Instituto Tecnológico y de Estudios Superiores de Monterrey
Campus Monterrey
School of Engineering and Sciences
Mathematical modeling of the enzymatic saccharification process of lignocellulosic biowaste
A thesis presented by
Vittorio Fedeli
Submitted to the
School of Engineering and Sciences
in partial fulfillment of the requirements for the degree of Master of Science
In Biotechnology
Monterrey Nuevo León, November 2020
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@2020 by Vittorio Fedeli All rights reserved
Dedication
To my brother Alessandro, thank you for being there every time I needed it and for all the advice and knowledge shared with me. Keep always moving and never stop being yourself.
To my mother, I admire you as you overcome every obstacle that presents to you. I love your strength and discipline.
To my father, thank you for always taking the time to advise and listen to me.
To PhD. Rodrigo Balam Muñoz Soto and his family. Thank you for all the time and for inculcating in me the passion for science.
To Pau, words are hard to find to thank you for all the affection and
unconditional love in the day to day.
v Acknowledgements
I want to thank MSc. Carlos Eduardo Gomez Sanchez and Ph.D.
Marco Arnulfo Mata Gomez for always pushing forward in my formation as a scientist, for your excellent guidance, patience, and the extra time dedicated to achieving this project.
I would like to express my acknowledgment to the Tecnologico de Monterrey, who provided support on tuition, and CONACyT, who provided the economic support for this work.
I would also like to express my sincere gratitude to MM SCM Hector Adiel Flores Nestor for the guidance and the time given to solve many of the obstacles presented during this project's development.
To Javier Ingelmo, for helping me with the Figures construction and your patience to understand my drawings.
To finish, to all my friends who always trust in me. Friendship is
something that I really appreciate.
vi
Mathematical modeling of the enzymatic saccharification process of lignocellulosic biowaste
by
Vittorio Fedeli
Abstract
Lignocellulose is a biowaste produced in large quantities by industries; approximately 181.5 billion tons are produced annually in the world. This makes this type of residue a qualifiable candidate resource of energy, which nowadays, is underutilized. It is estimated that the food processing industry produces around 1.3 billion tons per year. In Mexico, the craft beer industry produces 3.8 thousand tons per year of brewers' spent grain. Being Mexico's fastest- growing industry, it can be considered a suitable source of biowaste. Brewers spent grain is considered a lignocellulosic material, which possesses a complicated structure containing lignin, hemicellulose, and cellulose. Due to its complexity, diversity, and recalcitrance to degradation, specific pretreatments to degrade it have been developed, such as biological, chemical, physical, and physicochemical. Notwithstanding, in nature, fungi are well-known microorganisms capable of degrading it through a tremendous battery of enzymes that are secreted in an ordered and systematic fashion.
Nonetheless, the full understanding of this process, and the order in which each enzyme acts on lignocellulose, is far to be elucidated. Therefore, the present thesis aims to develop fungi bio-inspired mechanistic mathematical model capable of describing the enzymatic degradation process of lignocellulose biomass (brewers spent grain) and evaluate it through different experiments. Sequential addition of enzymes to biowaste, as well as experiments involving the addition of a pool without key enzymes that were further added at a specific time, were evaluated. Overall, results revealed that the lignin is not the most resilient and dense layer of lignocellulose as it has been believed. On the contrary, it seems lignin forms pore-like structures and diffuses through all different layers of this substrate. When hemicellulases (xylanases and pectinases) were not present in the enzyme pool, the reaction was not favored, indicating the importance of this polymer in lignin structure. These results gave an idea of how fungi work in nature and how the polymer layers are organized in lignin.
However, to fully confirm these findings, more tests need to be performed to generate a robust and proven mechanistic mathematical model, enabling us to lay the foundations of a potential industrial-scale process.
vii
List of Figures
Fig. 1. Lignocellulose Molecule a theoretical based proposal of its structure ... 6
Fig. 2. Pectin molecules and degrading enzymes ... 8
Fig. 3. Schematic illustration of hemicellulose type molecules. ... 13
Fig. 4. Hemicellulose structures and enzymes. ... 14
Fig. 5. Schematic representation of cellulose enzymatic hydrolysis ... 17
Fig. 6. Longitudinal cross-section of barley kernel. ... 18
Fig. 7. Lignocellulose biomass pretreatment... 22
Fig. 8. Mathematical model abstraction ... 26
Fig. 9.First mechanistic mathematical model plot result. s. ... 34
Fig. 10. Thermal degradation of substrate. ... 36
Fig. 11. Single Enzyme Influence test. ... 37
Fig. 12. Dynamic dual enzyme test.. ... 39
Fig. 13. Dynamic enzyme test results.. ... 41
Fig. 14. Second approach mathematical abstraction. ... 42
viii
List of Tables
Table 1. Brewers spent grain lignocellulose composition. ... 20
Table 2. Dynamic enzyme test methodology. ... 32
Table 3. Dextrose quantification methodology. ... 47
Table 4. Enzyme pool test methodology... 47
Table 5. Laccase influence test methodology. ... 48
Table 6. Xylanase Influence test methodology. ... 48
Table 7. Alpha amylase influence test methodology. ... 48
Table 8. Pectinase Influence test methodology... 49
Table 9. Cellulase influence test methodology. ... 49
Table 10. Dynamic test of Xylanase at 60 min and Pectinase at 120 min. ... 50
Table 11. Dynamic test of Pectinase at 60 min and Xylanase at 120 min methodology. .... 50
Table 12. Dynamic test of Cellulase at 60 min and Alpha amylase at 120 min methodology. ... 51
Table 13. Dynamic test of Alpha amylase at 60 min and Cellulase at 120min methodology. ... 51
Table 14. Dynamic test of Laccase at 60 min and Cellulase at 120 min methodology. ... 51
Table 15. Dynamic test of Cellulase at 60 min and Laccase at 120 min methodology. ... 52
ix
Contents
Abstract vi
List of Figures vii
List of Tables viii
Chapter I ... 1
1.1 Introduction... 2
1.2 Hypothesis ... 3
1.3 General Objective ... 3
1.4 Specific Objective ... 3
2 Chapter II ... 4
2.1 Lignocellulose Molecule and Degradation ... 5
2.1.1 Lignin molecule ... 5
2.1.2 Pectin Molecule: ... 7
2.1.3 Hemicellulose ... 11
2.1.4 Cellulose ... 16
2.2 Brewers spent grain and applications ... 17
2.3 Successful mathematical models for enzymatic degradation of polymers. ... 20
2.4 Existing lignocellulose pretreatments. ... 21
Chapter III ... 25
3 Materials and methods ... 25
3.1 Mathematical model ... 26
3.1.1 Conception of the model ... 26
3.1.2 Differential equations ... 27
3.2 Experimental section ... 28
3.2.1 DNS Reactant preparation ... 28
3.2.2 100 mM Acetate buffer pH 5.0 ... 28
x
3.2.3 Enzyme stock solutions ... 29
3.2.4 Dialysis of enzyme stock solutions ... 29
3.2.5 Brewers spent grain raw material ... 29
3.2.6 Substrate thermal degradation test ... 30
3.3 Experimental enzyme tests ... 30
3.3.1 Enzyme pool test ... 31
3.3.2 Single enzyme influence tests. ... 31
3.3.3 Dynamic dual enzyme tests. ... 31
3.3.4 Dynamic and sequential enzyme test ... 32
4 Chapter IV ... 33
4.1 Results and Discussion ... 34
4.1.1 First approach for mechanistic mathematical model: a theoretical approach... 34
4.1.2 Thermal degradation of brewer’s spent grain ... 35
4.1.3 Single enzyme influence test ... 36
4.1.4 Dynamic dual enzyme test ... 38
4.1.5 Dynamic and sequential enzyme test ... 40
4.1.6 Second mechanistic mathematical model proposal ... 41
5 Chapter V... 45
5.1 Conclusion and perspectives ... 46
6 Appendix ... 47
6.1 Appendix A: Dextrose Quantification ... 47
6.2 Appendix B: Enzyme pool test ... 47
6.3 Appendix C: Single enzyme influence tests. ... 48
6.3.1 Influence of Laccase. ... 48
6.3.2 Influence o Xylanase. ... 48
6.3.3 Influence of Alpha amylase. ... 48
6.3.4 Influence of Pectinase. ... 49
6.3.5 Influence of Cellulase. ... 49
6.4 Appendix D: Dynamic dual enzyme test ... 50
6.4.1 Addition of Xylanase at 60 min and Pectinase at 120 min. ... 50
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6.4.2 Addition of Pectinase at 60 min and Xylanase at 120 in. ... 50 6.4.3 Addition of Cellulase at 60 min and Alpha amylase at 120 min.
51
6.4.4 Addition of Alpha amylase at 60 min and Cellulase at 120 min.
51
6.4.5 Addition of Laccase at 60 min and Cellulase at 120 min. ... 51
6.4.6 Addition of Cellulase at 60 min and Laccase at 120 min. ... 52
7 References ... 53
1
Chapter I
Introduction
2
1.1 Introduction
Lignocellulosic biowastes are produced mainly by agricultural practices, food processing industry, fabric, and paper industries, and can be defined as a complex heteropolymer consisting of four molecules: lignin, hemicellulose, pectin (not in all cases), and cellulose.
Upon the monomeric units of these structures, resides its energetic potential, which remains unexploited. Depending on the source, individual concentration may vary. In general terms, lignin represents between 10 to 25%, hemicellulose from 23 to 32%, and cellulose between 38 to 50% of the total plant composition (Paul & Dutta, 2018).
Lignocellulose global production consists of approximately 181.5 billion tons, qualifying as an abundant resource, however underutilized (Paul & Dutta, 2018). One of the fastest- growing industries in Mexico corresponds to the craft brewing industry, with an average growth rate of 53.3 % and approximate lignocellulosic waste generation of 3.8 thousand tons per year of the so-called Brewers' spent grain (ACERMEX, 2018). This biowaste is composed of insoluble proteins and grain coverings rich in lignin, hemicellulose (arabinoxylan and xylan), cellulose, and the most common disposal method is as an animal feed (low value) (Carvalheiro et al., 2004).
Due to its complex structure and recalcitrance to degradation, specific methods have been developed, such as biological, chemical, physical, and physicochemical pretreatments, being the former the most straightforward. However, in most cases, significant drawbacks such as high environmental cost, high process costs, and scaling challenges have prevented them from being widely accepted (Haghighi Mood et al., 2013).
In nature, fungi secrete, in a sequential fashion, a set of enzymes such as cellulases, pectinases, hemicellulases, and xylanases that synergically degrade lignocellulosic materials efficiently. On the contrary, in industry, this set of enzymes is added to the biowaste as pools, and a high yield of conversion is achieved only when adding great amounts of enzymes. We believed that by sequentially adding the enzymes as mimicking what fungi do in nature, a higher degradation of biowaste could be attained. Nonetheless, the process of how fungi do it is far from being understood.
Some authors have tried to understand this biowaste and its degradation by establishing mathematical models (Pollegioni et al., 2015). Nevertheless, these models end up being challenging to solve, with high quantities of parameters that are difficult to determine.
Besides, they are fed with arbitrary data instead of being calculated with experimental data and considers only cellulose degradation as a homogeneous substrate instead of considering it as a heterogeneous and multi-structural system (Ye & Berson, 2011).
Therefore, the lack of mechanistic mathematical models that integrates lignocellulose as a heterogenic substrate consisting of lignin, hemicellulose, pectin (when present), and cellulose with the interaction between enzymes and different layers of the substrate opens the possibility for the development of a fungi bio-inspired mechanistic mathematical model, object of the present thesis.
3 In this work, we have developed a mathematical model that explains how enzymes degrade lignocellulosic material from brewer spent grain over time in a mechanistical fashion. It is believed that this model, along with experimental trials consisting of sequential addition of enzymes to the biowaste, will allow understanding not only how the different layers are arranged in the biomaterial but also to give an idea of how fungi do this process in nature.
This would allow optimizing processes with no need of adding great amounts of enzymes, thus reducing costs.
1.2 Hypothesis
The development of a mechanistic mathematical model bio-inspired in fungi that describes the enzymatic degradation process of lignocellulosic biowaste will allow us to understand how layers of the polymers are arranged, enzyme synergism, and enzyme-substrate interactions.
1.3 General Objective
To develop a fungi bio-inspired mechanistic mathematical model capable of describing the enzymatic degradation process of lignocellulose biomass.
1.4 Specific Objective
▪ To develop a mechanistic mathematical model that considers the enzymes' sequential interactions and interaction with the lignocellulosic material, as found in the literature.
▪ To experimentally evaluate the synergism of the main enzymes through sequential addition of enzymes to the brewers' spent grain to understand how layers are arranged.
▪ To evaluate the mechanistic mathematical model through experimental results.
▪ To reformulate the mathematical approach based on the results obtained.
4
2 Chapter II
Background
5
2.1 Lignocellulose Molecule and Degradation 2.1.1 Lignin molecule
Lignin represents one of the three constituents of lignocellulosic biomass. Its amorphous structure and chemical reactivity represent the most arduous component for degradation through a physical, chemical, or enzymatic process. Adding to scientists' low interest to develop efficient strategies capable of degrading it due to its absence of carbohydrates in its backbone (low energy potential). Nonetheless, some scientists, such as Levasseur et al., (2008), consider lignin a renewable source for aromatic polymers. The importance of lignin degradation relies on this compound's resilience to be degraded by enzymes and lignocellulose location. This is located at the outer cell wall that protects the rich saccharides cell wall polymers as pectin, hemicellulose, and cellulose. It is defined as an aromatic heteropolymer formed by polymerization of ρ-hydroxyphenyl (ρ- coumaryl alcohol building block), guaiacyl (Coniferyl alcohol building block) and syringyl (Sinapyl alcohol building block) units linked by three different types of bonds such as heterocyclic linkages, biphenyl bonds and β-aryl ether linkages (Zandleven et al., 2005).
The main lignin-degrading microorganisms correspond to white-rot fungi. With less efficiency, brown-rot fungi, some of these microorganisms are Coprinipsis cinereal, Phanerochaete chrysosporium, Ustilago maydis, Aspergillus nidulans, and Trichoderma reesei (Martínez-Trujillo et al., 2020). All lignin-degrading microorganisms can synthesize a group of extracellular enzymes that allow mineralized lignin. These enzymes can be classified into two groups: (1) Lignin modifying enzymes and (2) Lignin accessory enzymes, which cannot degrade lignin by their action; instead, they eliminate steric components and facilitates the action of lignin modifying enzymes. All lignin-degrading enzymes are related to the group of oxidoreductases.
2.1.1.1 Backbone degrading enzymes:
Lignin modifying enzymes that are responsible for attacking the lignin backbone are: lignin peroxidases ( EC 1.11.1.14) that catalyzes the depolymerization of lignin non-phenolic compounds through the oxidation of a heme group from Fe (III) to Fe(IV) by an oxidative specie of hydrogen peroxide (Pollegioni et al., 2015). Manganese peroxidase (EC 1.11.1.13) which attack lignin phenolic compounds by oxidation of a heme group from Mn (II) to Mn(III) by hydrogen peroxide and a chelator agent (Giardina et al., 2010); the oxidant power of this enzyme is transferred to Mn(III),which diffuses through lignin and attach to phenolic ring from the internal site (Ander & Marzullo, 1997). Versatile peroxidase (EC 1.11.1.16) that acts through various catalytic sites, Mn(II) heme group responsible for electron transfer and high redox potential to oxidize aromatic substrates, veratryl alcohols, non-phenolic and phenolic lignin, between others (Janusz et al., 2017), and laccase (EC 1.10.3.2) that is a
6 polyphenol oxidase that uses the low redox potential of four copper ions to catalyze the oxidation of an array of aromatic substrates (Ander & Marzullo, 1997; Janusz et al., 2017).
Peroxidases such as Lignin peroxidases (EC 1.11.1.14) and manganese peroxidases (EC 1.11.1.13) from Phanerochaete chrysosporium were induced by limiting nitrogen; if present in excess, lignin degradation will be inhibited in most white-rot fungi (Ai et al., 2014).
2.1.1.2 Accessory enzymes:
The lignin accessory enzymes are glyoxylate oxidase (EC1.2.3.5) that possess a free radical coupled to copper active site and produces H2O2 ,which is used as a substrate for lignin modifying enzymes to oxidize the product of lignin peroxidase such as glycolaldehyde (Janusz et al., 2017), and aryl-alcohol oxidase (EC 1.1.3.7) that oxidizes molecular oxygen into hydrogen peroxide mainly through the dehydrogenation of phenolic and non-phenolic aryl alcohols, such as aromatic secondary alcohols or polyunsaturated primary alcohols into the corresponding aldehydes (Martens-Uzunova & Schaap, 2009). Another enzyme is
Fig. 1. Lignocellulose Molecule a theoretical based proposal of its structure
7 pyranose oxidase (EC 1.1.3.10), which acts through the reduction of O2 into H2O2 needed for most of lignin-degrading enzymes. It can reduce various quinones and radicals produced by laccases avoiding re-polymerization of lignin (van den Brink & de Vries, 2011). Other relevant enzymes are glucose oxidase (EC 1.1.3.4) that oxidizes glucose to gluconolactone through two electron transfer to a flavin prosthetic group, which is then re-oxidized through a second two-electron transfer, and glucose dehydrogenase (EC 1.1.5.9) that catalyzes anomeric hydroxy group of glucose and FAD group is used as an electron acceptor (Kauppinen et al., 1995).
2.1.2 Pectin Molecule:
It is a heteropolymer present in the middle lamella of the primary plant cell wall and constitutes one-third of the total cell wall composition, being the less abundant lignocellulose component compared to lignin, hemicellulose, and cellulose (Kauppinen et al., 1995; Wong, 2008; Zandleven et al., 2005). The backbone of pectin is constituted mainly by α-1,4 linked D-galacturonic acid residues that can be methyl-esterified or substituted with acetyl groups (Martens-Uzunova & Schaap, 2009; van den Brink & de Vries, 2011).
The monomeric constituent of pectin is of interest for some wood degrading fungi and bacteria as it can be metabolized as a carbon source and for scientific purposes as it contains an energetic potential that can be used for multiple purposes such as biofuel production and added value monosaccharides manufacturing. Pectin polysaccharides create a matrix that embedded the cellulose and hemicellulose substrates, which are rich in monosaccharides.
Therefore, this matrix's correct degradation enables hemicellulose and cellulose-degrading enzymes to access their corresponding substrates (van den Brink & de Vries, 2011).
Pectin molecules are classified into four general groups, as shown in Figure 2. (1) Homogalacturonan that consists of a linear polymer composed of approximately 200 - 1,4- linked α-D- galacturonic acid residues. Some regions of the galacturonic acid monomer can either be methyl-esterified at the carboxylic acid group (C-6 position) or acetylated (C-3 or C-2 position) (Nighojkar et al., 2019). (2) Xylogalacturonan, where galacturonic acid residues from the homogalacturonan backbone are β-1,3-Linked to β-D-xylose residues as simple ramifications.
(3) Rhamnogalacturonan I, in which its backbone is formed by alternating α-1,2 linked L- rhamnose and α-1,4-linked galacturonic acid residues. On this molecule, L-rhamnose can be branched with O-4 attached L-arabinose or D-galactose side chains that can vary in the degree of polymerization. The more complex ramifications for this molecule are formed by L-arabinose and D-galactose, forming arabinogalactan side chains, and L-arabinose D- glucose ramifications can be substituted with ferulic acids.
(4) Rhamnogalacturonan II is the most complex (highly substituted) and less common pectin found in nature, and its backbone consists of D-galacturonic acid residues linked by α-1,4- linkages branched with rare sugars; these ramifications follow a conserved structural pattern.
8 The basic structure of pectin depends on the distribution of the groups mentioned above.
Nonetheless, pectin smooth regions are constituted of homogalacturonan, and the hairy or ramified regions are composed of xylogalacturonan, rhamnogalacturonan I, or rhamnogalacturonan II (Atanasova et al., 2018). Therefore, degradation of hairy regions requires an extra battery of accessory enzymes due to its complexity and steric impediment that ramification represents for pectin backbone degrading enzymes.
The most studied fungi that produce a battery of enzymes to fully degrade pectin correspond to Aspergillus niger, as shown by the studies performed by Kumar et al., (2008).
Nevertheless, other fungi have reported to produced sufficient enzymes that confer them the ability to degrade pectin; these are Rhizopus cryzae, Aspergillus tubingensis, Aspergillus nidulans, and Aspergillus aculeatus (Mayans et al., 1997).
Due to the complexity and diversity of pectin groups, the classification of pectin degrading enzymes is divided based on the molecule that they act on and by recognizing specific sites located on the backbone or the substituents groups.
Fig. 2. Pectin molecules and degrading enzymes
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2.1.2.1 Backbone degrading enzymes:
2.1.2.1.1 Homogalacturonan degrading enzymes:
The homogalacturonan degrading enzymes that attack the backbone corresponds to exo-poly- α-digalacturonosidase (EC 3.2.1.82), an exo-acting enzyme that targets the second α-1,4- glycosidic bond from the non-reducing end of homogalacturonan molecule to generate digalacturonate molecules (Mayans et al., 1997; van den Brink & de Vries, 2011). Pectate disaccharide - lyase (EC 4.2.2.9) that is responsible for the eliminative cleavage of glycosidic linkages in the homogalacturonan molecule from the non-reducing end; it is essential to mention that this enzyme is dependent on Ca+2 and it produces 1,4-galacturonic acid, 4,5- galacturonate polysaccharide and unsaturated digalacturonic acid (Giovane et al., 2004).
Endo-polygalacturonase (EC 3.2.1.15) that generates cleave of 1,4-α-D-galactosiduronic linkages from non-reducing ends to produce D-galacturonate and galacturonic acid units (Kumar et al., 2008). Another enzyme is the endo-pectin lyase (EC 4.2.2.10), which recognizes highly methylated forms of homogalacturonan with an action mechanism that consists of the eliminative cleavage of 1,4-α-D-galacturonan methyl ester to produce oligosaccharides (Wong, 2008).
Pectate lyase (EC 4.2.2.2) is an enacting enzyme that possesses higher activity over pectate molecules, demethylated or low esterified forms of pectin, and is dependent on Ca+2 to be active; the action mechanism consists of eliminative cleavage of the 1,4-α-D-galacturonan to produce oligosaccharides with 4-deoxy-α-D-galact-4-enurosyl groups at non-reducing ends (Rozeboom et al., 2013). Finally, an accessory enzyme for the homogalacturonan molecule is pectinmethylesterase (EC 3.1.1.11) that oversees the removal of methyl substituents from galacturonic acid residues to produce methanol and pectate. Its action allows access to backbone degrading enzymes (Wong, 2008).
2.1.2.1.2 Xylogalacturonan degrading enzymes:
The degradation of xylogalacturonan depends on several enzymes that involve backbone degrading enzymes and accessory enzymes. The backbone degrading enzymes are endo- polygalacturonase (EC 3.2.1.15), which is in charge of the internal cleavage of 1,4-α-D- galactosiduronic linkages of the non-reducing ends of both homogalacturonan and xylogalacturonan molecules to produce D-galacturonate and Galacturonic Acid polysaccharides (Wong, 2008).Galacturan 1,4-α-galacuronidae (EC 3.2.1.67) is an exo type enzyme that acts by releasing β-D-xylose-1,3-α-D- galacturonic acid and D-xylose-D- galacturonic acid disaccharides from non-reducing ends (Kumar et al., 2008; van den Brink
& de Vries, 2011). The xylogalacturonan hydrolase (EC 3.2.1.-) recognizes two adjacent β- xylose substituted galacturonic acids and hydrolyses the glycosidic bond (Mayans et al., 1997; van den Brink & de Vries, 2011).
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2.1.2.1.3 Rhamnogalacturonan I and II degrading enzymes:
Rhamnogalacturonan type I and II are the most complex pectin type; more enzymes are required to degrade it. The backbone degrading enzyme consists of a battery of enzymes.
Rhamnogalacturonan hydrolase (EC 3.2.1.171) cleaves the bond between the L- rhamnose and the D-galacturonic acid by hydrolysis to produce rhamnogalacturonan polysaccharides with β-D-galacturonic acid residues at the reducing ends. The presence of acetyl substituents represents a steric impediment; therefore, this enzyme act synergistically with rhamnogalacturonan acetylesterase (EC 3.1.1.86) (Ronald P. de Vries and Jaap Visser, 2001).
Rhamnogalacturonan galacturonohydrolase (EC 3.2.1.173) is an exo-acting enzyme in charge of removing terminal galacturonic acid residues from the non-reducing end of rhamnogalacturonan molecule and some rhamnogalacturonan oligosaccharides. This is possible through a single displacement mechanism to release D-galacturonic acid (Hövel et al., 2003; Saha & Bothast, 1999). Rhamnogalacturonan rhamnohydrolase (EC 3.2.1.174) is another exo-acting enzyme that removes the terminal rhamnosyl residues from the non- reducing end from rhamnogalacturonan and its oligosaccharides to release β-L- rhamnopyranose; substituents such as galactose block the hydrolytic action of the enzyme (de Lima et al., 2016). The α-L-rhamnosidase (EC 3.2.1.40) is responsible for the cleavage of the non-reducing α-L-rhamnose residues from rhamnogalacturonan I to release α-L rhamnose and its polysaccharide (Mølgaard et al., 2000).
Unsaturated rhamnogalacturonyl hydrolase (EC 3.2.1.172), which is an exo-acting enzyme that acts on the rhamnogalacturonan generated by rhamnogalacturonan endolyase (EC 4.2.2.23) and releases unsaturated galacturonic acid, α-D-galacturonic acid, 4-deoxy-β-L- threo-hex-4-enepyranosyl uronic acid, and its function is related to the unsaturated glucuronyl hydrolase (EC 3.2.1.179) (Mayans et al., 1997; van den Brink & de Vries, 2011;
Wong, 2008). Rhamnogalacturonan exolyase (EC 4.2.2.24) attacks the rhamnogalacturonan produced by rhamnogalacturonan endolyase (4.2.2.23) containing α-L-rhamnopyranose at the reducing position and 4-deoxy-4,5-unsaturated D-galactopyranosyluronic acid at the non- reducing position. It cleaves the α-L-rhamnopyranosyl-1,4-α-D-galactopyranosyluronic acid to produce disaccharide 2-O-4-deoxy-β-L-threo-hex-4-enopyranuronosyl-α-L- rhamnopyranose and a shorter rhamnogalacturonan oligosaccharide containing one-4-deoxy- 4,5-unsaturated D-galactopyranosyluronic acid at the non-reducing end (Gírio et al., 2010).
2.1.2.2 Accessory enzymes
The accessory enzymes that allow the complete degradation of rhamnogalacturonan are arabinan endo-1,5-α-L-arabinase (EC 3.2.1.99) that is responsible for the internal hydrolysis of the α-1,5-linkages of arabinan polysaccharides present as substituents groups and
11 producing α-D-arabino-pentofuranosyl and α-L-arabinofuranosyl and possess higher activity with non-branched arabinans (Ren & Sun, 2010).
Non-reducing end α-L-arabinofuranosidase (EC 3.2.1.55), an exo-acting enzyme responsible for the degradation of side chains and substituents in xylan hemicellulose and rhamnogalacturonan I molecules. Cleaving the L-arabinose substituents at the non-reducing ends by hydrolysis of the ester linkages between xylose or arabinose molecules and producing α-L-arabinofuranose, arabinobiose, α-1,5-L-arabinofuranotriose, and L-arabinose (Gírio et al., 2010). Arabinogalactan endo-β-1,4-galactanase (EC 3.2.1.89) that is responsible for the degradation of arabinogalactan side chains by hydrolysis of β-1,4-galactosidic bonds in arabinogalactan and galactan side chains to produce D-galactose, β-1,6-galactose, and D- galactooligosaccharides (Gírio et al., 2010; Ren & Sun, 2010).
The rhamnogalacturonan acetylesterase (EC 3.1.1.86) is in charge of the deacetylation of rhamnogalacturonan I by hydrolytic cleavage of 2-O-acetyl groups from substituted α-D- galacturonic acid residues (Ren & Sun, 2010).
Moreover, rhamnogalacturonan endolyase (EC 4.2.2.23) is specific for pectin hairy regions containing rhamnogalacturonan I and cleaves the L-α-rhamnopyranosyl-1,4-α-D galactopyranosyluronic acid bonds leaving the pectin molecule with an L-rhamnopyranose at the reducing end and 4-deoxy-4,5-unsaturated D-galactopyranosyluronic acid at the reducing end. With this, the pectin molecule is ready to be degraded by rhamnogalacturonan exolyase (EC 4.2.2.24) (van den Brink & de Vries, 2011).
2.1.3 Hemicellulose
Hemicellulose corresponds to the second most abundant material preceded by cellulose in the world. It constitutes around one-fourth of the total plant cell wall material, and it is composed of a variety of monosaccharides such as mannose, xylose, galactose, fucose, arabinose, glucose, galacturonic acid, and glucuronic acid. The hemicellulose integrates with the other lignocellulose components by hydrogen bonds to cellulose and by α-benzyl-ether linkages (covalent bond) to lignin (Pedersen et al., 2009).
Depending on the source, hemicellulose can be structurally diverse, and therefore, it is classified into four groups, depending on the monosaccharides expressed on the backbone.
(1) xyloglycan, (2) mannoglycan, (3) xyloglucan, and (4) mixed β-glucans. In turn, Xyloglycans divides into two groups (a) homoxylans and (b) heteroxylans. The latter is divided into five subgroups which are (i) glucuronoxylan, (ii) arabino(glucurono)xylan, (iii) Glucurono(arabino)xylan, (iv) arabinoxylan, and (v) complex heteroxylan, as shown in Figure 3 (van Zyl et al., 2010).
Xyloglycans are the most common hemicellulose found in hardwoods and herbaceous plants and constitute around 20% to 30% of secondary cell walls. It consists of β-(1,4) linked-D- xylopyranose molecules with carbohydrate side chains that vary between L-arabinose,
12 glucuronic acid (or its 4-O-methyl ether), and oligosaccharides such as D- or L-galactose, D- xylose, L-arabinose, and D-glucose (van Zyl et al., 2010).
Homoxylans are β-(1,3)- or β-(1,4)- linked D-xylopyranose, and its occurrence in nature is rare. Glucuronoxylan is mainly formed with xylose with side chains at 2-position of α-D- glucuronic acid or 4-O-methyl-D-glucuronic acid (Bajpai, 2014; Heinen et al., 2017).
The arabino(glucurono)xylan is the principal hemicellulose found in grasses and cereals and possesses a β-(1,4)-D-xylopyranose substituted backbone by 4-O-methyl-D-glucuronic acid and α-L-arabinofuranose (Bajpai, 2014; Heinen et al., 2017).
For glucurono(arabino)xylan, the main monomer of the backbone is arabinoxylan with few xylopyranose residues that are double substituted with α-L-arabinofuranose and uronic acid, and depending on the source, the number of substituents vary in terms of 4-O-methyl, xylose to arabinose ratio, and disaccharide side chains of α-L-arabinofuranose-(1,3)-α-L- arabinofuranose and β-D-xylopyranose-(1,2)-α-L-arabinofuranose (van Zyl et al., 2010).
Arabinoxylan is formed by a linear backbone of xylopyranose monomers with single or double substitutions of α-L-arabinofuranose residues. It is present in various cereals like barley, rice, rye, corn, or wheat, specifically located at the starchy endosperm and the outer layers of the kernel. Complex heteroxylans are (1,4)- β-D-xylopyranose backbone with arabinose and uronic acid residues, which also have various mono- and oligoglycosyl side chains (van Zyl et al., 2010)
There are two types of mannoglycans, galactomannans are constituted by (1,4)- linked D- mannopyranose and glucomannan, which are D-mannopyranose, and D-glucopyranose linked with β-(1,4) linkages. These two types can have a backbone branched in different degrees with D-galactopyranose residues. At last, xyloglucans are a β-(1,4)-D-glycopyranose backbone with side chains or D-xylopyranose, and the distribution of this side chain can occur in two forms, two glucopyranoses followed by two xylopyranoses or a single glucopyranose unit and three xylopyranoses (Hövel et al., 2003; Saha & Bothast, 1999).
For hemicellulose degradation, the most studied microorganism is Aspergillus niger.
Nonetheless, there are a variety of fungi that can degrade this plant cell wall polysaccharide.
In a study where 13 different fungi species were compared based on the carbohydrate-active enzymes contained in their genome, the most capable fungi for degrading hemicellulose where: Aspergillus nidulans, Aspergillus niger, Aspergillus oryzae, Penicillium chrysogenum, Fusarium graminearun, Neurospora crassa, Podospora anserina, Trichoderma reesei, Saccharomyces cerevisiae, Phanerochaete chrysosporium, and Schizophyllum commune (Bajpai, 2014). Also, in a screening study for hemicellulose- and cellulose-degrading enzymes in Ulocladium fungi species, it was demonstrated that most species of Ulocladium produce broad enzyme profiles capable of degrading hemicellulose (Bajpai, 2014).
13
Fig. 3. Schematic illustration of hemicellulose type molecules.
14
Fig. 4. Hemicellulose structures and enzymes.
15
2.1.3.1 Backbone degrading enzyme
Mannoglycan backbone degrading enzymes act in a systematic order where 1,4-β-D-mannan mannohydrolase (EC 3.2.1.78) is in charge of the mannan's hydrolysis backbone molecule by an endo-acting degradation of the internal glycosidic bonds to generate short β-1,4- manno-oligosaccharides (Aureli et al., 2018). These manno-oligosaccharides and mannobiose are hydrolyzed by 1,4-β-D-mannopyranoside hydrolase (EC 3.2.1.25) by the internal attack of linkages at the non-reducing ends to release mannose monomers (van Zyl et al., 2010).
Another enzyme that acts on mannose oligomers produced from the degradation of glucomannan and galactoglucomannan is 1,4-β-D-glucoside glucohydrolase (EC 3.2.1.21) through the removal of 1,4-glucopyranose units at the non-reducing ends to produce glucopyranose monomers (Bajpai, 2014; Tenkanen & Siika-Aho, 2000).
Xylan backbone degrading enzyme battery consists of endo-1,4-β-xylanase (EC 3.2.1.8) that is in charge of the cleavage of monosaccharides, disaccharides, and trisaccharides of β-D- xylopyranosyl, as well as xylopyranosyl oligomers through the hydrolysis of the β-1,4- glycosidic bonds (Szakacs et al., 2006). Then, the xylan-1,4-β-xylosidase (EC3.2.1.37) degrades those xylooligosaccharides and xylobiose by the successive remotion of D-xylose residues from non-reducing ends to produce xylose units (Heinen et al., 2017).
Finally, α-glucuronidase (EC 3.2.1.139) can attack either glucuronoxylans or glucuronoarabinoxylans backbone by hydrolysis of α-1,2-glycosidic bonds between xylose and glucuronic acid or 4-O-methyl-D-glucuronic acid (S. I. Mussatto et al., 2006).
2.1.3.2 Accessory enzymes
Mannoglycan accessory enzymes consist of 1,6-α-D-galactoside galactohydrolase (EC 3.2.1.22) that release the α-1,6-linked D-galactopyranosyl substituents from the galactomannan backbone (S. I. Mussatto et al., 2006). Acetyl mannan esterase (EC 3.1.1.6) is in charge of acetyl groups' remotion from the galactoglucomannan backbone (Pinheiro et al., 2019).
Arabinoxylan accessory enzymes are non-reducing end α-L-arabinofuranosidase (EC3.2.1.55), which is capable of also attacking rhamnogalacturonan I molecule (pectin) as well and is responsible for the cleavage of L-arabinose units at non-reducing ends by hydrolysis of the ester linkage between xylose or arabinose (Meneses et al., 2013). Feruloyl esterase (EC 3.1.1.73) is in charge of cleaving ester bonds between arabinose and ferulic acid side groups to release ferulic acid (Pinheiro et al., 2019).
ρ- coumaroyl esterase (EC 3.1.1.B10) cleaves ester bonds between arabinose and p-coumaric acid side groups to release p-coumaric acid (Pinheiro et al., 2019), and glucuronoarabinoxylan endo-1,4-β-xylanase (EC 3.2.1.136) is an enzyme produced by Bacillus licheniformis and is in charge of the hydrolysis of highly branched arabinoxylan
16 molecules into small oligomer containing arabinose and xylan (Solange I. Mussatto &
Roberto, 2006).
Glucuronoxylan accessory enzymes are xylan α-1,2-glucuronidase (EC 3.2.1.131) that is in charge of the hydrolysis of the α-1,2-linkages between 4-O-methyl glucuronic acid and xylose to liberate 4-O-methyl glucuronic acid from xylooligosaccharides and xylan polymers (Heinen et al., 2017), and acetylxylan esterase (EC 3.1.1.72), which cleave the ester linkages of the acetyl groups from β-D-xylopyranosul residues to produce acetyl units (Kanauchi et al., 2018).
2.1.4 Cellulose
Cellulose is the most abundant component among lignocellulose biomass. It represents around 30% to 60% of the total biomass (Szakacs et al., 2006) , and it is of significant interest for researchers and industries owing to its potential of becoming a source of renewable energy or chemical feedstock (Bhat, 2000), as it possesses in its structure around 500 to 1400 glucose molecules, depending on the degree of polymerization of the cellulose molecule and its source (Bhat, 2000).
Cellulose molecule is formed by linear chains of β-1,4-D glucans composed of cellobiose attached by β-1,4 glycosidic bonds. Each linear cellulose chain is oriented in parallel and depending on the degree of hydrogen bonds, cellulose forms highly ordered crystalline (not easily accessible for endo-acting enzymes and insoluble) cellulose or poorly arranged amorphous cellulose (easily accessible for enzymes), and the presence of these depends on the source(Szakacs et al., 2006).
Filamentous fungi represent the primary source for industrial application cellulases.
Nonetheless, some bacteria and marine organisms possess the ability to produce them. Some of the first studies pursued in fungi that produce cellulases were done in Trichoderma koningii, Penicillium funiculosum, Trichoderma viridie, Trichoderma reesei, Penicillium pinophilum, Fusarium solani, and Aspergillus niger. Since lignocellulose has gained more interest, other species have been classified as cellulases production microorganisms, such as Chaetomium thermophilum, Talaromyces emersonii, Neocallimastix frontalis, Bacillus spp., among others (Szakacs et al., 2006).
For a complete degradation of cellulose, both cellulose molecules (crystalline and amorphous) need to be fully degraded. For this, 1,4-β-D-glucan glucanohydrolases (EC 3.2.1.4) random cleave the β-1,4-glycosidic bonds within unsubstituted or substituted cellulose to produce glucose, cellobiose, or glucose polysaccharide chains (Bedford &
Partridge, 2010). Then cellulose 1,4-β-cellobiosidase (EC 3.2.1.91) cleaves the cellobiose from the non-reducing or reducing ends of polysaccharide chains produced by endoglucanase (EC 3.2.1.4) (Szakacs et al., 2006). Finally, β-D-glucoside glucohydrolases (EC 3.2.1.21) completes the hydrolysis by cleaving cellobiose and other glucose oligosaccharides to glucose (Rozeboom et al., 2013).
17 Another enzyme that can depolymerize cellulose through two-electron oxidation (FAD à FADH2) of the reducing-end of cello-oligosaccharides to produce cellobiose- 1,5-lactones, which are hydrolyzed spontaneously to cellobionic acid ((Gírio et al., 2010; Haghighi Mood et al., 2013).
Fig. 5. Schematic representation of cellulose enzymatic hydrolysis
2.2 Brewers spent grain and applications
Barley is the main constituent of the grain bill in most beer styles due to its starch, diastatic, and protein richness. Barley kernel consists of three parts: the embryo, the endosperm (aleurone and starch granules), and the grain covering consisting of four layers: the seed coat or testa, which covers the aleurone membrane, the fruit coat or pericarp, the epidermis, and the husk (Kunze, 2014 ) (Carvalheiro et al., 2004). After the mashing process, all starches and soluble proteins at the endosperm are extracted, leaving the insoluble proteins and the grain covering as the brewers' spent grain (BSG) main constituents that are rich in lignin, hemicellulose (arabinoxylan), cellulose, and many proteins and lipids. Also, some remaining starches could be present, depending on the mash step's extraction efficiency (Silva et al., 2004).
18 Brewer's spent grain is defined as the solid waste produced in breweries after completing the barley malt mashing process (ACERMEX, 2018). It is generated in a large amount (between 0.2 Kg per liter and 0.3 Kg/ liter), depending on the brewing recipe. In Mexico, the craft beer industry generates approximately 3.8 thousand tons per year of brewers spent grain, with an average growth rate of 53.29% of the industry, and the industrialized beer companies generate around 2.4 million tons per year of this waste (Xiros et al., 2008).
In a previous study by Pinheiro et al., (2019), six different brewer's spent grains batches from the same brewery were analyzed in terms of chemical composition (total glucan, soluble glucose polysaccharide, xylan, arabinan, total lignin, proteins, ashes, and extractives) to achieve high ethanol concentrations and yields during fermentations. The results showed that the lignocellulose constituents’ concentrations were glucans ranging from 26.5% to 32.1 %, lignin with values ranging from 8.9% to 14.5%, xylan ranging from 7.0% to 12.5%
(hemicellulose), arabinans ranging from 2.6% to 5.8 % (hemicellulose). To finish, extractable- soluble glucose polysaccharides with values as low as 2.2% and as high as 7.6%, which are expected in the non-converted starch fraction caused by the mashing process's low efficiency, a common drawback of craft brewing processes (Pinheiro et al., 2019).
In a research presented by Meneses et al., (2013)e subjected to acid hydrolysis pretreatment to recover hemicellulose and evaluate different process conditions (acid concentration, liquid-to-solid ratio) repercussion on the hemicellulose extraction rates. The substrate was provided from a local brewery (Lorena, Brazil) containing 100% barley malt. The brewers spent grain were washed with water at reception to eliminate residual sugars from the brewing process and dried at 50 ± 5 °C until 10% moisture was reached. The results showed that the brewers spent grain is composed of (g/100g of dry weight) lignin (27.8), hemicellulose (28.4), cellulose (16.8).
Fig. 6. Longitudinal cross-section of barley kernel.
19 Another study pursues by Xiros et al., (2008) evaluates the influence of a protein-rich and fibrous foodstuff produced with brewers spent grain, containing (% weight) lignin (20.1), hemicellulose (36.9), and cellulose (43.0), by converting % weight into g/100g dry weight, lignin (11.9), hemicellulose (21.8) and cellulose (25.4). The results indicate that the foodstuff improves defecation and prevented diarrhea and inflammation in colitis models.
Also, Solange I. Mussatto & Roberto, (2006) obtained brewers spent grain from a local craft brewery (Central de Cervejas, Vialong, Portugal) with approximately 80% moisture and treated it by drying at 50 °C to decrease moisture by about 10%. The pretreatment used correspond to autohydrolysis with water to enhance xylooligosaccharides with different temperatures and reaction times to evaluate its repercussion on extraction. The lignocellulose composition of the brewers spent grain consisted (g/ 100 g dry weight) of cellulose (21.9), xylan (20.6), arabinan (21.7), and lignin (24.6).
Brewers spent grain characterized by Kanauchi et al., 2018; S. I. Mussatto et al., (2006) consisted of (g/100g of dry weight) cellulose (25.3), hemicellulose (41.9), reported as a holocellulose (hemicellulose + cellulose), and lignin (16.9). The aim was to evaluate this waste as an adsorbent for acid orange seven, a dye used in paper and textile industries that causes environmental problems, resulting as a successful adsorbent for water dissolved dye.
Another application of brewers spent grain carried out by Carvalheiro et al., (2004), evaluating this substrate in terms of antioxidant phenolic compounds, demonstrating that it can be a valuable source. The substrate was supplied by UNICER bebidas de Portugal (S.mamede de infesta, Portugal) with 80% moisture, then dried at 60 °C to 10% moisture.
The composition consisted of (g/ 100gdry weight) cellulose (21.73), hemicellulose (19.27), xylans (13.63), arabinan (5.64), and lignin (19.40).
To finish, Silva et al., (2004) tested the ethanol production properties of Neurospora crassa with brewers spent grain as a substrate. The lignocellulosic composition consisted of (g /100g dray weight) cellulose (12.0), hemicellulose (40.2), and lignin (11.5). The composition of different brewers' spent grain is shown in Table 1.
The monomeric composition of brewers spent grain, the high percentage of the craft brewing industry's average growth rate - waste generation, the higher extractable-soluble glucose content (less spent grain), and its availability make this substrate a suitable candidate for further investigation and understanding lignocellulosic waste degradation as carried in nature by fungi.
20
Table 1. Brewers spent grain lignocellulose composition.
Component
[g /100gdry weight]
(Haghighi Mood et al., 2013)a
(Haghighi Mood et al., 2013)b
(Chang et al., 2011)
(Gírio et al., 2010)
(Gírio et al., 2010;
Haghighi Mood et al., 2013)
(Gírio et al., 2010; Haghighi
Mood et al., 2013)
(Gírio et al., 2010)
(Sadiq et al., 2011)
Cellulose 32.1 26.5 21.73 12.0 16.8 25.4 21.9 25.3
Hemicellulose 9.6 15.8 19.27 40.2 28.4 - 29.6 41.9
Xylan 7.0 10.6 13.63 - 19.9 - 20.6 -
Arabinan 2.6 5.2 5.64 - 8.5 21.8 9.0 -
Lignin 8.9 11.4 19.40 11.5 27.8 11.9 21.7 16.9
a data corresponding to BSG E b data corresponding to BSG A - not determined.
2.3 Successful mathematical models for enzymatic degradation of polymers.
A mathematical model is a tool to understand and structure a problem or phenomenon through its simplification and translation into an objective, punctual, and quantitative mathematical language that later can be challenged, fed, and proved through experimental work. Mathematical models are used to describe and answer a variety of phenomena. Herein, we describe some successful mathematical models for enzymatic degradation of heterogeneous polymers, principles, and results.
Ye & Berson, (2011) established a Michaelis-Menten based mathematical model to describe the concentrations of individual species (monomers or intermediate) produced by an enzyme degradation of a heteropolymer, focusing not just on the substrate concentration, but also on the product generation, as the products of one reaction are the substrates of another enzyme until completion of the degradation of all intermediate species. The model is compared with an in vitro HIV-1 protease-catalyzed Gag polyprotein for producing mature capsid proteins process, generating a mathematical model that accurately follows the time evolution and intermediate species formation in the Gag system.
21 Another mathematical model that describes the cellulose hydrolysis kinetic was developed by Y.-H. Percival Zhang, (2006) . A first-order cellulase's kinetic inactivation constant due to enzyme adsorption was considered through the Langmuir adsorption model, as it is one of the possible causes of slow kinetics and the rate reduction caused by this phenomenon. As a result, the comparison between the proposed model and the experimental data predicted 10%
of the experimental results, suggesting that Vmax decreases due to the inactivation.
Hydrolysis rate (33min-1) and inactivation rate (k=0.38h-1) were predicted within a 5%.
A more synergistic model is presented by Haghighi Mood et al., (2013), which includes two phenomena that cause gaps between mathematical models and experimental data. These are the degree of polymerization and the accessible bonds to cellulose hydrolysis in a Trichoderma cellulose-degrading model considering the action of three enzymes, cellobiohydrolase I, cellobiohydrolase II, and endoglucanase I. This results in a successful description of different enzymatic behaviors during hydrolysis in a variety of cellulosic substrates.
Up to date, many details based on the structure and function of enzymes and lignocellulosic biowaste are understood. Nonetheless, the interactions between substrate and enzymes, and the synergistic mechanism used by fungi in nature to degrade this substrate, are far to be fully comprehended. As shown above, the majority of existing mathematical models consider only cellulose degradation in a homogeneous substrate system, making evident the lack of mathematical models that study the synergism between the enzymes needed to degrade all lignocellulosic substrates and the interaction between them and all lignocellulose components. Therefore, we aim to develop a dynamic mechanistic mathematical model that considers a multi-enzyme complex to degrade all lignocellulose components that help understand the substrate, enzymes involved, and their interactions as well as the arrangement of the different polymer layers described in the previous sections.
2.4 Existing lignocellulose pretreatments.
Lignocellulose pretreatments' objective is to increase enzyme accessibility to allow efficient conversion of the lignocellulose polysaccharides into fermentable sugars. For lignocellulose, dependent processes (such as bioethanol and methane production), lignin and hemicellulose represent an impediment to obtaining considerable amounts of monomeric sugars. Therefore, a variety of methods are used to increase yields in terms of cellulose hydrolyzation.
Nonetheless, they can also produce inhibitors such as weak acids (levuliniacid, acetic acid, and formic acid), furan derivatives (5-hydroxy-2-methyl furfural), and furfural and phenolic compounds. Pretreatments classification consists of four groups that are physical, chemical, physicochemical, and biological pretreatments (Haghighi Mood et al., 2013), as shown in Figure 7.
22
Fig. 7. Lignocellulose biomass pretreatment
(1) Physical pretreatments aim to reduce the particle size to increase the contact surface area and decrease the degree of polymerization. These are subclassified as (a) Milling pretreatment, which consists of the use of mils (hammer milling, two-roll milling, disk milling, colloid milling, and ball milling) being the last one which reports better yields of extraction (Haghighi Mood et al., 2013).
(b) Extrusion pretreatment seeks physical and chemical alteration through mixing, heating, and shearing the lignocellulosic biomass (thermophysical). (c) Microwave pretreatment enables cellulose breakdown through the molecular collisions caused by a dielectric polarization, allowing selectivity, short process times, and less energy consumption (Gírio et al., 2010; Haghighi Mood et al., 2013). (d) Freeze pretreatment consists of freezing the sample for a specific time and then thawed at room temperature to produce structural changes in the lignocellulose material (Gírio et al., 2010; Haghighi Mood et al., 2013).
23 (2) Chemical pretreatments are classified into five subcategories: (a) Acid pretreatment is an acid-catalyzed process that allows the hydrolysis of lignocellulose into monosaccharides or oligosaccharides, allowing higher accessibility of cellulose-degrading enzymes. The acid concentration can be divided into high acid concentration/low temperature and low acid concentration/high temperature. The hydrolysis can occur to different extents, depending on the acid type and process conditions. The most common employed acids are hydrochloric acid, trifluoroacetic acid, sulfuric acid, and peracetic acid, being the last two the most utilized and the strongest (solubilize lignin), respectively (Haghighi Mood et al., 2013).
(b) Alkaline pretreatments are known to be a selective method as they allow the solubilizations of lignin, uronic acid, and acetyl substituents with medium impact on hemicellulose and low impact on cellulose hydrolysis (Haghighi Mood et al., 2013). The delignification is carried out by disrupting the ester bonds that crosslink the lignin and the hemicellulose.
(c) Ionic liquid pretreatment consisting of organic salts catalyzed process in which their melting point is usually below 100 °C conferring them with high thermal and chemical stability and low vapor pressure (Haghighi Mood et al., 2013). The organic salts can break the hydrogen bonding network of cellulose to solubilize it. It is effective against woods and crystalline structures of cellulose without solubilizing hemicellulose lignin (Haghighi Mood et al., 2013).
(d) The organosolv pretreatment utilize organic or inorganic acids as catalysts (such as hydrochloric and sulfuric acid) and solvents like ethanol, acetone, methanol, ethylene glycol, and tetrahydrofurfuryl alcohol. These, in combination, are capable of breaking internal linkages of lignocellulose to hydrolyze and solubilize hemicellulose and lignin (Haghighi Mood et al., 2013).
(e) Ozonolysis pretreatment breaks down the lignin and hemicellulose part of lignocellulose to increase cellulose biodegradation through an oxidation process carried by ozone gas; this pretreatment can release soluble compounds and formic acid and acetic acid molecules.
(3) Physicochemical pretreatments consist of (a) Steam explosion pretreatment, also considered a thermomechanochemical method. Consists of heating the lignocellulosic particles with pressurized steam (thermal forces); then, it is suddenly decompressed to atmospheric pressure (mechanical forces), and the moisture is evaporated (chemical forces), producing hydrolysis of glycosidic bonds of lignocellulose.
The hydrolysis of hemicellulose is accomplished by generating organic acids (such as acetic acid, formic acid, and levulinic acid) derived from substituent groups associated with hemicellulose molecule (Gírio et al., 2010; Haghighi Mood et al., 2013). (b) Ammonia fiber explosion pretreatment is similar to the steam explosion with the difference in liquid ammonia, which expands, causing the cleavage of lignin and the physical disruption of biomass fibers, opening the structure increasing the polymer surface area for a better enzyme digestibility (Gírio et al., 2010; Haghighi Mood et al., 2013).
24 (c) CO2 explosion pretreatment, also called supercritical CO2 explosion, is similar to steam and ammonia explosions. It is less costly, non-toxic, and needs lower temperatures than the others. CO2 possesses a small particle size (similar to water and ammonia), allowing it to penetrate small lignocellulose pores to produce hydrolysis (Haghighi Mood et al., 2013).
(d) Liquid hot water pretreatment (or Autohydrolysis) is an effective method for different lignocellulose types and can solubilize most hemicellulose but does not affect lignin and cellulose. It utilizes high temperatures and high pressures to avoid water evaporation and keeping it in contact with the substrate (Haghighi Mood et al., 2013). (e) Wet oxidation involves water and oxygen/air at high temperature and pressure. These conditions promote lignin solubilization and hemicellulose hydrolyzation through organic acids and oxidative reactions generated during the process (Pinheiro et al., 2019).
(4) Biological pretreatment consists of an enzymatic saccharification process in which microorganisms, especially fungi, are used to convert lignocellulose biowaste into a hydrolyzed substrate. The most common fungi employed in this pretreatment consist of white-, brown-, and soft-fungi, the white-rot fungi Phanerochaete chrysosporium, the one with the best-reported efficiencies to degrade lignocellulose. This technique's major drawback relies on the long process time, constant monitoring of microorganism growth, high-technology requirements, and the most important, the poor understanding of the fungi enzymatic mechanism (Pinheiro et al., 2019).
25
Chapter III
3 Materials and methods
26
3.1 Mathematical model
3.1.1 Conception of the model
Before running experiments, it was decided to create a mathematical as a first approach that was further recreated by using experimental data. This first approach was made considering all the information included in the literature regarding how the lignocellulosic residue is composed (Figure 1). Furthermore, to establish the mathematical model's principle, the following considerations were made:
(1) The external layer corresponds to lignin, a phenolic compound covering all the plants' cell wall polysaccharides. The second layer is hemicellulose and pectin, composed of high-interest monosaccharides such as D-xylose, D-glucose, D-galacturonic acid, and L-arabinose. Finally, the cellulose and starch layer, formed by cellobiose and D- glucose (the starches comes from a deficient mash process at the brewery) (Ye &
Berson, 2011)
(2) To initiate correct degradation of brewers spent grain, it is necessary to oxidize the most external layer that is lignin (Lynch et al., 2016; Solange I. Mussatto, 2014;
Pinheiro et al., 2019) through the action of laccase. Then degrade the hemicellulose and pectin layers by the action of pectinases and xylanases. Finally, degrade the cellulose and starch layers through the action of pectinases and xylanases.
(3) The remaining reaction products consist of reducing sugar.
Based on these considerations, and the structure from Figure1, an abstraction of the lignocellulose molecule was produced to generate a visual reference for the establishment of stoichiometric equations and the differential equations. The abstraction is shown in Figure 8.
Fig. 8. Mathematical model abstraction showing how polymer layers are arranged according to literature.
27 Figure 8 shows a hypothetical simplification of the composition of the brewer’s spent grain in terms of lignin, hemicellulose, pectin, cellulose, and starch. This abstraction is presented in a layer model, in which the degradation pattern must occur from the outer layers to the inner layer. According to this, the stoichiometric equations, which describe the enzymatic degradation and exposure of each subsequent layer were established as follow:
𝑟1.(𝑎)𝐿𝑖𝑔𝑛𝑖𝑛 𝐿𝑎𝑐𝑐𝑎𝑠𝑒 → (𝑏)𝐻𝑒𝑚𝑖𝑐𝑒𝑙𝑙𝑢𝑙𝑜𝑠𝑒 +(𝑐)𝑃𝑒𝑐𝑡𝑖𝑛 +(𝑓)𝑃ℎ𝑒𝑛𝑜𝑙𝑠
𝑟2.(𝑏)𝐻𝑒𝑚𝑖𝑐𝑒𝑙𝑙𝑢𝑙𝑜𝑠𝑒 𝑋𝑦𝑙𝑎𝑛𝑎𝑠𝑒 → (𝑑)𝐶𝑒𝑙𝑙𝑢𝑙𝑜𝑠𝑒 +(𝑔)𝑋𝑦𝑙𝑎𝑛 +(𝑛)𝑆𝑡𝑎𝑐ℎ +(ℎ)𝐴𝑟𝑎𝑏𝑖𝑛𝑜𝑠𝑒 𝑟3.(𝑐)𝑃𝑒𝑐𝑡𝑖𝑛 𝑃𝑒𝑐𝑡𝑖𝑛𝑎𝑠𝑒
→ (𝑑)𝐶𝑒𝑙𝑙𝑢𝑙𝑜𝑠𝑒 +(𝑔)𝑋𝑦𝑙𝑎𝑛 +(𝑛)𝑆𝑡𝑎𝑟𝑐ℎ +(𝑖)𝐺𝑎𝑙𝑎𝑐𝑡𝑢𝑟𝑜𝑛𝑖𝑐 𝐴𝑐𝑖𝑑
𝑟4.(𝑔)𝑋𝑦𝑙𝑎𝑛 𝑋𝑦𝑙𝑎𝑛𝑎𝑠𝑒 → (𝑗)𝑋𝑦𝑙𝑜𝑠𝑒 𝑟5.(𝑛) 𝑆𝑡𝑎𝑟𝑐ℎ 𝐴𝑙𝑝ℎ𝑎−𝑎𝑚𝑦𝑙𝑎𝑠𝑒
→ (𝑚)𝐺𝑙𝑢𝑐𝑜𝑠𝑒 𝑟6.(𝑑)𝐶𝑒𝑙𝑙𝑢𝑙𝑜𝑠𝑒 𝐶𝑒𝑙𝑙𝑢𝑙𝑎𝑠𝑒 → (𝑙)𝐶𝑒𝑙𝑙𝑜𝑏𝑖𝑜𝑠𝑒
3.1.2 Differential equations
The differential equations were then established based on the degradation of each component and the stoichiometric coefficient, which are related to the change of concentration of lignin, hemicellulose, and cellulose over the time. Initial concentration for each polymer was expressed according to Table 1. The differential equations were set as follow:
𝑑[𝐿𝑖𝑔𝑛𝑖𝑛]
𝑑𝑡 = −(𝑎)𝑟1 (1)
𝑑[𝐻𝑒𝑚𝑖𝑐𝑒𝑙𝑙𝑢𝑙𝑜𝑠𝑒]
𝑑𝑡 = (𝑏)𝑟1 − (𝑏)𝑟2 (2)
𝑑[𝑋𝑦𝑙𝑎𝑛]
𝑑𝑡 = (𝑔)𝑟2 + (𝑔)𝑟3 − (𝑔)𝑟4 (3)
𝑑[𝑆𝑡𝑎𝑟𝑐ℎ]
𝑑𝑡 = (𝑛)𝑟2 + (𝑛)𝑟3 − (𝑛)𝑟5 (4)
𝑑[𝐶𝑒𝑙𝑙𝑢𝑙𝑜𝑠𝑒]
𝑑𝑡 = (𝑑)𝑟2 + (𝑑)𝑟3 − (𝑑)𝑟6 (5)
𝑑[𝑋𝑦𝑙𝑜𝑠𝑒]
𝑑𝑡 = (𝑗)𝑟4 (6)
𝑑[𝑋𝑦𝑙𝑜𝑠𝑒]
𝑑𝑡 = (𝑗)𝑟4 (7)
𝑑[𝐺𝑎𝑙𝑎𝑐𝑡𝑢𝑟𝑜𝑛𝑖𝑐 𝑎𝑐𝑖𝑑]
𝑑𝑡 = (𝑖)𝑟3 (8)
𝑑[𝐺𝑙𝑢𝑐𝑜𝑠𝑒]
𝑑𝑡 = (𝑚)𝑟5 (9)
𝑑[𝐶𝑒𝑙𝑙𝑜𝑏𝑖𝑜𝑠𝑒]
𝑑𝑡 = (𝑙)𝑟6 (10)
𝑑[𝑃ℎ𝑒𝑛𝑜𝑙𝑠]
𝑑𝑡 = (𝑓)𝑟1 (11)
28 The enzyme reactions rates were defined according to the Michaelis-Menten equation, in which the reaction rate (r0) depends on the initial enzyme concentration [E]0, the catalytic turnover (kcat), and the initial substrate concentration [S]0 divided by the Michaelis-Menten constant (KM), the half-saturation constant, and initial substrate concentration [S]0.
𝑟0 = ([𝐸]0𝑘𝑐𝑎𝑡)[𝑆]0
𝐾𝑀+ [𝑆]0 (12)
The ordinary differential equations were substituted in equation (12), derived, and solved using Mathematica software (Wolfram Research, Inc. Mathematica, Version 12.1, 2020) for substrate concentration [𝑆]0, and plotted to obtain a graph that expresses the substrate concentration (consumed or produced) trough time.
3.2 Experimental section
3.2.1 DNS Reactant preparation
In a 4000 ml beaker, 800 ml of distilled water was placed, then 10.6 g of DNS (3,5- dinitrosalycilic acid, Sigma Aldrich, D0550-500G), 19.8 g of NaOH (CTR- Scientific,CTR03108), 306 g of Sodium potassium tartrate, 4-hydrate (J.T.Baker, 3262-01), 8.132 g of phenol crystals (High Purity, F1020), and 8.3 g of sodium metabisulfite (CTR- Scientific,CTR-01541) were weighed and added under agitation with a magnetic stirrer at 200 rpm (IKA RCT). Once homogenized, the rest of the water is added, and the solution is distributed into two amber bottles until its use.
3.2.2 100 mM Acetate buffer pH 5.0
A volume of 0.427 ml of acetic acid (Sigma Aldrich, 320099-2.5L) was added into a 2000 ml volumetric flask containing a small volume of distilled water (500 ml). Then, 1.443 g of sodium acetate (High Purity, A1160) were added to the volumetric flask and stirred vigorously. Once the suspension was homogenized, the rest of the distilled water was added, and the pH was adjusted at 5.0 with a potentiometer (Hanna checker HI98103, Hanna Instruments).
