ~55 ~35 ~35 ~45 ~100 (Gao et al., 2013) Avicel Douglas fir Douglas fir  Kraft pulping RMPa Celluclast Novozym 188 40 20 90 60 80 10 100 75 20 (Boussaid and Saddler, 1999) Corn Stover Dilute acid Spezyme CP 72 28 81 (Zhu, Sathitsuksanoh and Zhang, 2009) Corn Stover SO2 catalyzed steam Accellerase 1000 65 35 75 (Pribowo, Arantes and Saddler, 2012) Corncob Aqueous Ammonia Spezyme-CP 44-64 36-56 91 (Shang et al., 2014) Filter Paper  Celluclast 1.5 L Novozym 188 60 40 60 (Kristensen, Felby and Jorgensen, 2009) Mixed hardwood Steam explosion T. reesei βG T. reesei EG 58 20 42 80 60 Slnitsyn et al. (1983) Mixed softwood Ethanol Celluclast Novozym 188 45 55 95 (Tu, Pan and Saddler, 2009) Wheat straw Dilute acid Dilute alkali T. reesei cellulase Novozym 188 ~40 ~40 ~60 ~60 ~60 ~100 (Qi et al., 2011)  aRefiner mechanical pulps; bnot determined. 28  In previous studies, it has been demonstrated that there is a strong correlation between enzyme adsorption and the hydrolytic performance of complete enzyme mixtures (Wang et al., 2012; Arantes and Saddler, 2011; Kristensen, Felby and Jorgensen, 2009; Klyosov, 1990). For example, in prior work by Klyosov (1990), it was demonstrated that the level of cellulose conversion is limited by the level or “tightness” of binding of cellulase enzymes to the substrate. Similarly, strong correlations have always been found between total cellulase adsorption and their overall hydrolytic performance across a range of cellulosic and lignocellulosic biomass (Wang et al., 2012; Arantes and Saddler, 2011; Kristensen, Felby and Jorgensen, 2009). Based on these previous studies, it may be possible that the enzymes that remain free in solution, which represents a significant amount of the added enzymes, may only play a minor role in determining the overall rate and yield of the hydrolysis process.  However, other studies have also reported that increased enzyme adsorption is not necessary for efficient hydrolysis (Pakarinen et al., 2014; Gao et al., 2013; Varnai, Siika-aho and Viikari, 2013). It was shown that those cellulase enzymes that have less binding affinity (removing their CBMs) did not influence their hydrolytic potentials during the hydrolysis of Avicel and hydrothermally pretreated wheat straw at high solid loadings (Pakarinen et al., 2014; Varnai, Siika-aho and Viikari, 2013). Similarly, other work reported that the reduced enzyme binding affinity towards cellulose III as compared to cellulose I and regenerated amorphous cellulose had no significant impact on saccharification yields (Gao et al., 2013).   Due to the conflicting observations between different studies in the literature, an accurate assessment of the roles and functions of adsorbed enzymes during hydrolysis continues to be challenging. As mentioned above, enzyme, substrate and hydrolysis condition factors can play a significant role in affecting the adsorption of cellulase enzymes. However, many studies have 29  conducted hydrolysis using individual purified enzymes or “older” (such as Celluclast) cellulase preparations on pure cellulosic substrates or cellulose allomorphs that are not characteristic of real lignocellulosic substrates (Gao et al., 2013; Varnai, Siika-aho and Viikari, 2013; Wang et al., 2012; Kristensen, Felby and Jorgensen, 2009; Klyosov, 1990). Furthermore, previous hydrolysis studies studying the correlations between hydrolysis rate or yield and the quantity of adsorbed enzymes were often performed in the presence of both adsorbed and free enzymes (Pakarinen et al., 2014; Gao et al., 2013; Varnai, Siika-aho and Viikari, 2013; Wang et al., 2012; Arantes and Saddler, 2011; Kristensen, Felby and Jorgensen, 2009; Klyosov, 1990). This is in part due to the lack of a hydrolysis strategy to selectively remove the free enzymes during hydrolysis to allow for subsequent hydrolysis with mostly adsorbed enzymes. It is worth noting that a recent study by Yu et al. (2013) has reported that the removal of a large amount of free enzymes by vacuum filtration only had a minimal effect on the hydrolysis of Avicel and hardwood pulps. However, as substrate factors can further influence enzyme adsorption and hydrolysis, additional work is still required to better understand the roles and functions of the adsorbed and free enzymes to the overall hydrolysis of “real-life” pretreated lignocellulosic biomass.  1.4. Challenges of accurate quantification of enzyme adsorption and performance In order to better assess the roles and functions of adsorbed and free enzymes during lignocellulose deconstruction, it is apparent that protein distribution and enzyme performance must first be accurately quantified. However, the accurate determination of total protein concentration has remained challenging (Adney et al., 2012; McMillan et al., 2011; Adney et al., 1996). Many protein assays are influenced by the enzymes and compounds added or generated during lignocellulosic biomass pretreatment and hydrolysis. As mentioned above, current commercial enzyme mixtures often contain both cellulases and additional accessory enzymes to 30  improve hydrolytic performance and increase sugar yields (Gupta and Lee, 2013). For example, the most recent commercial cellulase preparation from Novozymes, Cellic CTec3, contains a significant number of different enzymes, which have unique amino acid compositions and physical properties (Figure 3). Additionally, during the pretreatment and subsequent hydrolysis of lignocellulosic biomass, a variety of compounds, such as monomeric and oligomeric sugars, and phenolic compounds are also released, potentially interfering with protein quantification (Kim et al., 2011; Ximenes et al., 2011; Ximenes et al., 2010). To further complicate matters, residual sugars from the fermentation broth as well as protein stabilizers and surfactants used to increase enzyme stability and to prevent unproductive binding to lignin, may also be present and interfere with protein determination (Adney et al., 2012; Chylenski et al., 2012; Eriksson, Karlsson and Tjerneld, 2002; Nieves et al., 1997). Therefore, a robust protein quantification assay that is compatible with these compounds and demonstrates minimal protein to protein variation is needed.       Figure 3. 2D gel electrophoresis of a commercial cellulase preparation, Cellic CTec3, from Novozymes.  14.4 21.5 31 45 66 97.5 116 kDa pH 3 10 31  Due to the complexity of lignocellulosic biomass derived samples, various assays have been used in the past by different researchers to monitor enzyme adsorption and performance during hydrolysis (Adney et al., 2012; McMillan et al., 2011; Adney et al., 1996). Unfortunately, the use of different techniques is highly problematic as significant challenges are often encountered when comparisons of enzyme activities and kinetics across different studies are required. In the following sections, the various methods of quantifying protein adsorption and performance are reviewed and assessed for their compatibility with lignocellulosic biomass hydrolysis derived samples.  1.5. Assays for quantifying protein adsorption 1.5.1. Gel-based assays  Gel-based assays are electrophoretic techniques that separate protein mixtures by molecular weight (sodium dodecyl sulfate polyacrylamide gel electrophoresis or SDS-PAGE), isoelectric point (pI) (isoelectric focusing or IEF) or a combination of both (2D-gel electrophoresis) (Vinzant et al., 2001). Following separation, the separated proteins can be visualized using different protein stains, such as Coomassie Brilliant Blue R-250. Based on the unique physical characteristics of each protein, the concentration of individual proteins can be identified and quantified semi-quantitatively by measuring the density of each protein band (Yu et al., 2013; Pribowo, Arantes and Saddler, 2012; Hu et al., 2010). Although these assays are relatively easy to perform, they are often unsuitable for complex cellulase mixtures, in part due to the similarities in molecular weight and isoelectric point between different enzyme components, which leads to the inability to resolve and quantify individual proteins (Yu et al., 2013; Pribowo, Arantes and Saddler, 2012; Hu et al., 2010).  32   An alternative gel-based assay for quantifying protein adsorption is the use of zymograms, which have been previously employed to measure the adsorption profiles of various cellulase components during hydrolysis (Pribowo, Arantes and Saddler, 2012). Similar to SDS-PAGE, IEF and 2D-gel electrophoresis, proteins are first separated by molecular weight and/or pI under non-denaturing conditions to preserve enzyme activity (Sun et al., 2008; Schwarz et al., 1987; Béguin, 1983). This is followed by the incubation of the separated proteins onto another gel containing different enzyme specific substrates. Enzymatic activity/adsorption can then be measured by the addition of a stain, such as Congo red, which displays clear bands against a dark background resulting from the hydrolysis of the added substrate (Sun et al., 2008; Schwarz et al., 1987; Béguin, 1983). This combination of gel electrophoresis and activity measurements provided by zymograms results in a semi-quantitative measurement of the individual adsorption profiles of different enzymes. However, the more routine use of zymograms is significantly restricted by the lack of enzyme specific substrates as individual cellulase enzymes have been shown to demonstrate activity towards multiple substrates (Zhang, Himmel and Mielenz, 2006). For example, both Cel7A and Cel7B are known to display activity towards 4-methylumbelliferyl-β-D-lactoside (Zhang, Himmel and Mielenz, 2006; van Tilbeurgh, Claeyssens and de Bruyne, 1982). As a result, individual enzyme adsorption profiles cannot be accurately quantified. Unfortunately, this limitation is expected to be further exacerbated as enzyme mixtures become increasingly complex to improve hydrolytic performance and increase sugar yields (Gupta and Lee, 2013). Therefore, zymograms are also unsuitable for the quantification of enzyme adsorption during the hydrolysis of lignocellulosic biomass.    33  1.5.2. Immunoassays  Another potential method of monitoring enzyme adsorption is the use of immunoassays, which include Western blots and enzyme-linked immunosorbent assays (ELISA) (Pribowo, Arantes and Saddler, 2012; Heiss-Blanquet et al., 2011; Dan et al., 2000; Aho et al., 1991; Riske, Eveleigh and Macmillan, 1990). Similar to gel-based assays, Western blots also require the prior separation of protein by gel electrophoresis. This is followed by the detection of a target protein by monoclonal antibodies (mAb), which only binds to a specific region of the protein of interest. Therefore, this method can be used to recognize and quantify individual enzyme components during hydrolysis and has been successfully employed for the quantification of Cel6A, Cel7A and Cel6B in a Trichoderma reesei culture (Aho et al., 1991). However, due to potential gel-to-gel variations, western blots are typically used as a semi-quantitative technique.   In contrast, an ELISA assay uses reporter enzyme linked antibodies to detect target proteins and can be performed directly in liquid samples (Pribowo, Arantes and Saddler, 2012; Heiss-Blanquet et al., 2011; Riske, Eveleigh and Macmillan, 1990). Upon successful binding of the antibodies, the quantity of a target protein can be subsequently detected by the addition of a substrate specific for the reporter enzyme and measuring the quantity of product produced, which provides a quantitative measurement of protein concentration. Similar to western blots, ELISA’s have been used by different researchers to quantify various cellulase monocomponents (Pribowo, Arantes and Saddler, 2012; Heiss-Blanquet et al., 2011; Riske, Eveleigh and Macmillan, 1990). In recent work by Pribowo et al. (2012), a double-sandwich ELISA was developed to track the distribution of cellulase monocomponents, Cel6A, Cel7A and Cel7B, during the hydrolysis of pretreated corn stover with two different commercial cellulase preparations.  34   Due to the high degree of specificity of antibodies, immunoassays offer a promising approach to monitor the adsorption profiles of individual enzymes within a complex mixture during hydrolysis. However, the high degree of specificity also leads to difficulties when quantifying enzyme cocktails from varying sources. This was demonstrated as antibodies developed for the detection of Cel6A in Celluclast (Novozymes, Franklinton, NC) displayed reduced binding affinity towards Cel6A present in Accellerase (Genencor) and Cellic CTec2 (Novozymes, Franklinton, NC) (Pribowo, Arantes and Saddler, 2012). These authors suggested that the difference in response may be due to a change in antigen recognition resulting from the use of a different Cel6A in the latter enzyme preparations. As a result, with the development of increasingly complex enzyme preparations, the development of a large repertoire of antibodies specific to the physical characteristics of each individual enzyme will be required. Unfortunately, this approach will likely be costly and time-consuming, limiting the applicability of immunoassays for the quantification of only a subset of enzyme monocomponents instead of a complete enzyme mixture. 1.5.3. Protein labelling assays Various authors have also studied the adsorption of cellulases with the use of protein labelling assays, which requires the direct attachment of a foreign molecule to a target protein/enzyme (Wang et al., 2012; Zhu et al., 2009; Hong, Ye and Zhang, 2007; Palonen, Tenkanen and Linder, 1999; Linder and Teeri, 1996; Chanzy, Henrissat and Vuong, 1984). For example, tritium labelling has been used to study the temperature dependency and reversibility of the binding of the CBM of Cel7A to bacterial microcrystalline cellulose (Linder and Teeri, 1996). Similarly, tritium labelled intact Cel7A and Cel5A as well as their respective catalytic 35  domains were also used to investigate the unspecific binding of cellulases to lignin (Palonen, Tenkanen and Linder, 1999). In addition to radionuclides, additional labelling strategies include the use of fluorescent proteins and metal particles. In previous work by Chanzy et al. (1984), Cel7A was labelled with 4-6nm gold particles, which subsequently enabled the direct visualization of its adsorption on the surface of cellulose microfibrils and microcrystals. Other studies using a recombinantly produced thioredoxin-green fluorescent protein-cellulase binding module fusion protein also allowed for the quantitative determination of the accessibility of cellulose present in pure and pretreated lignocellulosic substrates to cellulases (Wang et al., 2012; Zhu et al., 2009; Hong, Ye and Zhang, 2007). Due to the use of a foreign molecule to label and quantify target proteins/enzymes, protein labelling assays are not anticipated to be affected by complex enzyme mixtures and lignocellulosic biomass hydrolysis conditions. This potential lack of interference thus suggests that protein labelling is an effective method of monitoring enzyme adsorption. Unfortunately, the method is often technically tedious to perform (Wang et al., 2012; Zhu et al., 2009; Hong, Ye and Zhang, 2007). This is in part due to the non-specific nature of the protein labels, which necessitates the need for an extensive purification step to isolate labelled and unlabelled proteins/enzymes prior to their application (Zhu et al., 2009; Hong, Ye and Zhang, 2007; Chanzy, Henrissat and Vuong, 1984). Additionally, the addition of labels to enzymes may affect enzymatic activity depending on the type of labelling method used (Palonen, Tenkanen and Linder, 1999; Chanzy, Henrissat and Vuong, 1984). It has been previously reported that 40% of the initial starting activity of Cel7A was lost when labelled with colloidal gold particles (Chanzy, Henrissat and Vuong, 1984). Therefore, due to the potential loss of activity and tedious nature of 36  the technique, protein labelling assays may not be an ideal method for quantifying enzyme adsorption during hydrolysis. 1.5.4. Activity dependent assays  The use of substrate dependent assays is another potential method of quantifying enzyme adsorption (Pribowo, Arantes and Saddler, 2012; Rad and Yazdanparast, 1998; Otter et al., 1989). In this method, enzyme adsorption is determined as the difference in enzymatic activity between the complete enzyme mixture and the adsorbed or free enzymes, which can be quantified by the release of sugars, chromogenic or fluorescent compounds or dyes or a reduction in viscosity (Zhang, Himmel and Mielenz, 2006). Depending on the specific substrate being employed, the activity/adsorption of an individual enzyme component, subset of enzymes in a cellulase mixture or the total cellulase mixture, can be measured.  Although substrate dependent assays have been applied in previous studies to quantify enzyme adsorption during the hydrolysis of lignocellulosic biomass, many disadvantages exist. Similar to zymograms, they suffer from the lack of specific substrates that can accurately quantify the activity of an individual enzyme component or the total cellulase mixture (Zhang, Himmel and Mielenz, 2006). Additionally, the presence of soluble compounds added or generated during pretreatment or hydrolysis are known to greatly affect enzyme activity. It has been reported that the presence of phenolic compounds can both inhibit and precipitate enzymes found in commercial cellulase preparations (Ximenes, Kim and Ladisch, 2013; Kim et al., 2011; Tejirian and Xu, 2011; Ximenes et al., 2011; Ximenes et al., 2010). Both monomeric and oligomeric sugars are also known to inhibit cellulase activity through end product inhibition (Zhang and Viikari, 2012; Xiao et al., 2004; Holtzapple et al., 1990). As a result, enzymatic 37  activity measurements may not accurately reflect the true adsorption profile of cellulase enzymes. 1.5.5. Total protein assays  Unlike many of the assays described above, total protein assays measure the total amount of protein present instead of the quantity/activity of an individual protein. Enzyme adsorption is measured based upon the mass difference of the initial amount of enzyme added and free/adsorbed enzyme quantified. In general, total protein assays can be divided into five different groups based on their respective detection mechanisms. These include copper-based, dye-based, direct spectrophotometric-based, nitrogen elemental analysis-based and amino acid-based assays.   Copper-based assays are based upon the reduction of Cu2+ to Cu+ by peptides and include the Bicinchoninic acid (BCA) assay and Lowry assay (Smith et al., 1985; Lowry et al., 1951). The resulting Cu+ is then reacted with Bicinchoninic acid or Folin-phenol reagent to form a coloured complex that can be measured spectrophotometrically at 560nm or 750nm, respectively (Smith et al., 1985; Lowry et al., 1951). Due to their relative simplicity, both methods have been previously employed for the quantification of cellulases (Jourdier et al., 2013; Varnai, Siika-aho and Viikari, 2013; Adney et al., 2012; McMillan et al., 2011). However, a number of studies have demonstrated that both methods suffer from the interference of phenolic compounds, protein glycosylation and reducing sugars, in part due to their abilities to reduce copper independently of peptides (Aguiar and Ferraz, 2007; Walker, 2002; Fountoulakis, Juranville and Manneberg, 1992; O'Sullivan and Mathison, 1970; Folin and Denis, 1912). Complex enzyme mixtures also pose a challenge for the BCA assay as bicinchoninic acid can be reduced directly 38  by cysteine, cystine, tyrosine and tryptophan, which results in higher colour yields than other amino acids, potentially overestimating protein concentrations (Wiechelman, Braun and Fitzpatrick, 1988). Similarly, the reduction of the Folin-phenol reagent can occur through direct interactions with the amino acid side chains of asparagine, cysteine, histidine, tyrosine and tryptophan (Legler et al., 1985; Chou and Goldstein, 1960). Due to these limitations, copper-based assays are not an ideal method for the quantification of protein under lignocellulosic biomass hydrolysis conditions.  In contrast to copper-based assays, dye-based assays, such as the Bradford and Pierce 660nm assays, detect protein through the addition of a dye that selectively binds to basic amino acids (Antharavally et al., 2009; Bradford, 1976). For example, the Bradford assay utilizes the Coomassie Brilliant Blue G250 dye, which reacts with the positively charged NH3+ group of arginine, histidine and lysine. Upon binding, the dye undergoes a shift in the adsorption maxima that can be quantified spectrophotometrically at 595nm. A significant advantage of dye-based assays is their robustness towards high concentrations of sugars, but several disadvantages limit their use under lignocellulosic biomass hydrolysis conditions (Fountoulakis, Juranville and Manneberg, 1992; Chauvet and Lamy, 1990; Mattoo, Ishaq and Saleemuddin, 1987). In particular, the dependence on amino acid composition, specifically basic amino acids, by dye-based assays may not represent the diversity of proteins/enzymes present in current commercial cellulase preparations (Adney et al., 2012; Kruger, 2002; Tal, Silberstein and Nusser, 1985). For example, in previous work by Adney et al. (2012), it was demonstrated that the absorbance responses of purified T. reesei Cel7A and Cel6A were 25 to 50 times lower than that of bovine serum albumin when performing the Pierce 660nm assay. Additionally, protein glycosylation 39  may sterically hinder the optimal binding of dyes to basic amino acids, resulting in the underestimation of protein concentration (Fountoulakis, Juranville and Manneberg, 1992).  Another type of total protein assay previously employed to quantify enzyme adsorption are direct spectrophotometric-based assays. This technique involves the measurement of protein at ultraviolet wavelengths of 205 or 280nm. Absorption at 280nm is typically used in favour of 205nm as many buffers and salts display strong absorbance at 205nm (Grimsley and Pace, 2001). At 280nm, total absorbance is dependent on the cysteine, tyrosine and tryptophan content in the protein sample (Aitken and Learmonth, 2002). Protein concentration can be subsequently calculated using the Beer-Lambert Law, A = amcl where A is absorbance, am is the molar extinction coefficient, c is the concentration of protein and l is the length of the path length in centimeters (Noble and Bailey, 2009). When performed under lignocellulosic biomass hydrolysis conditions, direct spectrometric-based assays demonstrate high tolerance towards most monosaccharides and protein stabilizers, such as sorbitol and glycerol, as they do not exhibit strong absorbance at ultraviolet wavelengths (Fan and Geveke, 2007; Grimsley and Pace, 2001; Barr and Chrisman, 1940). However, other pretreatment derived furan and phenolic derivatives, such as furfural, HMF and lignin, display strong absorbance at 280nm (Koch and Schmitt, 2013; Martinez et al., 2000; Turunen and Turunen, 1967). Another limitation of direct spectrometric-based assays is the inability to quantify complex enzyme mixtures. Due to the varying amino acid composition of different proteins, each individual protein possesses a unique molar extinction coefficient (Banka and Mishra, 2002). For example, a difference of 15% can be observed between the molar extinction coefficients of Cel7A (78800 M-1cm-1) and Cel6A (92000 M-1cm-1) (Kipper, Valjamae and Johansson, 2005). Due to this potential source of error, an absolute protein concentration cannot be obtained accurately for a protein mixture when 40  quantifying protein concentrations at 280nm. As a result, direct spectrometric-based assays are also unsuitable for the quantification of cellulase mixtures.   Quantitative measurements of cellulases have also been performed using nitrogen elemental analysis methods, such as Kjeldahl Nitrogen Analysis and Dumas Combustion (Gao et al., 2014; Haven and Jorgensen, 2013; Kumar and Wyman, 2008). Both techniques determine protein concentration by measuring the amount of nitrogen in a sample using either combustion in an oxygen rich atmosphere at 950 to 1050oC (Dumas Combustion) or concentrated sulfuric acid hydrolysis (Kjeldahl Nitrogen Analysis) (Owusu-Apenten and FOODnetBASE, 2002). Subsequently, the amount of protein is calculated by the multiplication of a nitrogen-to-protein conversion factor, which is dependent on the amino acid composition of a sample (Owusu-Apenten and FOODnetBASE, 2002). Since nitrogen elemental analysis methods are only specific for nitrogen containing compounds, it is highly compatible with lignocellulosic biomass hydrolysis conditions as the primary source of nitrogen will likely be from the addition of cellulase enzymes during enzymatic hydrolysis. Unfortunately, a quantitative quantification of protein adsorption is often difficult to achieve due to the lack of a single nitrogen-to-protein conversion factor that can represent the diversity of enzyme-substrate adsorption profiles across different hydrolysis conditions (Gao et al., 2014; Pribowo, Arantes and Saddler, 2012; Kumar and Wyman, 2008; Owusu-Apenten and FOODnetBASE, 2002). Inaccuracies may thus arise when attempting to quantify the total protein concentration of adsorbed or free cellulases with nitrogen elemental analysis methods.  In recent studies, the ninhydrin assay, which is an amino acid-based assay, has been employed as a method of quantifying the adsorption of cellulases (Haven and Jorgensen, 2013; Pribowo, Arantes and Saddler, 2012; Arantes and Saddler, 2011; Hu, Arantes and Saddler, 2011; 41  Zhu, Sathitsuksanoh and Zhang, 2009). During analysis, protein samples are first hydrolyzed at 100oC with 6M hydrochloric acid for 24 hours to degrade polypeptides into amino acids (Starcher, 2001). The resulting amino acids then react with triketohydrindene hydrate (ninhydrin) to form a colored complex known as Ruhemann’s purple that can be measured spectrophotometrically between 560 to 580nm (Friedman, 2004; Starcher, 2001). Due to its specific detection of amino acids, the ninhydrin assay thus demonstrates decreased susceptibility to protein-protein variation and interference to most compounds encountered during lignocellulosic biomass hydrolysis (Kang and Lubec, 2011; Zhu, Sathitsuksanoh and Zhang, 2009; Friedman, 2004; Starcher, 2001; Chauvet and Lamy, 1990; Marks, Buchsbaum and Swain, 1985). However, the interference from sugar degradation products remains problematic, limiting the applicability of the ninhydrin assay towards lignocellulosic biomass hydrolyzates (Zacharius and Porter, 1967; Schilling, Burchill and Clayton, 1963; Amber and Snider, 1932). The ideal protein quantification assay should be able to quantify complex enzyme mixtures in the presence of added as well as pretreatment and hydrolysis generated compounds. Unfortunately, most of the assays described in the literature appear to be incompatible with conditions commonly encountered during the processing of lignocellulosic biomass (Table 3). Although the ninhydrin assay appeared to be the most promising assay, due to its specificity for amino acids, the interfering effects from sugar degradation products remain problematic.  42  Table 3. Compatibility of different protein assays with lignocellulosic biomass hydrolysis conditions a: compatible; X: incompatible; DOC: depends on concentration of selected compound Type of assay Examples Compatible witha Complex enzyme mixtures Carbohydrates Phenolic Compounds Protein stabilizers Surfactants Gel-based SDS-PAGE; IEF 2D-gel electrophoresis; Zymograms X     Immunoassays Western blots; ELISA X     Protein labelling Fluorescent/Particle/ Radiolabeling X     Substrate dependent Complete/Individual enzyme assays X X X X X Total protein BCA Assay X X X DOC DOC LowryAssay X X X DOC DOC Bradford assay X  X DOC DOC UV X  X DOC DOC Kjeldahl analysis X     Dumas combustion X     Ninhydrin  X    43  1.6. Thesis objectives Although many studies have attempted to study the relationship between enzyme adsorption and hydrolytic performance of cellulase enzymes, an accurate assessment of the role that adsorbed enzymes play in achieving effective cellulose hydrolysis has continued to be challenging. This is in part due to the lack of both an accurate protein quantification method to monitor enzyme adsorption/desorption and a hydrolysis strategy that minimizes enzyme loading. Most studies have employed pure cellulosic substrates or cellulose allomorphs, which are not representative of industrially relevant lignocellulosic substrates. Therefore, a primary objective of this thesis was to improve our current understanding of the roles of adsorbed and free enzymes during the hydrolysis of lignocellulosic biomass. It was hoped that, by better understanding their respective roles we could develop new strategies to reduce enzyme loadings. The specific objectives of each chapter within the thesis are summarised below. First (Chapter 3.1), a modified and rapid ninhydrin assay was developed to more accurately quantify the enzymes/protein present in lignocellulosic biomass derived hydrolyzates and to better monitor the adsorption/desorption of cellulases during hydrolysis. The addition of sodium borohydride was assessed for its ability to eliminate the interference of sugars and its effect on protein measurement. Elevated hydrolysis temperatures above the traditional conditions of 100oC were also investigated as a potential method to reduce the hydrolysis time required for complete protein hydrolysis. Additionally, the applicability of the modified ninhydrin assay under lignocellulosic biomass hydrolysis conditions was assessed by comparing the concentration of cellulase preparations suspended in the water soluble fractions obtained from the steam pretreatment of agricultural residues, hardwoods and softwoods. 44  I then (Chapter 3.2) assessed the roles and functions of adsorbed enzymes in determining the rate and extent of hydrolysis of lignocellulosic biomass. A two-stage hydrolysis strategy was developed to separate the free and adsorbed enzymes after reaching adsorption equilibrium for subsequent hydrolysis. Hydrolysis was performed on both model and lignocellulosic substrates, including dissolving pulp, steam pretreated poplar and steam pretreated lodgepole pine using whole enzyme preparations as well as adsorbed cellulases. Differences in hydrolytic rate and yield as well as enzyme distribution on model and lignocellulosic substrates were then evaluated using the modified ninhydrin assay developed in Chapter 3.1 to determine the possible relationship between enzyme adsorption and hydrolytic performance.    45  2. Materials and methods 2.1. Preparation of pretreated lignocellulosic biomass Two lignocellulosic substrates, poplar and lodgepole pine, were pretreated using SO2-catalyzed steam pretreatment in a 2L StakeTech III steam gun (Stake Technologies, Norvall, ON, Canada). Prior to steam pretreatment, poplar and lodgepole pine wood chips (DW; dry weight of 300 g) were first impregnated with SO2 in a re-sealable plastic bag and allowed to react overnight. The plastic bags containing unadsorbed SO2 were then opened and allowed to vent for 60 minutes. Subsequently, 50g of the biomass were pretreated under near optimal conditions based on previous studies in the steam gun (Table 4) (Kumar et al., 2010; Bura et al., 2003). After pretreatment, the water insoluble fractions of the pretreated biomass were separated from the liquid fractions by vacuum filtration and stored at -20oC until further analysis. Dissolving pulp (DsP) was used as a model cellulosic substrate.   Table 4. Pretreatment conditions for various biomass substrates used in this study. Pretreatment Species Temperature (oC) Time (min) SO2 (%w/w) Abbreviation Steam pretreatment Poplar 190 5 3 SPP190 Poplar 200 5 3 SPP200 Lodgepole pine 200 5 4 LPP200  2.2. Compositional analysis of pretreated biomasses The chemical compositions of the water insoluble fractions of the steam pretreated materials were determined using the Technical Association of the Pulp and Paper Industry (TAPPI) standard method T222 om-88 as previously described by Bura et al. (2003). The 46  monomeric sugars in the acid hydrolyzate were subsequently quantified by high performance liquid chromatography (HPLC) (Dionex DX-3000, Sunnyvale, CA) using a CarboPac PA1 column and fucose as the internal standard as described elsewhere (Arantes and Saddler, 2011). Acid soluble lignin was also quantified at 205nm using a UV-Vis spectrometer (Varian Cary 50, Belrose, Australia) and acid insoluble lignin was quantified using gravimetric filtration using a medium coarseness Gooch type filtering crucible (Pyrex, Corning, NY). All compositional analyses were carried out in triplicate.  2.3. Cellulase enzymes  Commercial cellulase enzyme mixtures, Celluclast 1.5 L, Cellic CTec2 and Cellic CTec3 were provided by Novozymes North America Inc. and derived from genetically modified strains of Trichoderma reesei. The commercial hemicellulase mixtures used in this study were HTec (Novozymes, Franklinton, NC) and Multifect Xylanase (Genencor US Inc., Palo Alto, CA). The protein concentration of each enzyme mixture was quantified using a modified ninhydrin assay with bovine serum albumin (BSA) as a standard.  2.4. Enzymatic hydrolysis  The enzymatic hydrolysis experiments were conducted at a total volume of 20mL in 50mL round-bottom centrifuge tubes (Pyrex, Corning, NY) at a solids loading of 2% w/v and 50oC with rotational mixing at 150 rpm using a hybridization incubator (FinePCR COMBI D-24, Korea). Prior to the addition of enzymes, DsP, SPP190, SPP200 and LPP200 were diluted to 2% w/v solids with water and 1M sodium acetate buffer to attain a final concentration of 50mM (pH=4.8) followed by incubation at 50oC for 1 hour with rotational mixing. After incubation, varying amounts of Cellic CTec3 were added directly to the reaction mixture. Hydrolysis was 47  performed over a period of 72 hours with periodic sampling of 1mL volume at 1, 2, 5, 24, 48 and 72 hours. Each collected sample was centrifuged at 13000rpm at 4oC for 10 minutes to separate the solid and liquid fractions. The liquid fractions were heated at 100oC for 10 minutes and stored at 4oC for further analysis. Total sugar produced was quantified by high-performance liquid chromatography (HPLC) (Dionex DX-3000, Sunnyvale, CA) as described elsewhere (Arantes and Saddler, 2011). All experiments were performed in duplicate and repeated at least twice. 2.4.1.  Two-stage hydrolysis  To assess the role of adsorbed and free cellulase enzymes in determining the hydrolysis kinetics and yield of lignocellulosic biomass, a two-stage hydrolysis strategy was developed to separate the respective enzyme fractions for subsequent hydrolysis (Figure 4). For all two-stage hydrolysis experiments, the total reaction volume was increased to 25mL from 20mL to facilitate the removal of free enzymes. Initially, substrates were incubated with water and sodium acetate buffer (as described in section 2.4) followed by the addition of Cellic CTec3. After the addition of enzymes, hydrolysis with the complete enzyme mixture was allowed to proceed for 1 hour to reach adsorption equilibrium between the liquid and solid phases. The resulting hydrolyzates were subsequently centrifuged at 4oC at 5000rpm for 15 minutes to separate the liquid and solid fractions. Different volumes of the liquid fraction, representing 25, 50 and 75% of the free enzymes in solution were removed and replaced with an equivalent volume of 50mM sodium acetate buffer (pH=4.8) unless otherwise specified. The re-suspended solids were then hydrolyzed for a further 71 hours for a total of 72 hours with the adsorbed and residual free enzymes. Periodic sampling and sugar quantification were conducted as described in section 2.4. The quantity of protein removed in the liquid fractions was quantified using the modified 48  ninhydrin assay described in section 2.5. Samples containing less than 50μg/g protein were lyophilised and re-suspended in 50mM sodium acetate buffer (pH=4.8) prior to protein quantification.   Figure 4. Two-stage hydrolysis strategy to separate free and adsorbed enzymes after 1 hour of hydrolysis with the complete enzyme mixture of Cellic CTec3. All hydrolysis experiments were conducted at 2% w/v solids loading. 2.4.2.  Multi-stage hydrolysis  To further elucidate the role of adsorbed enzymes, a multi-stage hydrolysis strategy with complete free enzyme removal was developed to ensure that hydrolysis was conducted only with the adsorbed enzymes (Figure 5). As previously described in section 2.4, substrates were initially pre-incubated with water and buffer at 50oC for 60 minutes with rotational mixing. The substrate-buffer mixture was subsequently cooled to 4oC prior to the addition of Cellic CTec3. To limit the initial hydrolysis rate, the reaction mixtures were maintained at 4oC and hydrolysis 49  was allowed to proceed for 1 hour. The liquid and solids fractions were then separated by centrifugation at 5000rpm for 15 minutes at 4oC. After centrifugation, the liquid fractions were removed and the solid fractions were re-suspended with 50mM sodium acetate buffer (pH=4.8) equivalent to the volume of liquid removed. This liquid-solid separation process was repeated until the total quantity of protein contained in the removed liquid fractions contained less than 5% of the initial protein added as determined by the modified ninhydrin assay (described in section 2.5). The washed solids containing only adsorbed enzymes were then re-suspended with 50mM sodium acetate buffer (pH=4.8) to the initial starting hydrolysis volume and incubated at 50oC for 71 hours for a total of 72 hours. For comparison, hydrolysis experiments under the same conditions were also conducted with a complete enzyme mixture at the same enzyme loadings without liquid-solid separation. Sampling was conducted at 1 and 72 hours followed by sugar quantification as described in section 2.4. For the determination of the quantity of protein adsorbed on the insoluble substrate after 1 hour of hydrolysis and subsequent washings, a volume of buffer equivalent to the difference of the initial starting volume and volume of liquid removed was added to the collected solids fractions to facilitate mixing. The pH of the re-suspended mixtures was adjusted to pH=13 using 50% w/w NaOH (Fisher Scientific) and incubated at room temperature for 2 hours with rotational mixing at 150 rpm using a hybridization incubator to desorb the adsorbed protein on the insoluble residue (solid fraction). The alkali-treated samples were subsequently centrifuged, after which the liquid fraction was collected and 1M sodium acetate buffer was added to a final concentration of 50mM to attain a final pH of 4.8. The desorbed proteins were stored at 4oC and quantified using the modified ninhydrin assay. Samples containing less than 50μg/g protein were 50  lyophilised and re-suspended in 50mM sodium acetate buffer (pH=4.8) prior to protein quantification.   Figure 5. Multi-stage hydrolysis strategy to retain only adsorbed enzymes for an additional round of hydrolysis from a commercial enzyme preparation, Cellic CTec3, after 1 hour of initial hydrolysis with the complete enzyme mixture. All hydrolysis experiments were conducted at 2% w/v.    51  2.5. Development of a modified ninhydrin assay 2.5.1. Traditional acidic ninhydrin assay Total protein concentration was measured using the ninhydrin assay following acid hydrolysis with bovine serum albumin (BSA, Sigma) as a protein standard. 100μL of each protein sample (protein concentration between 0-800μg/g) was mixed with 50μL of water in a 0.5mL screw-cap microcentrifuge tube (Fisherbrand). This was followed by the addition of 300μL of 9M hydrochloric acid (HCl) to a final concentration of 6M and subsequently heated in a dry heating bath (MBI Lab Equipment, Kirkland, PQ) at 100oC for 24 hours. After cooling to room temperature, 100μL of the hydrolyzate was transferred into a 1.5mL microcentrifuge tube (Fisherbrand) and neutralized with 100μL of 5M sodium hydroxide (NaOH) (Fisher Scientific). Upon neutralization, 200μL of 2% ninhydrin reagent (Sigma) was added and heated at 100oC for 10 minutes. Samples were then cooled to room temperature prior to the addition of 500μL of 50% ethanol. Subsequently, 200μL of the colored solution was transferred to a 96 well microplate (Corning) and the absorbance was read at 560nm by a microplate reader (PerkinElmer VICTOR3, Woodbridge, ON). All experiments were performed in triplicate and repeated at least twice. 2.5.2. Evaluating possible glucose interference of the traditional ninhydrin assay  To evaluate the possible interfering influence of glucose on the traditional ninhydrin assay, which is the predominant sugar produced during enzymatic hydrolysis of lignocellulosic biomass, BSA (100, 800 and 1600μg protein/g) and glucose (0, 5, 10, 20 and 40g/kg) (Sigma) were dissolved in 50mM sodium acetate buffer (pH=4.8). 50μL of the protein solution and 50μL of the sugar solution were mixed together, resulting in a final glucose concentration of 2.5, 5, 10 52  and 20g/kg and BSA concentrations of 50, 400 and 800μg/g respectively. 50μL of water was then added to the protein-sugar mixture. Alternatively, 50mM sodium acetate buffer (pH=4.8) was added in place of the sugar solution for the 0g/kg glucose condition. All samples were then quantified using the traditional ninhydrin assay.  2.5.3. Elimination of glucose interference with sodium borohydride As sodium borohydride (NaBH4) had been previously used to reduce monomeric sugars to sugar alcohols, we investigated the potential use of NaBH4 to remove the interference of sugars on the traditional ninhydrin assay (Abdek-Akher, Hamilton and Smith, 1951). 100μL of a 20g/kg glucose solution dissolved in 50mM sodium acetate buffer (pH=4.8) was incubated with 50μL of NaBH4 solution prepared by dissolving NaBH4 pellets (Sigma-Aldrich) in water followed by the addition of 0.02% v/v Antifoam O-30 (Sigma-Aldrich). Although higher sugar concentrations (>20g/kg) can be anticipated during high dry matter hydrolysis, these conditions are also likely accompanied by the lack of free water and a subsequent increase in free protein. Therefore, this may necessitate the dilution of samples prior to the quantification of protein and thus, the total sugar concentration used in subsequent work was limited to a maximum of 20g/kg. Three different NaBH4 concentrations of 4, 6 and 13.3g/kg were prepared, representing ratios of 1:10, 1:6.67, 1:3 NaBH4:glucose w/w respectively. Samples were incubated at room temperature and the reaction was stopped at 5, 15, 30, 60 and 120 minutes by the addition of 300μL of 9M HCl (final concentration of 6M HCl). Following NaBH4 treatment the samples were assayed by the traditional ninhydrin assay. Once the required incubation time and concentration of NaBH4 needed to remove glucose interference was determined, the same conditions were then used to assess the possible influence of hemicellulose derived sugars 53  (arabinose, galactose, mannose and xylose), at concentrations of 20g/kg. As discussed in more detail, the required ratio of NaBH4: glucose and incubation time required to completely eliminate sugar interference was determined to be 1:3 NaBH4:glucose w/w and 60 minutes. These conditions were subsequently used to develop the modified ninhydrin assay. 2.5.4. Influence of sodium borohydride on protein measurements   To further investigate the possible influence of NaBH4 on the ninhydrin assay, BSA (0-1600μg/g) and glucose (40g/kg) were dissolved in 50mM sodium acetate buffer (pH=4.8). 50μL of the protein solution and 50μL of the sugar solution were mixed together, resulting in a final glucose concentration of 20g/kg and maximum BSA concentration of 800μg/g. This protein-sugar mixture was then subjected to NaBH4 treatment as described above. As a comparison without the addition of NaBH4, 50μL of the protein solution was mixed with 50μL of 50mM sodium acetate buffer followed by the addition of 50μL of water. All samples were then subjected to acid hydrolysis at 100oC for 24 hours followed by quantification with the traditional ninhydrin assay. Upon establishing the effects of NaBH4 on BSA, the protein concentrations of commercial cellulase preparations (Celluclast 1.5 L, Cellic CTec2, Cellic CTec3) and commercial hemicellulase preparations (HTec, Multifect Xylanase) were also compared with and without the addition of NaBH4. The concentrations of sugars present in the enzyme preparations were measured using high-performance liquid chromatography (HPLC) (Dionex DX-3000, Sunnyvale, CA) as described elsewhere (Arantes and Saddler, 2011). 2.5.5. Reduction of time required for complete protein hydrolysis  As previous work had shown that the time required for complete protein hydrolysis can be reduced by increasing the hydrolysis temperature, we also investigated if the total hydrolysis 54  time can be reduced by increasing the hydrolysis temperature from 100oC to 130oC (Fountoulakis and Lahm, 1998). Previous work has hydrolyzed protein at temperatures as high as 160oC to reduce the hydrolysis time, but these conditions often required specially-designed equipment (Chiou and Wang, 1989). A temperature of 130oC was thus selected as a compromise condition between hydrolysis time and the need for specialized equipment. BSA, Cellic CTec2 and HTec were incubated using the NaBH4 conditions described above and hydrolyzed with 9M HCl (final concentration of 6M HCl) at 100oC and 130oC in a dry heating bath (MBI Lab Equipment, Kirkland, PQ). Samples were hydrolyzed in triplicates for 0, 2, 4, 6, 12, 18, and 24 hours at 100oC for 0, 15, 30, 60, 120 and 240 minutes at 130oC. The extent of protein hydrolysis was quantified using the ninhydrin assay. Upon determining the required hydrolysis time to attain complete protein hydrolysis at 130oC, the protein concentrations of commercial cellulase preparations (Celluclast 1.5 L, Cellic CTec2, Cellic CTec3) and commercial hemicellulases preparations (HTec, Multifect Xylanase) were compared between the traditional hydrolysis conditions of 24 hours at 100oC and the hydrolysis conditions at 130oC. As discussed later in more detail, complete protein hydrolysis was attained with a reduced time of 2 hours at 130oC across all of the enzyme preparations. These conditions were subsequently used for the modified ninhydrin assay. 2.5.6. The development of a modified ninhydrin assay to quantify protein under lignocellulosic biomass hydrolysis conditions The modified ninhydrin method was developed to improve the overall accuracy, compatibility and speed of the assay. 100μL of protein containing samples (protein concentration between 0-800μg/g) were first incubated with 50μL of NaBH4 for 60 minutes at a ratio of 1:3 NaBH4:total sugar (w/w) in a 0.5mL screw-cap microcentrifuge tube with BSA as a protein 55  standard unless otherwise specified. This was followed by the addition of 300μL of 9M HCl (final concentration of 6M HCl) and subsequent heating in a dry heating bath at 130oC for 2 hours. After cooling to room temperature, 100μL of the hydrolyzate was transferred to a 1.5mL microcentrifuge tube and neutralized with 100μL of 5M NaOH. Upon neutralization, 200μL of 2% ninhydrin reagent was added and heated at 100oC for 10 minutes. Samples were then cooled to room temperature prior to the addition of 500μL of 50% ethanol. Subsequently, 200μL of the colored solution was transferred to a 96 well microplate and the absorbance was read at 560nm. All experiments were performed in triplicate and repeated at least twice. 2.5.7. Compatibility of the modified ninhydrin assay with compounds encountered during lignocellulosic biomass hydrolysis  In addition to monosaccharides, lignocellulosic hydrolyzates can also contain other compounds, such as oligosaccharides and phenolic compounds. To determine the influence of these compounds, a commercial cellulase preparation, Cellic CTec3, was quantified using the modified ninhydrin assay described above after dilution in 50mM sodium acetate buffer (pH=4.8) or the water soluble fractions (WSFs) obtained from the SO2 catalyzed steam pretreatment of corn stover (SPCS), poplar (SPP) and lodgepole pine (SPLPP). These conditions were selected to mimic “worse-case scenario” hydrolysis conditions where high concentrations of phenolic derivatives and sugars should be present. After dilution, the final total sugar concentration in each enzyme/WSF mixture was always below 20g/kg. Samples without NaBH4 treatment were also quantified to assess the degree of interference by the WSFs. As mentioned above, the water soluble fractions (WSFs) were derived from SO2 catalyzed steam pretreated corn stover (SPCS), poplar (SPP) and lodgepole pine (SPLPP), which had been pretreated under near optimal conditions that maximized the recovery of carbohydrates 56  and resulted in cellulose-rich substrates that were susceptible to enzymatic hydrolysis (Arantes and Saddler, 2011). Compositional analyses of the water soluble fractions were carried out in duplicate according to procedures from the National Renewable Energy Laboratory (NREL) (Sluiter et al., 2006). Additionally, the total amount of phenolics was measured in triplicate using the Lowry-Folin method with phloroglucinol as a standard (Zhang et al., 2006). 2.5.8. Elucidation of possible interfering effects of individual lignocellulosic components  To elucidate the individual interfering effects of different lignocellulosic components present in the WSFs, Cellic CTec3 was diluted in the different WSFs and quantified using the modified ninhydrin assay after three different treatments. NaBH4 treatment alone was used to isolate the effects of oligosaccharides as only monosaccharides and the reducing end subunit of oligomeric sugars were reduced (Ng and Zeikus, 1986; Abdek-Akher, Hamilton and Smith, 1951). Alternatively, samples were autoclaved at 121oC for 60 minutes with 4% w/w sulfuric acid following the NREL procedure, without subsequent NaBH4 treatment, to determine the effects of oligosaccharides without prior NaBH4 treatment (Sluiter et al., 2006). A combination of both treatments (acid hydrolysis with NaBH4 treatment) was used to determine the effects of lignin derived compounds from steam pretreatment. The pHs of the autoclaved samples were all readjusted to 4.8 using 50% w/w NaOH. All samples were then hydrolyzed and quantified according to the modified ninhydrin assay described above. 2.5.9. Comparison of Current Protein Assays and the Modified Ninhydrin Assay   In order to assess the compatibility of traditional protein assays with lignocellulosic hydrolyzates, Cellic CTec3 was measured in triplicate in the presence of 50mM acetate buffer or pretreatment WSFs as described above using the BCA, Bradford and Lowry assays (Thermo 57  Fisher Scientific Inc., Rockford, IL) with BSA as the protein standard. Acetone precipitation was conducted prior to protein quantification by the assays according to the manufacturer’s specifications to remove any interfering compounds present in the WSFs. The determined protein values were then compared against those obtained by the modified ninhydrin assay.   58  3. Results and discussion 3.1. Development of a NaBH4 coupled ninhydrin based assay for the quantification of protein/enzymes during the enzymatic hydrolysis of pretreated lignocellulosic biomass 3.1.1.  Background The cost-effective production of sugars from lignocellulosic biomass continues to be challenging for the bioconversion industry. This is primarily due to the high cost associated with large amounts of enzyme/proteins needed to achieve efficient saccharification of cellulose and hemicellulose to monomeric sugars (Klein-Marcuschamer et al., 2012; Stephen, Mabee and Saddler, 2012). Not surprisingly, a considerable amount of effort has been dedicated to decreasing enzyme production costs, increasing enzyme performance and developing enzyme recycling strategies. As a result, enzyme/protein concentration has become a key indicator in increasing numbers of technoeconomic analyses (Davis et al., 2013; Humbird et al., 2011). Therefore, in order to assess the efficacy of these approaches from both a fundamental enzyme kinetic and techno-economic perspective, an accurate method of assessing cellulase activity/performance is needed.  Traditionally, the filter paper activity (FPA) assay is often employed to evaluate cellulase activity/performance and this assay is currently the recommended method by the International Union of Pure and Applied Chemistry (IUPAC) (Ghose, 1987). The filter paper activity assay measures cellulase activity on a volumetric basis by determining the amount of cellulase enzymes required to solubilize 4% of a 50mg Whatman #1 filter paper strip within 60 minutes. Although widely employed, it is well known that, in addition to the composition of an enzyme preparation, substrate characteristics can also play a significant role in affecting enzymatic performance (Adney et al., 2012; Arantes and Saddler, 2011). As a result, the filter paper activity 59  assay is often not reflective of enzyme performance on pretreated lignocellulosic substrates. Due to this inherent limitation of the filter paper assay, many researchers have increasingly shifted away from enzymatic activity based assays in favour of total protein quantification assays as a method of assessing enzyme performance (Adney et al., 2012; McMillan et al., 2011; Dowe, 2009). Unfortunately, the selection of a total protein quantification assay for the accurate determination of total protein concentration in a “cellulase mixture” has remained challenging. This is in part due to the complexity of a cellulase mixture, which contain both cellulases and other accessory enzymes, such as hemicellulose-degrading enzymes (Gupta and Lee, 2013). Additionally, interferences can arise simply from the complexity of commercial cellulase preparations, which contain a combination of cellulase and accessory enzymes, each with unique amino acid sequences and physical properties, such as glycosylation (Adney et al., 2012). Previously, a comparative study by Adney et al. (1996) compared the protein concentration of multiple commercial cellulase preparations using the modified Lowry assay, Bradford assay, Bicinchoninic acid (BCA) assay, Kjeldahl Nitrogen Analysis and UV absorbance at 280nm. It was found that, depending on the protein quantification method performed, the total protein concentration varied by as much as 90%. Similar results were also obtained by McMillan et al. (2011) where the protein concentration of four cellulase preparations varied by 2.6 - 4.8 times when measured by the Bradford and BCA assay. This large variability in protein concentrations when utilizing different quantification methods is of significant concern, particularly when the evaluation of enzyme activities and kinetics is being considered. To further complicate the current challenge of total protein quantification, samples from the enzymatic hydrolysis of lignocellulosic biomass often contain monomeric and oligomeric 60  sugars in addition to protein (Zhang and Lynd, 2004). Unfortunately, several of the protein assays that have been used to quantify cellulase enzymes are known to be incompatible with sugars (Walker, 2002; Schilling, Burchill and Clayton, 1963). In most previous works, the interfering effects of sugars were overcome by diluting the samples below interfering concentrations as low sugar and high protein concentrations were often encountered. However, with the increasing hydrolytic efficiency present in newer cellulase preparations, high sugar and low protein concentrations are likely. As a result, the dilution of samples can no longer effectively eliminate the interfering effects of sugars, resulting in significant compatibility issues with many protein assays when quantifying total protein under lignocellulosic biomass hydrolysis conditions. Additionally, interfering effects are also anticipated from other potential compounds found during lignocellulosic biomass hydrolysis, such as phenolic compounds, protein stabilizers, sugar degradation products and surfactants (Chylenski et al., 2012; Aguiar and Ferraz, 2007; Walker, 2002; Martinez et al., 2000; Chauvet and Lamy, 1990; Turunen and Turunen, 1967; Schilling, Burchill and Clayton, 1963; Folin and Denis, 1912). For example, furfural, 5-hydroxymethylfurfural (HMF) and phenolic compounds can arise from the degradation of sugars and lignin during the pretreatment of lignocellulosic biomass (Ximenes, Kim and Ladisch, 2013; Kim et al., 2011; Tejirian and Xu, 2011; Ximenes et al., 2011; Ximenes et al., 2010). Further interference can also be encountered if surfactants or protein stabilizers have been added to increase enzyme stability or to prevent unproductive binding to lignin (Adney et al., 2012; Chylenski et al., 2012; Eriksson, Karlsson and Tjerneld, 2002; Nieves et al., 1997). Therefore, the identification of a compatible total protein quantification method that is 61  unaffected by the complexity of cellulase preparations and conditions encountered during lignocellulosic biomass processing would be of value. As mentioned earlier, although the ninhydrin assay appears to be a promising assay due to its specificity for protein and compatibility with most compounds from lignocellulosic samples, it also has several limitations (Kang and Lubec, 2011; Friedman, 2004; Starcher, 2001; Marks, Buchsbaum and Swain, 1985; D'Aniello et al., 1985). These include the incompatibility with sugars, long hydrolysis times to attain complete protein hydrolysis and potentially wide-ranging standard deviations (Haven and Jorgensen, 2013; Starcher, 2001; Schilling, Burchill and Clayton, 1963). Therefore, in the work described below, a modified, accurate and rapid ninhydrin assay was developed and used to quantify total protein in the presence of lignocellulosic biomass hydrolysis derived compounds. In summary, the interfering effects of sugars were eliminated by subjecting protein samples to a sodium borohydride treatment and a six-fold reduction in total assay time was also achieved by increasing hydrolysis temperatures from the traditional 100oC to 130oC. 3.1.2.  Evaluation of glucose interference with the ninhydrin assay Prior to protein quantification by the ninhydrin assay, the complete hydrolysis of protein to amino acids is necessary. This is traditionally accomplished by heating the protein samples at 100oC for 24 hours with 6M HCl (Starcher, 2001). As other work has also recommended the use of alkaline conditions where the proteins are hydrolyzed using a strong base, such as 13.5M NaOH, we first compared the relative merits of the acid or alkaline approach to protein breakdown (Haven and Jorgensen, 2013; Zhu, Sathitsuksanoh and Zhang, 2009). Although the 62  acidic and alkaline ninhydrin assays showed a similar linear protein quantification range of 0-800μg/g, the alkaline ninhydrin assay has been reported to be influenced by various biomass derived components (Haven and Jorgensen, 2013). Previous work which used the alkaline ninhydrin method reported that protein concentrations were underestimated by up to 15% in the presence of the pretreatment liquid obtained from hydrothermally pretreated wheat straw (Haven and Jorgensen, 2013). Another potential drawback of the alkaline method is the potential high losses of serine and threonine, which can constitute as much as 25% of the amino acid composition of cellulase enzymes produced by Trichoderma reesei (Banka and Mishra, 2002; Fountoulakis, Juranville and Manneberg, 1992). Therefore, acidic hydrolysis conditions were selected in preference to alkaline conditions. Typically, some residual sugars from the fermentation broth are present in cellulase preparations and sugars are also produced during the hydrolysis of lignocellulosic biomass (Zhang and Lynd, 2004; Nieves et al., 1997). As expected, the acidic ninhydrin assay appeared to be highly sensitive to the presence of glucose, which is the predominant sugar released by cellulases (Figure 6). This is in part due to the dehydration of hexose sugars during the traditional protein conditions of 100oC, 6M HCl for 24 hours, resulting in the generation of sugar degradation products, such as levulinic acid. In previous work, it had been shown that levulinic acid can react anomalously with the ninhydrin reagent by forming a cyclic intermediate with the five-carbon ring of ninhydrin, leading to the formation of a red condensation product that absorbs at 560nm (Schilling, Burchill and Clayton, 1963). We observed that an abnormal red colour instead of the expected chromophore, Ruhemann’s purple, was formed in the spiked samples. However, the severity of the interference appeared to be dependent on the protein concentration (Figure 6) as no significant interference (<5%) was observed until the glucose 63  concentration was increased beyond 10g/kg, when protein concentrations were above 400μg/g. In contrast, due to the lower absorbance responses at lower protein concentrations (50μg/g), significant overestimation (>40%) was observed, even at the lowest concentration of glucose tested (5g/kg). As enzyme loadings can be anticipated to be further reduced with improvements in enzyme efficiency, this suggest that, in order to employ the acidic ninhydrin assay as a method of quantifying protein, the interference of sugars must first be overcome.  Figure 6. Effect of increasing glucose concentration on the measured protein concentration of BSA by the traditional ninhydrin assay. 3.1.3.  Overcoming sugar interference Several techniques have been proposed as a way of eliminating the effects of interfering compounds on protein assays. In particular, the precipitation of protein by trichloroacetic acid or acetone has often been employed (Noble and Bailey, 2009). As mentioned earlier, with increasing improvements in enzyme efficiency, low protein concentrations are expected during 64  the hydrolysis of lignocellulosic biomass. Consequently, precipitative methods may lead to the loss of protein during the precipitation process, resulting in difficulties when accurate quantification of protein is necessary. Sodium borohydride (NaBH4) is a mild reducing agent used in organic chemistry for the reduction of aldehydes and ketones to alcohols (Periasamy and Thirumalaikumar, 2000). Prior work showed that it could reduce monomeric sugars to their respective alcohols (Abdek-Akher, Hamilton and Smith, 1951). As a result, the addition of NaBH4 was investigated as a potential method of eliminating sugar interference by preventing the formation of sugar degradation products during acid hydrolysis. To mimic the sugar concentrations that may be expected from the dilution of samples after high solids loading hydrolysis of lignocellulosic biomass, glucose (20g/kg) was incubated with different ratios of NaBH4 (w/w) for up to 120 minutes, hydrolyzed under acidic conditions followed by quantification by the traditional ninhydrin assay. Regardless of the concentration of NaBH4 added, it was apparent that the reduction of glucose occurred rapidly and no further changes were observed between 60 and 120 minutes (Figure 7a). At a ratio of 1:10 and 1:6.67 NaBH4:glucose (w/w), the interfering effects of glucose continued. However, as the ratio of NaBH4 to glucose was increased to 1:3 (w/w), no absorbance was observed at 560nm after 60 minutes, indicating the minimization of glucose interference on the ninhydrin assay. 65    Figure 7. (a) Influence of different NaBH4 treatment conditions on the absorbance response of ninhydrin in the presence of 20g/kg glucose (b) Interference of sugars with and without NaBH4 treatment at a ratio of 1:3 NaBH4:sugar (w/w) for 60 minutes.  66  To try to further assess the reactivity of NaBH4 with other sugars produced during enzymatic hydrolysis, hemicellulosic sugars were also incubated at a ratio of 1:3 NaBH4:sugar (w/w) for 60 minutes and quantified using the ninhydrin assay (Figure 7b). As expected, without the addition of NaBH4, galactose and mannose resulted in significant interference due to their decomposition to levulinic acid. In contrast, although levulinic acid was not produced during the degradation of pentose sugars, both arabinose and xylose appeared to interfere with the assay. This increase in absorbance was likely due to the development of a yellow colour resulting from the production of furfural during acid hydrolysis of the pentose sugars. However, with the addition of NaBH4, the interfering effects of hemicellulosic sugars were completely eliminated as demonstrated by the lack of absorbance at 560nm. Therefore, a ratio of 1:3 NaBH4:total sugar (w/w) and incubation time of 60 minutes was employed in all subsequent work. This removed the interfering effects of up 20g/kg sugars on the ninhydrin assay. Alternatively, at sugar concentrations above 20g/kg, the dilution of the samples appeared sufficient to minimize interference depending on the protein concentration present after dilution. 3.1.4.  Compatibility of sodium borohydride with protein Having established that the interference of up to 20g/kg of sugars could be minimized with the addition of NaBH4, its compatibility with protein was evaluated. BSA was quantified using the ninhydrin assay in the absence or presence of glucose. Both protein samples were hydrolyzed using traditional acidic hydrolysis conditions with the NaBH4 only applied to the glucose containing samples. The absorbance at 560nm was then plotted against the protein concentration of BSA, resulting in a linear protein standard curve (Figure 8). A comparison of the slope between samples without NaBH4 treatment (y=2.1132x, R2=0.9996) and with NaBH4 67  treatment (y=2.0579x, R2=0.9995) suggested that the addition of NaBH4 had no significant effect on the protein quantification.  Figure 8. Comparison of the protein concentration determined after BSA (20g/kg glucose) was diluted in 50mM sodium acetate buffer, using either the traditional ninhydrin assay or the traditional ninhydrin assay after NaBH4 treatment. To further evaluate the compatibility of NaBH4 with the ninhydrin assay, commercial enzyme preparations were also quantified with and without NaBH4 treatment. It was likely that they contained some soluble compounds, such as residual sugar from the fermentation broth and surfactants that act as protein stabilizers that would interfere with the assay (Nieves et al., 1997). It was apparent that Cellic CTec2 contained the high sugar concentrations of 265.5g sugar/kg enzyme preparation (Table 5).   68  Table 5. Concentration of monomeric sugars present in commercial enzyme preparations.  Concentration (g sugar/kg enzyme preparation) Enzyme Preparations Arabinose Galactose Glucose Xylose Mannose Celluclast 1.5 L b.d.la 0.3 5.8 0.1 0.9 Cellic CTec2 b.d.la 3.3 265.5 0.5 3.2 Cellic CTec3 b.d.la 2.2 17.3 0.5 2.6 HTec b.d.la 0.5 0.9 0.01 0.9 Multifect Xylanase b.d.la 0.9 5.9 0.1 1.0  a b.d.l: below detectable limit Since high sugar concentrations were detected in some of the enzyme preparations, it was initially anticipated that the protein measurements by the traditional ninhydrin assay may be overestimated due to the generation of sugar degradation products during the acid hydrolysis step. Interestingly, the addition of NaBH4 had no effect on the protein concentration of the quantified cellulase preparations (Figure 9). This lack of interference from sugars was likely due to the large dilution factors (>100x) required to dilute the enzyme preparations prior to quantification due to the high initial concentration of protein and sugars present. These results confirmed that NaBH4 treatment is not necessary when the dilution of a protein sample is sufficient to reduce sugar concentration while maintaining a high protein:sugar ratio. However, under typical conditions, the dilution of samples alone is unlikely to eliminate the interference of sugars as high protein:sugar ratios are unlikely to be typically encountered. Thus, a NaBH4 treatment prior to protein quantification by the ninhydrin assay might be necessary, as shown in more detail below. 69   Figure 9. Comparison of total protein concentrations of different commercial enzyme preparations quantified using the traditional ninhydrin assay with and without NaBH4 treatment. 3.1.5.  Reduction of hydrolysis time to achieve complete protein hydrolysis Although the addition of NaBH4 can greatly improve the compatibility of the ninhydrin assay when being used under typical lignocellulosic biomass hydrolysis conditions, long hydrolysis times continue to be required for the complete hydrolysis of protein to amino acids. Previous work has shown that the complete hydrolysis of chicken egg white lysozyme can be attained within 45 minutes at 160oC (Chiou and Wang, 1989). Thus, we next investigated whether the complete hydrolysis of cellulases could be attained within a shorter period of time. To evaluate the effects of elevated hydrolysis temperatures, BSA, a commercial cellulase preparation (Cellic CTec2), and a xylanase preparation (HTec) were hydrolyzed at 100oC and 130oC, which was selected as a compromise condition between hydrolysis time and the necessity for specialized equipment, such as specially designed hydrolysis tubes. Complete hydrolysis was defined as the lack of increase in absorbance at 560nm upon quantification by the ninhydrin 70  assay. Among all of the protein samples hydrolyzed, it was evident that no further hydrolysis occurs after 18 hours at 100oC. In contrast, only 2 hours was required at 130oC to reach the same level of hydrolysis, suggesting that the overall speed of the ninhydrin assay can be significantly increased by over 80% at elevated temperatures above 100oC (Figure 10a-c).    Figure 10. Comparison of the time required to achieve complete hydrolysis of different proteins/enzyme preparations at 100oC and 130oC. (a) bovine serum albumin (b) Cellic CTec2, a commercial cellulase preparation (c) HTec, a commercial xylanase preparation. Since different proteins hydrolyze at different rates depending on their amino acid composition, the applicability of the reduced hydrolysis time was assessed across different 71  enzyme mixtures (Fountoulakis and Lahm, 1998). The protein concentration of three different commercial cellulase and two hemicellulase preparations hydrolyzed for 24 hours at 100oC and 2 hours at 130oC were thus compared (Figure 11). It was apparent that both hydrolysis conditions resulted in identical protein concentrations across all five enzyme preparations. Additionally, the standard errors obtained at 130oC (<5%) were also similar to those obtained from the traditional hydrolysis conditions. These results suggested that the two major operational drawbacks of the ninhydrin assay, incompatibility with sugars and long hydrolysis times to attain complete protein hydrolysis, can be overcome by prior treatment with NaBH4 followed by acidic hydrolysis at 130oC.   Figure 11. The effect of different hydrolysis conditions on the total protein concentrations of different enzyme preparations. Enzyme preparations were subjected to NaBH4 treatment and hydrolyzed at 100oC for 24 hours or 130oC for 2 hours with 6M HCl before quantification by the ninhydrin assay.  72  3.1.6.  Compatibility with lignocellulosic biomass hydrolysis conditions  Although the complementary actions of NaBH4 and elevated hydrolysis temperature allow the ninhydrin assay to be conducted accurately and rapidly in the presence of sugars, the compatibility of the modified ninhydrin assay with pretreatment derived compounds after acidic hydrolysis remains unclear. As mentioned earlier, lignocellulosic biomass samples are often complex and may contain lignin, lignin-derived phenolic fragments and polysaccharides (monomeric and oligomeric) in addition to protein. To assess the applicability of the modified ninhydrin assay with pretreatment derived compounds, we next quantified the protein concentration of a commercial cellulase preparation, Cellic CTec3, in the presence of the water soluble fractions (WSFs) obtained from the steam pretreatment of corn stover, poplar and lodgepole pine (Table 6). These conditions were selected to mimic “worse-case scenario” hydrolysis conditions where high concentrations of phenolic derivatives and sugars might be encountered.  Table 6. Compositional analysis of WSFs from steam pretreated substrates.   Concentration (g/kg) WSF a  Ara Gal Glu Xyl Man Phe SPCS Monomeric 1.5 0.9 2.2 13.8 0 6.0 Oligomeric 0.7 0.2 0.5 1.9 0.7 SPP Monomeric 0.8 2.3 20.7 40.5 2.9 6.9 Oligomeric 0.2 0.7 5.5 6.6 1.2 SPLPP Monomeric 3.2 7.5 19.2 12.3 22.7 3.0 Oligomeric 0.5 2.2 4.6 1.4 7.8  a SPCS, steam pretreated corn stover; SPP, steam pretreated poplar; SPLPP, steam pretreated lodgepole pine; Ara, arabinan; Gal, galactan; Glu, glucan; Xyl, xylan; Man, mannan; Phe, phenolics. 73  Without the addition of NaBH4, it was apparent that the water soluble fractions from the three types of steam pretreated lignocellulosic biomass all suffered from interference (Figure 12). The protein concentration of Cellic CTec3 was overestimated by 5, 22 and 22% when dissolved in the WSFs derived from SPCS, SPP and SPLPP respectively. However, the NaBH4 treated samples all showed no interference. This was unexpected as it was initially anticipated that oligosaccharides would still be present after the NaBH4 treatment and would degrade to monosaccharides and sugar degradation products that might interfere with the ninhydrin assay during acid hydrolysis. This would be in part due to the reaction mechanism of NaBH4, which reduces monosaccharides and only the reducing end subunit of oligomeric sugars to their respective alcohol forms (Ng and Zeikus, 1986; Abdek-Akher, Hamilton and Smith, 1951). Consequently, the remaining subunits forming the oligosaccharide were anticipated to be capable of forming degradation products during the acidic hydrolysis process. To investigate if oligosaccharides in addition to monosaccharides could cause interference, the enzyme/WSF mixtures were autoclaved at 121oC with 4% w/w sulfuric acid to acid hydrolyze the oligosaccharides to monosaccharides. This was followed by quantification using the modified ninhydrin assay without prior NaBH4 treatment. It was apparent that the additional increase in monosaccharide concentration resulted in a further increase in the overestimation of protein concentration across all of the tested WSFs (Figure 12). This suggested that both monosaccharides and oligosaccharides can form sugar degradation products that react with ninhydrin. A possible mechanism for the reduced interference occurring with oligosaccharides upon reduction by NaBH4 is the decrease in the number of monomeric subunits capable of degradation during acidic hydrolysis. For example, previous work has shown that the acid hydrolysis of cellobiose previously reduced with NaBH4 would produce one unit of glucose 74  and sorbitol respectively (Ng and Zeikus, 1986). As Cellic CTec3 was quantified at a protein concentration of 200 - 400μg/g, the possible influence of the oligosaccharides was likely below effective levels. Although the interference of oligosaccharides may be significant at low protein concentrations, it was apparent that carrying out an initial acid hydrolysis with 4% w/w sulfuric acid at 121oC followed by NaBH4 treatment minimized any interference resulting from either mono- or oligosaccharides (Figure 12). However, under typical enzymatic biomass hydrolysis conditions, the concentration of oligosaccharides is likely to be relatively low due to the presence of β-glucosidase and other accessory enzymes in enzyme preparations to facilitate monosaccharide production (Gupta and Lee, 2013). As a result, NaBH4 treatment alone should be enough to overcome the interference of sugars. In contrast, the phenolic derivatives from steam pretreatment did not appear to affect the ninhydrin assay as the removal of sugar interference was sufficient to eliminate any interfering effects arising from the water soluble fractions (Figure 12). This is in agreement with previous work that demonstrated plant derived extractives and lignin did not interfere with the ninhydrin reagent (Chauvet and Lamy, 1990; D'Aniello et al., 1985). Therefore, it appears that the modified ninhydrin assay can be used as a method for the accurate and rapid quantification of total protein concentration up to 800μg/g in lignocellulosic biomass hydrolysis derived samples containing a maximum sugar concentration of 20g/kg. 75   Figure 12. Quantification of total protein concentration of Cellic CTec3 diluted in the WSFs from steam pretreatment by the modified ninhydrin assay. 3.1.7. Comparison of the modified ninhydrin assay versus current total protein quantification assays  We next quantified Cellic CTec3 in 50mM sodium acetate buffer or the WSFs obtained from steam pretreatment using either the BCA, Bradford or Lowry assays. These assays have been previously employed by various workers to quantify total protein in cellulase mixtures. However, they all suffer from various interfering effects, as described earlier. The measured protein concentrations were then compared against values obtained by the modified ninhydrin assay, which is highly specific for amino acids and compatible with lignocellulosic biomass hydrolysis derived compounds. As expected, in comparison to the modified ninhydrin assay, the copper-based and dye-based assays resulted in significantly higher and lower protein concentrations respectively, even in the presence of 50mM sodium acetate buffer (Figure 13). As discussed earlier, this difference in the measured protein concentrations was anticipated as both 76  types of assays have been shown to be influenced by amino acid composition, phenolic compounds and protein glycosylation.   Figure 13. Quantification of total protein concentration of Cellic CTec3 diluted in different WSFs obtained after steam pretreatment using various protein assays. It was apparent that all of the tested assays were significantly influenced by compounds present in the WSFs from steam pretreatment, even after an additional acetone precipitation step to remove interfering compounds (Figure 13). The protein concentration of Cellic CTec3 was underestimated up to 13% and 18% by the BCA and Lowry assay respectively and overestimated by up to 14% by the Bradford assay when diluted in WSFs compared to 50mM sodium acetate buffer. In contrast, the modified ninhydrin assay showed good agreement and reproducibility, in the presence of either 50mM sodium acetate buffer or WSFs. Additionally, standard errors of less than 5% were observed across all of the samples, when performing the modified ninhydrin assay, whereas acetone precipitated samples from the BCA, Bradford and Lowry assays showed large standard errors, likely due to incomplete protein recovery after precipitation. Thus, it 77  appears that the modified ninhydrin assay can be reproducibly used to quantify total protein/enzyme concentrations under typical lignocellulosic biomass hydrolysis conditions. 3.1.8. Conclusions Previous protein quantification methods used to quantify cellulases have all suffered from some sort of interference from other materials present during the enzymatic hydrolysis of lignocellulosic biomass. A modified and rapid ninhydrin assay that can accurately measure total protein concentration was developed. A combination of sodium borohydride treatment followed by acid hydrolysis at 130oC eliminated the interfering effects of sugars up to a concentration of 20g/kg and reduced the total assay time 6-fold. The method was also shown to be compatible across a range of different lignocellulosic biomass hydrolysis conditions, allowing for the accurate determination of enzyme kinetics and performance.   78  3.2. The relative roles that adsorbed and free enzymes play in determining the rate and extent of hydrolysis of lignocellulosic biomass and the potential of a two-stage hydrolysis strategy to reduce enzyme loadings 3.2.1.  Background Previous work (section 3.1) had shown that a modified ninhydrin assay consisting of the combination of sodium borohydride treatment followed by acid hydrolysis at 130oC was able to accurately and rapidly quantify protein under lignocellulosic biomass hydrolysis conditions. As mentioned previously, the interpreted relationship between cellulase adsorption and these enzymes hydrolytic performance was likely influenced by the lack of an accurate protein quantification method to monitor enzyme adsorption/desorption during hydrolysis (Adney et al., 2012; McMillan et al., 2011; Adney et al., 1996). Additionally, experimental conditions in many previous studies, where individual purified enzymes and/or simplified cellulase mixture were used on “pure” cellulosic substrates, may also not represent the complexity of current hydrolysis processes where multiple enzyme components (cellulases and various accessory enzymes) work together to break down complex, pretreated lignocellulosic substrates. Therefore, we next used the improved protein assay to try to better elucidate the various roles and functions of adsorbed enzymes (using Cellic CTec3) during the hydrolysis of various industrially relevant pretreated lignocellulosic substrates. As was described in detail earlier, enzymatic hydrolysis of cellulose is thought to occur in at least two major phases where the insoluble cellulose is first hydrolyzed to soluble cello-oligosaccharides followed by the rapid hydrolysis of soluble cello-oligosaccharides to cellobiose and glucose (Arantes and Saddler, 2011; Lynd et al., 2002). Due to the insoluble nature of lignocellulosic biomass, it is well known that the diffusion and adsorption of cellulases to the 79  solid substrate is a key prerequisite prior to the hydrolysis of lignocellulosic biomass. Interestingly, as mentioned earlier, a review of the literature suggests that a relatively large proportion of the added enzymes remain free in solution regardless of the substrates, pretreatment conditions or enzyme mixtures used while achieving high glucan conversion yields (Table 2). In previous works, it was reported that the removal of the free enzymes during hydrolysis only has a minimal effect on the hydrolysis of pure cellulosic substrates (Yu et al., 2013; Lee and Fan, 1983; Mandels, Kostick and Parizek, 1971). Based on these observations, the use of primarily adsorbed enzymes for hydrolysis may represent a significant opportunity to reduce the currently high enzyme loadings required for cellulose hydrolysis.  However, there are several contradictory observations and conclusions regarding the roles and functions that adsorbed enzymes play in determining the hydrolysis rate and yield of lignocellulosic biomass (Pakarinen et al., 2014; Gao et al., 2013; Varnai, Siika-Aho and Viikari, 2013; Arantes and Saddler, 2011; Wang et al., 2011; Kristensen, Felby and Jorgensen, 2009; Klyosov, 1990). This is in part due to the use of individual purified enzymes or older commercial cellulase preparations (such as Celluclast 1.5 L) on pure cellulosic substrates or cellulose allomorphs that are not characteristic of real lignocellulosic. As described previously, both substrate and enzyme related factors are known to influence enzyme adsorption. Previous work has demonstrated that interactions between different individual enzyme components and their respective physical structures can influence enzyme adsorption/desorption profiles. This is likely to be enhanced with the increasing complexity of current commercial cellulase preparations (Payne et al., 2013; Palonen et al., 2004; Linder et al., 1995; Reinikainen et al., 1992; Ståhlberg, Johansson and Pettersson, 1991; Tomme et al., 1988; Van Tilbeurgh et al., 1986). Additionally, other work that has investigated the influence of substrate properties on enzyme adsorption 80  indicated that lignin and hemicellulose may act as a physical barrier that restricts the access of cellulose to cellulases (Meng and Ragauskas, 2014; Kumar et al., 2012; Arantes and Saddler, 2011; Hu, Arantes and Saddler, 2011; Mooney et al., 1998; Eriksson et al., 1991). Related work has also showed that lignin may unproductively bind cellulase enzymes via electrostatic, hydrophobic and hydrogen bonding interactions (Rahikainen et al., 2013; Del Rio, Chandra and Saddler, 2011; Nakagame et al., 2011; Nakagame, Chandra and Saddler, 2010; Berlin et al., 2006; Palonen et al., 2004; Sewalt, Glasser and Beauchemin, 1997). Therefore, in the work reported here, the roles and functions of adsorbed enzymes from a current commercial enzyme mixture during the hydrolysis of pretreated lignocellulosic biomass were determined. A two-stage hydrolysis strategy was initially evaluated where a “model” cellulosic substrate (dissolving pulp) was hydrolyzed, and after a set time, the enzymes that were free in solution were removed, with only the adsorbed enzymes contributing to subsequent cellulose hydrolysis. In subsequent work, steam pretreated poplar and lodgepole pine were hydrolyzed with either the adsorbed enzymes or the entire enzyme mixture, using Cellic CTec3. Regardless of the substrate used, the selective removal of free enzymes had only a relatively limited effect on the hydrolysis potential of CTec3 preparations. By using a two-stage hydrolysis approach to remove the “free” enzymes, the required CTec3 enzyme dosage could be reduced by up to 53% while maintaining effective cellulose hydrolysis. 3.2.2.  Time course of enzymatic hydrolysis of DsP To minimize the possible influence of hemicellulose and lignin, the possible contributions of the adsorbed and free enzymes were initially assessed during the hydrolysis of dissolving pulp (DsP). The minimal enzyme loading required to achieve industrially relevant glucan conversion 81  yields (70-100%) was targeted and, as shown in Figure 14, an enzyme loading of 30mg Cellic CTec3/g glucan resulted in the hydrolysis of 94% of the pulp after 72 hours.   Figure 14. Time course of hydrolysis of DsP after the selective removal of different proportions of free enzymes after 1 hour of hydrolysis at a solids loading of 2% w/v, 50oC with an enzyme loading of 30mg Cellic CTec3/g glucan. DsP: dissolving pulp. 3.2.3.  Two-stage hydrolysis strategy To assess the relative contributions of adsorbed/free enzymes during hydrolysis, a two-stage hydrolysis strategy was developed that involved the selective separation of the adsorbed and free enzymes by centrifugation after 1 hour of hydrolysis. Previous work had shown that most cellulases reached adsorption equilibrium with the substrate after 30 to 60 minutes of incubation (Pribowo, Arantes and Saddler, 2012). Following initial hydrolysis and adsorption, different volumes of the liquid fraction, representing different amounts of free enzymes, were subsequently removed and the hydrolysis continued for a further 71 hours, for a total of 72 hours. The hydrolysis rates and yields were then compared to assess if there was any difference in 82  hydrolytic performance between the complete enzyme mixture and hydrolysis with just the adsorbed and residual free enzymes.  Previously, it was suggested that the re-suspension of enzyme-substrate pellets separated by centrifugation resulted in the irreversible binding of purified Cel7A, Cel6A and Cel7B onto the cellulosic substrates (Pellegrini et al., 2014). This was thought to be due to the compression of the substrate, leading to the delayed release of the adsorbed enzymes. Although it appeared there was irreversible adsorption of the individual cellulase enzymes, it was not known if this would also occur with complex commercial enzyme mixtures. Previous works have shown that the synergistic interaction between different celluloytic and non-celluloytic enzymes changed the adsorption and desorption profiles of the various enzymes (Maurer, Bedbrook and Radke, 2012; Igarashi et al., 2011; Eriksson, Karlsson and Tjerneld, 2002; Medve et al., 1998; Ryu, Kim and Mandels, 1984). Although other techniques, such as ultracentrifugation and vacuum filtration have been used to separate adsorbed and free enzymes, both techniques result in the loss of some of the insoluble substrate (Yu et al., 2013; Ramos and Saddler, 1994). Recent work by Yu et al. (2013) also showed the transfer of enzymes that were used to hydrolyze Avicel and hardwood pulps to the filter paper used during vacuum filtration. Therefore, due to the reduced loss of the insoluble substrate and protein, centrifugation was selected as probably the best method to try to fractionate the adsorbed and free enzymes. 3.2.4.  Selective removal of free enzymes during the hydrolysis of DsP As mentioned earlier, 94% of the original cellulose was hydrolyzed after 72 hours using an enzyme loading of 30mg/g glucan of the complete enzyme mixture. At this enzyme dosage, 70% of the added protein was located in solution after 1 hour of hydrolysis. As previous work by 83  Yu et al. (2013) reported that the removal of free enzymes had an insignificant effect on the hydrolysis of Avicel and hardwood pulps, we wanted to determine if the hydrolysis of DsP could be achieved using only adsorbed enzymes. Thus a two-stage hydrolysis strategy was used to selectively remove different quantities of free enzymes and to assess the possible role of adsorbed enzymes in determining the rate and extent of hydrolysis. It was apparent that the free enzymes only contributed marginally to the overall hydrolysis of DsP, as indicated by the similar rate and hydrolysis yields that were achieved (Figure 14). Cellulose hydrolysis yields of 93% and 88% could still be achieved when 25% and 50% of the free enzymes, which represented 18% and 37% of the initial protein added, were removed after one hour by centrifugation. When the amount of free enzymes/protein removed was further increased to 75%, more than 85% of the DsP was hydrolyzed after 72 hours, corresponding to only a 10% decrease in yield despite removing nearly 55% of the initial protein added. Interestingly, it appeared that hydrolysis yields decreased as the quantity of free enzymes removed was increased. Due to the large amount of enzyme removed during the two-stage hydrolysis strategy it was thought that this might be due the loss of β-glucosidase, which is primarily found in solution, resulting in the accumulation of cellobiose and product inhibition of the cellulase enzymes (Xiao et al., 2004). When Novozym 188, a commercial β-glucosidase preparation, was added after the removal of 75% of the free enzymes at a protein loading equivalent to 20% of the initial protein added, no significant improvement was observed. This suggested that sugar inhibition was not the main reason for the slightly lower hydrolysis yields observed with increasing removal of free enzymes. 84  Earlier work has shown that incomplete cellulose hydrolysis might occur, even when using very high protein loadings and prolonged hydrolysis times, due to the increasing recalcitrance of the residual cellulose within the substrate as hydrolysis proceeds (Mooney et al., 1998; Hogan and Mes-Hartree, 1990). Due to the near complete hydrolysis of DsP, it is possible that enzymes that were potentially essential for the hydrolysis of the remaining recalcitrant cellulose were partially removed during the two-stage hydrolysis. This was evident as lower conversion yields were only observed when 50 and 75% of the free enzymes in solution were removed. However, despite the slightly lower hydrolysis yields, the adsorbed enzymes were able to hydrolyze the DsP to almost the same extent as when using the complete mixture. Thus, nearly 55% less enzyme could be used to achieve almost the same level of hydrolysis. Based on these observations, it appeared that the enzymes initially adsorbed to the substrate are primarily responsible for the overall hydrolysis and represented a potential opportunity to lower enzyme loadings. 3.2.5.  Effect of the selective removal of free enzymes on the enzymatic hydrolysis of lignocellulosic biomass Although the removal of free enzymes/protein appeared to have little effect on the hydrolysis of DsP, it remained unclear if the same phenomenon extends to more complex and recalcitrant lignocellulosic substrates. Unlike DsP, lignocellulosic biomasses are naturally heterogeneous, containing hemicellulose and lignin in addition to cellulose. In previous work, it was suggested that the accessibility to cellulose by cellulases is one of the major factors that affects the ease of hydrolysis of lignocellulosic biomass (Gourlay, Arantes and Saddler, 2012; Arantes and Saddler, 2010; Mansfield, Mooney and Saddler, 1999; Grethlein, 1985). This is in part due to the restriction of fibre swelling by both lignin and hemicellulose, which limit the 85  penetration of the cellulase to the individual cellulose fibres (Hu et al., 2013; Kumar et al., 2012; Arantes and Saddler, 2011; Jeoh et al., 2007; Eriksson et al., 1991). Additionally, unproductive binding may also occur between cellulases and lignin through hydrophobic, electrostatic and hydrogen bonding interactions (Del Rio, Chandra and Saddler, 2011; Nakagame et al., 2011; Nakagame, Chandra and Saddler, 2010; Berlin et al., 2006; Palonen et al., 2004; Sewalt, Glasser and Beauchemin, 1997). During acid based pretreatments, pseudolignin may also be produced from the degradation of sugars, which can further limit cellulose accessibility and unproductively bind cellulases (Kumar et al., 2013; Sannigrahi et al., 2011). As a result, the differences in substrate composition and properties between model cellulosic substrates and lignocellulosic biomass may influence the distribution of cellulases and subsequent hydrolysis by the adsorbed enzymes. Therefore, to further assess if the limited role of free enzymes in determining the rate and extent of hydrolysis were dependent on the nature of the substrate, the two-stage hydrolysis strategy was also performed on a range of different SO2 catalyzed steam pretreated lignocellulosic substrates (Table 7). Table 7. Steam pretreatment conditions and chemical composition of pretreated lignocellulosic substrates Substrates Pretreatment conditions Composition of substrates (%)a  SO2-steam pretreatment Ara Gal Glu Xyl Man AIL Abbreviation Poplar 190oC, 5min, 3% SO2 0.1 0.2 57.8 4.9 1.1 29.9 SPP190 Poplar 200oC, 5min, 3% SO2 0.1 0.07 57.7 1.5 0.9 33.6 SPP200 Lodgepole pine 200oC, 5min, 4% SO2 0.3 0.5 52.4 1.2 2.1 40.8 LPP200  aAra, arabinan; Gal, galactan; Glu, glucan; Xyl, xylan; Man, mannan; AIL, acid insoluble lignin. 86  Acid catalysed steam pretreatment has been shown to be an effective pretreatment technique that works on a range of lignocellulosic substrates (Chandra et al., 2007). As had been observed previously, although the lignin content in steam pretreated lodgepole pine (softwood) was higher than poplar (hardwood), the cellulose content of the steam-pretreated substrates was similar, ranging from about 52 to 58%. As expected, xylan and mannan were the primary hemicellulose derived sugars found in poplar and lodgepole pine, respectively. When the poplar wood chips were pretreated at two different severities (T=190°C and T=200°C), to try to generate substrates that differed in their xylan content (to determine if the amount of residual hemicellulose might influence the adsorption of cellulases and subsequent hydrolysis), three times more xylan was retained after the lower severity treatment. As was used with DsP hydrolysis, an enzyme loading that resulted in industrially relevant cellulose hydrolysis yields of between 70 and 100% was initially used. At enzyme loadings of 7.5mg/g, 11.5mg/g and 60mg Cellic CTec3/g glucan, hydrolysis yields of 78, 87 and 88% were achieved after 72 hours of hydrolysis of SPP200, SPP190 and LPP200 respectively (Figure 15). In contrast to DsP and the previous enzyme adsorption studies where a significant fraction of the added enzymes remained in solution, most (60 to 85%) of the added enzymes/protein was adsorbed after 1 hour of hydrolysis for all three lignocellulosic substrates.  As both enzyme and substrate properties are known to influence enzyme adsorption, it was possible that the difference in the distribution of adsorbed and free enzymes was due to the use of different enzyme preparations and substrates in the current and past studies. In particular, due to the solubilisation of hemicellulose and minimal removal of lignin during steam pretreatment, the residual lignin in the pretreated substrates might unproductively adsorb the added enzymes. Additionally, current commercial cellulase preparations are known to be 87  complex and multi-component, containing accessory enzymes, such as xylanase and LPMOs, in addition to cellulases (Gupta and Lee, 2013; Merino and Cherry, 2007). Previous work has shown that xylanases can enhance hydrolysis by removing the hemicellulose derived barrier that limits the access of cellulases to cellulose (Hu et al., 2014; Hu et al., 2013; Hu, Arantes and Saddler, 2011; Bura, Chandra and Saddler, 2009; Öhgren et al., 2007). Other work has also shown that the β-glucosidase which is included in newer enzyme preparations adsorbed strongly to a range of steam pretreated substrates, further contributing to the higher amount of enzyme adsorbed observed on the SPP200, SPP190 and LPP200 substrates (Haven and Jorgensen, 2013). Although relatively large amounts of the initial protein added were adsorbed, it was apparent that relatively high hydrolysis yields were still achieved, suggesting that the free enzymes in solution had minimal influence on the overall hydrolysis. To try to elucidate the role of the adsorbed enzymes during the hydrolysis of SPP200, SPP190 and LPP200, any free enzymes in solution were selectively removed using the two-stage hydrolysis strategy while retaining the adsorbed enzymes for subsequent hydrolysis. A similar rate and extent of hydrolysis was observed after the removal of 75% of the free enzymes, which corresponded to a reduction of 11, 19 and 30% of the initial protein loading of SPP200, SPP190 and LPP200 respectively (Figure 15). Specifically, the time course of cellulose hydrolysis of the SPP190 and SPP200 substrates were identical while a 10% decrease in cellulose hydrolysis was observed for LPP200 when using the adsorbed enzymes as compared to the complete enzyme mixture. Thus, it appeared that the adsorbed enzymes determined the rate and extent of hydrolysis regardless of substrate composition and properties.  88   Figure 15. Time course of hydrolysis after the selective removal of 75% of free enzymes in solution after 1 hour of hydrolysis. (a) SPP200 (7.5mg/g Cellic CTec3/g glucan). (b) SPP190 (11.5mg/g Cellic CTec3/g glucan). (c) LPP200 (60mg/g Cellic CTec3/g glucan). SPP200: steam pretreated poplar at 200oC, 5min, 3% SO2; SPP190: steam pretreated poplar at 190oC, 5 min, 3% SO2; LPP200: steam pretreated lodgepole pine at 200oC, 5 min, 4% SO2. 89  3.2.6.  Effect of selective free enzyme removal at different enzyme loadings on enzymatic hydrolysis  Having established that the removal of free enzymes had only a minor to negligible effect on the rate and yield of hydrolysis of pure cellulosic and lignocellulosic substrates, we next wanted to determine if the two-stage hydrolysis strategy was enzyme loading dependant. Thus, all of the DsP, SPP190, SPP200 and LPP200 substrates were hydrolyzed at enzyme loadings that resulted in hydrolysis yields of between 10-40% and 40-70% (Figure 16).  Figure 16. Cellulose hydrolysis of a range of lignocellulosic biomass at different enzyme loadings after 72 hour using a commercial enzyme mixture as compared to two-stage hydrolysis. DsP: dissolving pulp; SPP190: steam pretreated poplar at 190oC, 5 min, 3% SO2; SPP200: steam pretreated poplar at 200oC, 5 min, 3% SO2; LPP200: steam pretreated lodgepole pine at 200oC, 5 min, 4% SO2. As expected, the use of lower enzyme loadings resulted in reduced sugar yields (Figure 16). However, it was apparent that the adsorbed enzymes continued to achieve similar hydrolysis rates and yields as were achieved using the complete enzyme mixture across all of the substrates 90  tested (Table 8). Interestingly, at the lowest enzyme loading tested for DsP, higher cellulose hydrolysis was attained with the adsorbed enzymes than the complete enzyme mixture. As Cellic CTec3 is a highly complex enzyme mixture and DsP is a relatively “pure” cellulosic substrate, the removed free enzymes may contain a large amount of unadsorbed accessory enzymes, which may not be required for the hydrolysis of cellulose. Consequently, there may be a reduced competition for adsorption sites between accessory enzymes and cellulases at lower enzyme loadings. Based on these results, it was apparent that efficient hydrolysis could be achieved using only the adsorbed enzymes regardless of enzyme loadings and substrate characteristics. 91  Table 8. Comparison of enzyme performance of a complete enzyme mixture and adsorbed enzymes after 72 hours of hydrolysis    Enzyme performance (mg glucose/mg protein) Substrates Protein Loading (mg/g glucan) Adsorbed enzymes (mg)a Complete enzyme mixture Adsorbed enzymesa DsP 6 1.9 ± 0.1 67.1 ± 3.3 92.1 ± 11.3 14 3.5 ± 0.2 46.1 ± 1.2 56.2 ± 4.1 22 5.0 ± 0.2 32.5 ± 1.5 40.9 ± 2.5 30 5.9 ± 0.1 26.9 ± 1.7 36.4 ± 1.3 SPP200 1.75 0.4 ± 0.04 135.3 ± 4.5 156.6 ± 6.4 3.75 1.0 ± 0.02 118.3 ± 4.3 131.6 ± 2.2 7.5 2.0 ± 0.05 100.7 ± 5.2 106.7 ± 2.9 SPP190 1.5 0.2 ± 0.008 164 ± 15.5 252.6 ± 10.3 3 0.5 ± 0.04 129 ± 10.1 182.7 ± 7.2 11.5 2.6 ± 0.1 76.6 ± 1.6 106.1 ± 2.7 LPP200 15 2.5 ± 0.01 20.8 ± 0.5 25.1 ± 0.7 37.5 6.6 ± 0.06 13.4 ± 0.7 14.0 ± 0.4 60 9.7 ± 0.06 10.9 ± 0.2 11.1 ± 0.3 aAdsorbed enzymes = Adsorbed enzymes + residual free enzymes after separation by the two-stage hydrolysis strategy 92  3.2.7.  Effect of the complete removal of free enzymes on the enzymatic hydrolysis of pure cellulosic and lignocellulosic substrates  Although it appeared that the removal of free enzymes had only a minor to negligible influence on the rate and extent of hydrolysis of pure cellulosic and lignocellulosic substrates at various levels of enzyme loading, it was apparent that a significant proportion of the substrate was hydrolyzed within the first hour of hydrolysis when using the complete enzyme mixture. As a small amount of free enzymes also remained after separation, this suggested that subsequent hydrolysis was not solely performed by the adsorbed enzymes. To try to further elucidate if the adsorbed enzymes were responsible for the majority of the hydrolysis of pure cellulosic and lignocellulosic substrates, a multi-stage hydrolysis strategy was developed (Table 9).  Table 9. Total protein recovery after separation of adsorbed and free enzymes after 1 hour of hydrolysis at 4oC.   Quantity of protein recovered (mg)a  Substrates Protein Loading (mg/g glucan) Adsorbed protein 1st wash 2nd wash 3rd wash 4th wash Protein mass balance (%) DsP 30 5.4 ± 0.4 6.0 ± 0.009 2.1 ± 0.005 0.9 ± 0.007 0.7 ± 0.007 106.0 SPP200 7.5 1.8 ± 0.07 0.6 ± 0.03 b.d.l b.d.l b.d.l 109.0 SPP190 11.5 1.9 ± 0.2 1.1 ± 0.05 0.3 ± 0.02 b.d.l b.d.l 104.8 LPP200 60 6.8 ± 0.7 4.4 ± 0.01 1.2 ± 0.007 0.6 ± 0.003 0.4 ± 0.001 93.1 ab.d.l: below detection limit 93  In this approach, the initial adsorption and hydrolysis of the various substrates was performed for 1 hour at 4oC instead of 50oC to minimize initial cellulose hydrolysis. Following centrifugation to separate the adsorbed and free enzymes, the insoluble substrates were thoroughly washed with 50mM sodium acetate buffer to remove the loosely bound and free enzymes while only retaining the adsorbed enzymes. Complete removal of free protein/enzymes was defined as achieving less than 5% of the initial protein added in solution after 1 hour of hydrolysis when quantified by the modified ninhydrin assay. The remaining adsorbed enzymes were subsequently used in another round of hydrolysis to assess their hydrolytic performance in the absence of free enzymes for 71 hours, resulting in a total hydrolysis time of 72 hours. As expected, the extent of hydrolysis of both the pure cellulosic and lignocellulosic substrates was only marginally reduced by the removal of the enzymes free in solution (Table 10). Using only the adsorbed enzymes, both SPP200 and SPP190 showed similar hydrolysis yields and DsP and LPP200 only showed slightly less (10%) as compared to those achieved using the complete enzyme mixture. When the enzyme performance between the complete enzyme mixture and adsorbed enzymes was compared the adsorbed enzymes demonstrated equal or higher enzyme performance despite achieving a lower overall conversion yield. These results thus further support the observation that the adsorbed enzymes were primarily responsible for determining the rate and extent of hydrolysis while the free enzymes played a minor role.    94  Table 10. The productivity and performance of a complete enzyme mixture, Cellic CTec3 as compared to adsorbed enzymes after the 72 hour of hydrolysis of pure cellulosic and lignocellulosic substrates  3.2.8.  The potential to reduce enzyme loadings by using a two-stage hydrolysis strategy   Having established that the hydrolysis of lignocellulosic biomass could be mostly achieved using adsorbed enzymes, it was possible that a two-stage hydrolysis approach represented a significant opportunity to reduce the overall enzyme loading require to achieve effective hydrolysis. To try to assess the potential for enzyme savings, four separate empirical models (based on the simplified HCH-1 model where a linear relationship exists between carbohydrate conversion and the natural logarithm of cellulase loading) were developed for each substrate tested (Holtzapple, Caram and Humphrey, 1984). Previous work has shown the linearity of the simplified HCH-1 model was applicable to both pure cellulosic and pretreated lignocellulosic substrates across a wide range of enzyme and solids loadings (Zhu et al., 2010;    Total cellulose hydrolysis (%) Enzyme performance (mg glucose/mg protein) Substrates Enzyme Loading (mg/g glucan) Adsorbed protein (mg) Complete enzyme mixture Adsorbed enzymes Complete enzyme mixture Adsorbed enzymes DsP 30 5.4 ± 0.4 75.5 ± 1.9 65.5 ± 1.9 24.6 ± 0.6 38.4 ± 3.8 SPP200 7.5 1.8 ± 0.07 67.0 ± 0.4 65.6 ± 0.3 88.9 ± 0.7 100.5 ± 0.3 SPP190 11.5 1.9 ± 0.2 94.4 ± 2.2 91.3 ± 0.4 83.5 ± 0.3 131.8 ± 3.7 LPP200 60 6.8 ± 0.7 80.2 ± 2.4 70.6 ± 0.8 13.5 ± 0.4 13.7 ± 0.3 95  O’Dwyer et al., 2007; Mandels, Kostick and Parizek, 1971). For example, a linear relationship was observed during the hydrolysis of lime-pretreated corn stover between 0.25–50 FPU/g dry biomass and solids loading between 10-100g/L (O’Dwyer et al., 2007). As the complete removal of free enzymes with multiple washing steps is unlikely to be employed industrially, this model was subsequently used to predict hydrolysis yields using adsorbed enzymes after two-stage hydrolysis with 75% free enzyme removal and a complete enzyme mixture.   When the 72-hour hydrolysis yields obtained using either the complete enzyme mixture or two-stage hydrolysis with 75% free enzyme removal was plotted against the initial enzyme loading, two distinct relationships were observed (Figure 17). Although, as expected, the overall hydrolysis of the different substrates was strongly dependent on the initial enzyme loading, it was apparent that lower amounts of adsorbed enzymes could be used to achieve similar hydrolysis yields as was obtained with the complete enzyme mixture, as indicated by the larger y-intercepts for all of the substrates tested. These results again suggested that a two-stage hydrolysis with the selective removal of free enzymes and subsequent hydrolysis with adsorbed enzymes could be used as a potential method to reduce enzyme loading.  96    Figure 17. Relationship between enzyme loading and total glucan conversion of a complete enzyme mixture (♦) as compared to the adsorbed and residual free enzymes (■) after separation. (a) DsP. (b) SPP200. (c) SPP190. (d) LPP200. DsP: dissolving pulp; SPP200: steam pretreated poplar at 200oC, 5min, 3% SO2; SPP190: steam pretreated poplar at 190oC, 5 min, 3% SO2; LPP200: steam pretreated lodgepole pine at 200oC, 5 min, 4% SO2. 97   As was used previously, a target of 70% cellulose hydrolysis was used to try to calculate the potential enzyme loading reduction that might be achieved by using a two-stage hydrolysis strategy (Figure 18). It was apparent that any reduction in enzyme loadings that was achieved was significantly influenced by the amount of free enzyme that was detected. For example, due to the high percentage of enzyme free in solution during the hydrolysis of DsP, up to 50% enzyme reduction could be achieved by just using the adsorbed enzymes. In contrast, only a 25-29% saving in protein loading could be achieved with the steam-pretreated substrates due to the higher absorption of the added protein to these substrates.   Figure 18. Relative reduction in enzyme loading to achieve 70% glucan conversion when using a complete enzyme mixture as compared to the adsorbed and residual free enzymes after separation. DsP: dissolving pulp; SPP200: steam pretreated poplar at 200oC, 5min, 3% SO2; SPP190: steam pretreated poplar at 190oC, 5 min, 3% SO2; LPP200: steam pretreated lodgepole pine at 200oC, 5 min, 4% SO2.   98  3.2.9.  Conclusions The work in this chapter confirmed how difficult it is to accurately track enzyme/protein distribution when using a complex “cellulase mixture” and a range of heterogeneous, lignocellulosic substrates. Much of the previous work in this area, where researchers had tried to assess the role that adsorbed cellulase enzymes play in determining the rate and extent of cellulose hydrolysis, had tried to simplify the process by using either individual enzymes and/or simplified cellulase mixture on “pure” cellulosic substrates. However, enzyme-substrate interactions are known to be greatly influenced by the composition/source of enzymes and the structural properties of the substrates, making interpretations from this earlier work problematic. By using the improved protein determination method outlined in Chapter 3.1, we were able to assess the relative contributions that the adsorbed and unabsorbed enzymes played during cellulose hydrolysis. It was apparent that the adsorbed enzymes played a critical role, with the majority of the hydrolytic action carried out by these enzymes. This was the case with both the “model” dissolving pulp substrate and with all of the pretreated lignocellulosic substrates. As anticipated, the enzyme adsorption profiles differed depending on the nature and type of the cellulosic substrates, likely due to differences in cellulose accessibility and lignin type and content. Surprisingly, regardless of the enzyme loadings or the nature of the substrates, by carrying out hydrolysis using just the adsorbed enzymes, similar hydrolysis yields and hydrolytic performances were achieved as were obtained when using the complete (those in solution and those adsorbed) enzyme mixture. This supported earlier work by others who had suggested that the enzyme components that were in close association with the substrate played the major role in achieving effective cellulose hydrolysis. By only using the adsorbed enzymes associated with the residual recalcitrant substrate, protein loadings could be reduced from 25% up to 53% on 99  SPP200 and DsP respectively, while achieving similar hydrolysis yields as were obtained using both soluble and adsorbed enzymes.    100  4. Final conclusions and future work 4.1. Conclusions The main focus of the thesis work was to better understand the relative contributions that the adsorbed and free cellulases might play during the hydrolysis of cellulosic substrates. Primarily because of the complexity of both the enzyme system and the heterogeneous nature of the substrate, most of the previous work that has looked at enzyme adsorption and hydrolytic performance has generally used individual enzymes and/or simplified cellulase mixture and “model” cellulosic substrates, such as Avicel, Solka floc or cotton. However, as the current work has shown, enzyme adsorption can be influenced by several factors, such as substrate and hydrolysis condition factors.  A modified ninhydrin based protein quantification assay was successfully developed that addressed several of the limitations of the currently available protein determination methods. A sodium borohydride treatment coupled with hydrolysis temperatures of 130oC reduced the assay time by 6-fold (to 4 hours) and accurately quantified the protein concentration of a commercial enzyme mixture in the presence of a range of steam pretreated biomass derived water soluble fractions. The modified ninhydrin assay was subsequently used to accurately monitor enzyme distribution and performance during the hydrolysis of a range of “model” and steam pretreated lignocellulosic substrates. As initially anticipated, when the modified ninhydrin assay was used to monitor the adsorption profile of enzymes present in the Cellic CTec3 commercial enzyme mixture, the initial protein distribution was shown to be dependent on the physicochemical nature of the substrate. When subsequent hydrolysis was carried out using just the adsorbed enzymes associated with the residual recalcitrant substrate, similar hydrolysis yields as were obtained when adding a complete enzyme mixture could be achieved. This occurred regardless 101  of the substrates or enzyme loadings employed. This strongly suggested that the adsorbed enzymes were primarily responsible for determining the rate and extent of hydrolysis.  4.2. Future work 4.2.1. The influence of initial enzyme distribution in determining the rate and extent of hydrolysis at high solids loading Typical industrial enzymatic hydrolysis processes operate at high solids loadings as it increases the sugar concentrations, reduces operational costs and minimizes energy consumption (Kristensen, Felby and Jorgensen, 2009; Hodge et al., 2008). However, previous works have observed that there is a decrease in hydrolysis yields with increasing solids (Varnai, Siika-Aho and Viikari, 2013; Wang et al., 2011; Kristensen, Felby and Jorgensen, 2009). One of the potential factors that have been proposed for this “high solids” inhibition effect is an increase in the physical constraint of water, which limits enzyme movement to reaction sites within the substrate (Hsieh et al., 2014; Selig et al., 2012; Roberts et al., 2011). Since the initially adsorbed enzymes were demonstrated in this thesis to be primarily responsible for determining the rate and extent of hydrolysis at low solids loading, it is likely that achieving efficient initial enzyme distribution might also be essential for efficient hydrolysis at high solids loadings. As the two-stage hydrolysis strategy applied in this thesis for the separation of the “bound” and “free” enzymes at the initial stage of hydrolysis can be used to achieve a concentrated biomass system with well distributed enzymes, it would be interesting to assess how this hydrolysis strategy will improve the hydrolytic performance of commercial cellulase preparations at high solid loading hydrolysis as well.  102  4.2.2.  Potential recycling of the free enzymes for further rounds of hydrolysis  The thesis work showed that a two-or-multi-stage hydrolysis strategy is an effective method of reducing enzyme loadings by performing hydrolysis with only adsorbed enzymes. However, the role and function of the free enzymes remain unclear. Previous works have demonstrated that the free enzymes in solution after separation continue to remain active, but have significantly reduced hydrolysis ability (Yu et al., 2013; Pribowo, Arantes and Saddler, 2012). It has been proposed that the loss in activity may be due to the removal of key essential enzyme activities for lignocellulose deconstruction (Yu et al., 2013). Therefore, the potential recycling of the free enzymes and the requirement for the supplementation of additional enzymes in further rounds of hydrolysis should be investigated. Subsequently, this information could then be used to develop new methods of reducing enzyme usages commercially.  For example, a potential application of free enzyme recycling during a commercial hydrolysis process is the implementation of a continuous hydrolysis process. In this process, the filtrate containing the free enzymes following solid-liquid separation may be added to new substrate instead of fresh water for a subsequent round of hydrolysis. Additional enzymes may then be supplemented to replenish enzyme activities required to achieve efficient hydrolysis. By employing this hydrolysis strategy, the amount of enzymes required to achieve effective hydrolysis might be reduced through multiple rounds of hydrolysis, potentially reducing operational costs.   103  4.2.3.  Further assessment of the role of adsorbed enzymes on different cellulose morphologies and pretreated lignocellulosic substrates This study showed that similar rate and extent of hydrolysis on pure cellulosic dissolving pulp (DsP) and steam pretreated lignocellulosic substrates were obtained with adsorbed enzymes as compared with a complete commercial enzyme mixture. However, other methods, such as mechanical, AFEX, organosolv, ionic liquid and hydrothermal pretreatments have also been investigated as potential pretreatment strategies in the industrial bioconversion process (Chandra et al., 2007; Mansfield, Mooney and Saddler, 1999). As the physicochemical nature of the substrate (cellulose accessibility/DP/crystallinity/allomorphs and hemicellulose/lignin properties and content) is known to greatly influence enzyme adsorption, it would be interesting to investigate if the adsorbed enzymes continue to be primarily responsible for enzymatic hydrolysis on those substrates (Van Dyk and Pletschke, 2012; Mansfield, Mooney and Saddler, 1999). Therefore, the role of adsorbed enzymes on different cellulose morphologies (PASC/Avicel/CNC/Cellulose II/Cellulose III) and pretreated lignocellulosic substrates will be of interest to further investigate.  104  References Abdek-Akher, M., Hamilton, J.K., Smith, F., 1951. 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