UNIVERSIDADE ESTADUAL PAULISTA “JÚLIO DE MESQUITA FILHO” INSTITUTO DE PESQUISA EM BIOENERGIA unesp PROGRAMA INTEGRADO (UNESP, USP E UNICAMP) DE PÓS-GRADUAÇÃO EM BIOENERGIA CELLOOLIGOSACCHARIDES AND XYLOOLIGOSACCHARIDES PRODUCTION FROM SUGARCANE BAGASSE AND COTTON RESIDUES USING CHEMICAL, PHYSICAL AND BIOLOGICAL APPROACHES JEFFERSON POLES FELIPUCI Tese apresentada ao Instituto de Pesquisa em Bioenergia de Rio Claro, Universidade Estadual Paulista, como parte dos requisitos para obtenção do título de Doutor em Bioenergia. Orientador(a): Prof. Dr. Michel Brienzo Coorientadores: Prof. Dr. Fernando Masarin e Profa. Dra. Rosana Goldbeck Novembro - 2024 PROGRAMA INTEGRADO (UNESP, UPS E UNICAMP) DE PÓS-GRADUAÇÃO EM BIOENERGIA CELLOOLIGOSACCHARIDES AND XYLOOLIGOSACCHARIDES PRODUCTION FROM SUGARCANE BAGASSE AND COTTON RESIDUES USING CHEMICAL, PHYSICAL AND BIOLOGICAL APPROACHES JEFFERSON POLES FELIPUCI Tese apresentada ao Instituto de Pesquisa em Bioenergia de Rio Claro, Universidade Estadual Paulista, como parte dos requisitos para obtenção do título de Doutor em Bioenergia. Orientador: Prof. Dr. Michel Brienzo Coorientadores: Prof. Dr. Fernando Masarin e Profa. Dra. Rosana Goldbeck Novembro - 2024 F315c Felipuci, Jefferson Poles Cellooligosaccharides and xylooligosaccharides production from sugarcane bagasse and cotton residues using chemical, physical and biological approaches / Jefferson Poles Felipuci. -- Rio Claro, 2024 127 p. : il., tabs., fotos Tese (doutorado) - Universidade Estadual Paulista (UNESP), Instituto de Pesquisa em Bioenergia, Rio Claro Orientador: Michel Brienzo Coorientador: Fernando Masarin, Rosana Goldbeck 1. Bagaço de cana. 2. Algodão. 3. Enzimas. 4. sacarídeos. I. Título. Sistema de geração automática de fichas catalográficas da Unesp. Dados fornecidos pelo autor(a). UNIVERSIDADE ESTADUAL PAULISTA Unidade Complementar - Rio Claro Instituto de Pesquisa em Bioenergia - Unidade Complementar - Rio Claro - Rua 10, 2527, 13500230 www.ipben.unesp.br CERTIFICADO DE APROVAÇÃO TÍTULO DA TESE: Cellooligosaccharides and xylooligosaccharides production from sugarcane bagasse and cotton residues using chemical, physical and biological approaches AUTOR: JEFFERSON POLES FELIPUCI ORIENTADOR: MICHEL BRIENZO COORIENTADOR: FERNANDO MASARIN COORIENTADORA: ROSANA GOLDBECK COELHO Aprovado como parte das exigências para obtenção do Título de Doutor em Bioenergia, área: Bioenergia pela Comissão Examinadora: Prof. Dr. MICHEL BRIENZO (Participaçao Presencial) Laboratorio de Caracterizacao de Biomassa / Instituto de Pesquisa em Bioenergia IPBEN Dr. FERNANDO ROBERTO PAZ CEDEÑO (Participaçao Virtual) Dale Bumpers College of Agricultural, Food and Life Sciences / Pós-doutorando - University of Arkansas (UARK, EUA) Profa. Dra. THAIS SUZANE MILESSI ESTEVES (Participaçao Virtual) Departamento de Engenharia Química / Universidade Federal de São Carlos - UFSCar Profa. Dra. PATRÍCIA FELIX ÁVILA (Participaçao Presencial) Faculdade de Engenharia de Alimentos / Universidade Estadual de Campinas - Unicamp Rio Claro, 28 de novembro de 2024 AGRADECIMENTOS Primeiramente, agradeço aos meus pais Sônia Maria Poles e Jair Felipuci e à minha esposa Mayara Mulato dos Santos por todo o apoio e amor durante a minha caminhada. Ao meu orientador Michel Brienzo pelos ensinamentos e por ter sido o melhor orientador que eu poderia pedir. Muito obrigado! Aos meus colegas de laboratório Carol, Rogério, Rosângela, Rodrigo e Hernán pela amizade e pela companhia no laboratório. À Danieli, à Nathália e novamente ao Hernán, pela imensa ajuda no último ano. À Luiza pela oportunidade de ser coorientador. À professora Derlene Attili de Angelis e ao professor Fernando Masarin pela coorientação e pela ajuda. À UNESP e ao IPBEN O presente trabalho foi realizado com apoio da Coordenação de Aperfeiçoamento de Pessoal de Nível Superior - Brasil (CAPES) - Código de Financiamento 001 ABSTRACT Cellooligosaccharides (COS) and xylooligosaccharides (XOS) are oligomers with interest in several areas, such as agriculture and food industry. The production of COS and XOS depends on the pretreatment involved in the biomass. This study was dedicated to COS and XOS production from two different biomasses: sugarcane bagasse and cotton residue. The methods involved chemical, physical and biological pretreatment, and a combination of them. Also, for both studies, x-ray, FTIR-ATR and scanning electron microscope (SEM) analysis were performed. For sugarcane bagasse, the objective was the production of COS and XOS by a combination of physical and biological pretreatments, while for cotton residue the objective was the production of COS using chemical, physical and biological pretreatments, individually. The sugarcane bagasse was biologically pretreated for 5 months with Coniophora puteana (CBMAI 0870), Gloeophyllum trabeum (CBMAI 0872), and Pleurotus ostreatus (CCIBt 2338). A part of the biomass was ball-milled, and the other part was knife-milled, then the enzymatic hydrolysis with cellulase and xylanase was applied in different enzyme loads, ranging from 20 to 50 IU/g. After five months of pretreatment with C. puteana, followed by enzymatic hydrolysis using cellulase at 50 IU/g in combination with knife milling, the yield of COS reached 26.23%. In contrast, when the same pretreatment was applied but with ball milling instead, the yield of COS increased to 36.65%. The best XOS result was 78.12% yield after the biological pretreatment with G trabeum after 1 month of pretreatment. For the cotton residue, NaOH was used with different concentrations and temperature, ball milling with three different times of reactions and biological pretreatment using Gloeophyllum trabeum (CBMAI 0872) growth for 15 days in liquid medium. The COS yield reached 8.68% from untreated cotton using 100 IU/g of enzyme, against 11.61% of COS which was alkali pretreated using 50% NaOH (w/w) at 100 ºC. The biological pretreatment reached 9.13% of COS yield using the same enzyme load. The ball milling method obtained the best COS yield, reaching 30.11% after 6 h of milling. For both studies, the x-ray, FTIR-ATR and SEM analysis showed differences in the biomasses in comparison to the untreated ones, mainly after the ball milling pretreatment, decreasing drastically the crystallinity index, from 65.38% to 47.85% after 3 h of ball milling. This study examines the effects of chemical, physical, and biological pretreatments on sugarcane bagasse and cotton in the production of COS and XOS. Keywords: oligosaccharides, biomass, sugarcane bagasse, cotton, enzymatic hydrolysis. RESUMO Celo-oligosacarídeos (COS) e xilo-ogosacarídeos (XOS) são oligômeros com interesse em diversas áreas, como a agricultura e a indústria alimentícia. A produção de COS e XOS depende do pré- tratamento envolvido na biomassa. Este estudo foi dedicado à produção de COS e XOS a partir de duas biomassas diferentes: bagaço de cana-de-açúcar e resíduo de algodão. Os métodos envolveram pré-tratamento químico, físico e biológico, e uma combinação deles. Também, para ambos os estudos, foram realizadas análises por raios-x, FTIR-ATR e microscópio eletrônico de varredura (MEV). Para o bagaço de cana, o objetivo foi a produção de COS e XOS por combinação de pré-tratamentos físicos e biológicos, enquanto que para o resíduo de algodão o objetivo foi a produção de COS utilizando pré-tratamentos químicos, físicos e biológicos, individualmente. O bagaço de cana foi pré-tratado biologicamente por 5 meses com Coniophora puteana (CBMAI 0870), Gloeophyllum trabeum (CBMAI 0872) e Pleurotus ostreatus (CCIBt 2338). Uma parte da biomassa foi moída em esferas e a outra parte foi moída com faca, então a hidrólise enzimática com celulase e xilanase foi aplicada em diferentes cargas de enzimas. Após cinco meses de pré- tratamento com C. puteana, seguido de hidrólise enzimática utilizando celulase a 50 UI/g em combinação com moagem com faca, o rendimento de COS atingiu 26,23%. Em contraste, quando o mesmo pré-tratamento foi aplicado, mas com moagem de esferas, o rendimento de COS aumentou para 36,65%. O melhor resultado do XOS foi 78,12% de rendimento após o pré- tratamento biológico com G. trabeum após 1 mês de pré-tratamento. Para o resíduo de algodão, foi utilizado NaOH com diferentes concentrações e temperaturas, moagem em esferas com três tempos diferentes de reações e pré-tratamento biológico utilizando crescimento de Gloeophyllum trabeum (CBMAI 0872) por 15 dias em meio líquido. O rendimento de COS atingiu 8,68% a partir do algodão não tratado com 100 UI/g de enzima, contra 11,61% de COS que foi alcalino pré-tratado com 50% de NaOH (m/m) a 100 ºC. O pré-tratamento biológico atingiu 9,13% do rendimento de COS utilizando a mesma quantidade de enzima. Além disso, o método de moagem por esferas obteve o melhor rendimento de COS, atingindo 30,11% após 6 h de moagem. Para ambos os estudos, as análises de raios-x, FTIR-ATR e MEV mostraram diferenças nas biomassas em comparação com as não tratadas, principalmente após o pré-tratamento da moagem de esferas, diminuindo drasticamente o índice de cristalinidade, de 65,38% do índice de cristalinidade da amostra não tratada para 47,85% após 3 h de moagem de esferas. Este estudo examina os efeitos de pré-tratamentos químicos, físicos e biológicos em bagaço de cana-de-açúcar e algodão na produção de COS e XOS. Palavras-chave: oligossacarídeos, biomassa, bagaço de cana-de-açúcar, algodão, hidrólise enzimática FIGURES LIST Figure 1.1 - Schematic representation of lignocellulosic biomass emphasizing the cellulose macromolecule……………………………………………………………………………………………..25 Figure 1.2 - Scanning electron micrographs of beech wood degradation by white-rot fungi after 120 days…………………………………………………………………………………………..………...28 Figure 1.3 - Simplified representation of lignocellulolytic enzymes and their action mode………………………………………………………………………………………………...………30 Figure 1.4 - Scanning electron microscope images on the surface of the Oil palm Empty Fruit Bunch…………………………………………………………………………………………………….….40 Figure 1.5 - Proposed process of degradation of the wheat straw cell wall by Phanerochaete chrysosporium………………………………………………………………………...……………………41 Figure 2.1 - A: ATR spectra of untreated and biologically pretreated with C. puteana (CBMAI 0870) bagasse samples; B: ATR spectra of untreated and biologically pretreated with G. trabeum (CBMAI 0872) bagasse samples. C: ATR spectra of untreated and biologically pretreated with P. ostreatus (CCIBt 2338) bagasse samples………………………………………………………………………………………………...…...73 Figure 2.2 - Scanning electron microscope image from the untreated and biologically pretreated sugarcane bagasse. The yellow arrows show the gaps formed by the fungi growth (G. trabeum 2nd month, G. trabeum 3rd month, C. puteana 1st month, C. puteana 2nd month, P. ostreatus 2nd month, P. ostreatus 3rd month and P. ostreatus 4th month) and the structural modifications in the lignocellulosic biomass (G. trabeum 1st month, C. puteana 2nd month, C. puteana 3rd month, P. ostreatus 2nd month, P. ostreatus 3rd month and P. ostreatus 4th month)……………………………………………………………………………………………………….75 Figure 2.3 - X-ray diffraction from sugarcane bagasse after biological pretreatment and ball milling……………………………………………………………………………………………………..…77 Figure 3.1 - X-ray diffraction from cotton residues comparing the untreated one and after pretreatments……………………………………………………………………………………………..100 Figure 3.2 - FTIR-ATR spectra of untreated and pretreated samples…………………………..….102 Figure 3.3 - SEM images from unetreated and pretreated cotton…………………………………..104 Figure 4 - Book published in 2020………………………………………………………………..…….112 Figure 5 - Book chapter published in 2020……………………………………………………..……..113 Figure 6 - Paper published in 2024…………………………….………………………………………114 TABLE LIST Table 1.1 - Properties of cellulases and hemicellulases action on lignocellulosic biomass…………………………………………………………………………………………………......37 Table 2.1 - Chemical composition of untreated and biologically pretreated sugarcane bagasse material…………………………………………………………………………………………………...…66 Table 2.2 - COS conversion (%) after enzymatic hydrolysis of biologically pretreated sugarcane bagasse using three different enzymatic charges of cellulase………………………………………..68 Table 2.3 - COS conversion (%) after enzymatic hydrolysis of ball-milled biologically pretreated sugarcane bagasse using cellulase…………………………………………………………………..….70 Table 2.4 - XOS production after enzymatic hydrolysis of untreated and biologically pretreated sugarcane bagasse…………………………………………………………………………………..……71 Table 2.5 - ATR absorbance wavebands in lignocellulosic biomass………………………………….72 Table 3.1 - Chemical composition of untreated and pretreated cotton residue……………………..93 Table 3.2 - Glucose, C2 and >C3 yield after enzymatic hydrolysis using cellulase………………..97 Table 3.3 - Total COS yield after enzymatic hydrolysis using cellulase at 20, 50 and 100 IU/g…………………………………………………………………………………………………..……...98 Table 3.4 - ATR absorbance wavebands in lignocellulosic biomass……………………………….101 Attachment Table 2.1 - ANOVA test for chemical characterization comparison between samples. G. trabeum (Gt); C. puteana (Cp); P. ostreatus (Po)……………………………………………..………115 Attachment Table 2.2 - Yield of glucose and COS (%) after enzymatic hydrolysis of biologically pretreated sugarcane bagasse using three different enzymatic charges of cellulase……………………………………………………………………………...…………………….117 Attachment Table 2.3 - ANOVA test for enzymatic hydrolysis (cellulase) comparison between samples for each enzymatic charge (20, 50 and 100 IU/g). G. trabeum (Gt); C. puteana (Cp); P. ostreatus (Po)……………………………………………………………………………..………………118 Attachment Table 2.4 - ANOVA test for enzymatic hydrolysis (cellulase) comparison between samples for 50 IU/g of enzymatic charge for ball milling method. G. trabeum (Gt); C. puteana (Cp); P. ostreatus (Po)…………………………………………………………………………………………..120 Attachment Table 2.5 - ANOVA test for enzymatic hydrolysis (xylanase) comparison between samples for 50 IU/g of enzymatic charge. G. trabeum (Gt); C. puteana (Cp); P. ostreatus (Po)……………………………………………………………………………………………….………..121 Attachment Table 3.1 - ANOVA test for chemical composition comparison between samples………………………………………………………………………………………...………….122 Attachment Table 3.2 - ANOVA test for glucose using 20, 50 and 100 IU/g of enzyme charge……………………………………………………………………………………………...………123 Attachment Table 3.3 - ANOVA test for C2 using 20, 50 and 100 IU/g of enzyme charge………………………………………………...……………………………………………………124 Attachment Table 3.4 - ANOVA test for >C3 using 20, 50 and 100 IU/g of enzyme charge……………………………………………………………………………………………...………125 Attachment Table 3.5 - ANOVA test for enzyme charge comparison used for COS production………………………………………………………………………………………...……….126 Attachment Table 3.6 - ANOVA test for pretreatment comparison used for COS production…………………………………………………………………………………………...…….127 TABLE OF CONTENTS THESIS INTRODUCTION……………………………………………………………..……………….…12 OBJECTIVES…………………………………………………………………………………………..…..13 Specific Objectives…………………………………………………………………..…………………..14 Thesis format and organization……………………………………………………….......................14 CHAPTER I: BIOTECHNOLOGICAL ASPECTS OF MICROBIAL PRETREATMENT OF LIGNOCELLULOSIC BIOMASS……………………………………………………………………..….18 ABSTRACT…………………………………………………………………………………………...……19 1.1 INTRODUCTION…………………………………………………………………………………...….20 1.2 BIOLOGICAL PRETREATMENT…………………………………………………………………....21 1.2.1 Lignocellulosic Biomass structure……………………………………………………………..23 1.2.2 Microorganisms in Biological pretreatments………………………………………………...26 1.2.2.1 White-Rot Fungi………………………………………………………………………………...…27 1.2.2.2. Brown-Rot Fungi………………………………………………………………………..………..28 1.2.2.3. Bacteria………………………………………………………………………………………...….29 1.2.3 Enzymes Involved in Biological Pretreatment…………………………………………..…...30 1.2.3.1 Cellulases……………………………………………………………………………………….....31 1.2.3.2 Hemicellulases……………………………………………………………………………..……..32 1.2.3.3 Ligninases……………………………………………………………………………………..…..34 1.2.4. Enzymatic Hydrolysis of Biological Pretreated Material………………………………..…35 1.2.5 Mechanisms of Cell Wall Degradation by Microorganisms……………...………………...39 1.3. ECONOMIC IMPACTS AND CHALLENGES ON INDUSTRIAL SCALE INVOLVING BIOLOGICAL PRETREATMENT………………………………………………….…………………….42 1.4. CONCLUDING REMARKS……………………………………………………………………...…..44 CHAPTER II: EFFICIENT PRODUCTION OF CELLOOLIGOSACCHARIDES AND XYLOOLIGOSACCHARIDES BY COMBINED BIOLOGICAL PRETREATMENT AND ENZYMATIC HYDROLYSIS PROCESS………………………………………………………………...57 ABSTRACT……………………………………………………………………………………...…………58 2.1 INTRODUCTION………………………………………………………………………………...…….59 2.2 MATERIALS AND METHODS……………………………………………………………………….60 2.2.1 Material and bagasse preparation………………………………………..…………………….60 2.2.2 Biological pretreatment……………………………………………………………………..……61 2.2.3 Chemical characterization………………………………………………………………………..61 2.2.4 Enzymatic hydrolysis………………………………………………………………………...…...62 2.2.5 Analysis of ATR‑FTIR……………………………………………………………………...………63 2.2.6 Biomass images from electron microscope………………………………………………….63 2.2.7 X‑ray analysis………………………………………………………………………………………63 2.2.8 Statistical analysis…………………………………………………………………………………64 2.3 RESULTS AND DISCUSSION……………………………………………………………………….64 2.3.1 Chemical characterization………………………………………………………………………..64 2.3.2 Enzymatic hydrolysis……………………………………………………………………...……...66 2.3.3 FTIR‑ATR analysis…………………………………………………………………………...…….71 2.3.4 Scanning electron microscope images………………………………………………………..73 2.3.5 X‑ray analysis……………………………………………………………………………………....76 2.4 CONCLUSIONS…………………………………………………………………………………….....77 CHAPTER III: CELLOOLIGOSACCHARIDES PRODUCTION FROM COTTON RESIDUE USING BIOLOGICAL PRETREATMENT, ALKALI AND MILLING APPROACHES………………………..84 ABSTRACT………………………………………………………………………………………………...85 3.1 INTRODUCTION………………………………………………………………………………...…….86 3.2 MATERIAL AND METHODS…………………………………………………………………………87 3.2.1 Material and cotton preparation…………………………………………………………………87 3.2.2 Ball milling………………………………………………………………..…………………………88 3.2.3 Alkali pretreatment………………………………………………………………………………...88 3.2.4 Biological pretreatment…………………………………………………………………………..88 3.2.5 Chemical characterization………………………………………..………………………………89 3.2.6 Enzymatic hydrolysis ………………………………………..…………………………………...89 3.2.7 X-ray analysis………………………………………..…………………………………………......90 3.2.8 FTIR-ATR analysis………………………………………..………………………........................90 3.2.9 Scanning electron microscope images ……………………………………………………….90 3.2.10 Statistical analysis………………………………………..……………………………………...90 3.3 RESULTS AND DISCUSSION………………………………………..…………………………......91 3.3.1 Chemical characterization………………………………………..………………………………91 3.3.2 COS yield after enzymatic hydrolysis……………………………………………………...…..93 3.3.3 - X-ray analysis………………………………………..……………………………………..…….98 3.3.4 FTIR-ATR analysis………………………………………..…………………………...………….100 3.3.5 Scanning electron microscope (SEM) images………………………………………………102 3.4 CONCLUSIONS………………………………………..…………………………………………….106 THESIS GENERAL CONCLUSION AND FUTURE PERSPECTIVE……………………………….111 ATTACHMENT………………………………………..……………………………………..…………...112 12 THESIS INTRODUCTION Biomass is a biological material that can be converted into energy or high added value products. Sugarcane bagasse and cotton residues are examples of biomasses that can be used to produce cellooligosaccharides and xylooligosaccharides, or second generation ethanol, in the sugarcane bagasse case. The potential of biomasses can be an ideal substitute for fossil fuels in energy industries, once the origin of the biomasses are natural resources. Bio-based products have been used over the years as a replacer or a diminisher for oil-based products, they are present in products such as engines, packaging, medicines, and others. The bioproducts can be originated from vegetal biomass like crops, vegetable oils, forests, and waste from agriculture, cities and industry (SCAPINI, et al., 2021; QASEEM, SHAHEEN, WU, 2021). Before being turned into a bioproduct, the biomass needs to pass through a process called pretreatment, which consists of modifying the lignocellulosic structure aiming to expose the macromolecules in it or reduce the recalcitrance. Pretreatments can be separated into chemical, physical and biological pretreatment, and a combination of them (FELIPUCI, et al., 2020). Chemical pretreatments consist of the use of acid or alkali reagents to modify the biomass structure: acid conditions solubilize hemicellulose, while alkali conditions change the lignin structure, reduce the cellulose crystallinity and partially solubilize the hemicellulose and lignin. Physical pretreatments consist of use of a physique condition, such as milling, reducing the lignocellulosic particle size and crystallinity. Usually, chemical and physical pretreatments are combined to reach a better result. However, chemical pretreatment usually generates undesirable compounds and demands much energy (FELIPUCI, et al., 2020; 2021). There are also the physic-chemic pretreatments, such as steam explosion, consisting in a saturated steam under a controlled pressure and temperature. Biological pretreatment is the use of microorganisms to degrade or modify vegetal biomass structure employing their special enzymatic complexes (IOELOVICH, 2016). Biological pretreatment is already known as a sustainable option to conventional methods used in industries due to does not generate inhibitory and toxic compounds and is a cheaper method compared to chemical and physical pretreatments. Biological pretreatment can be combined with chemical and physical methods to reach better results using less chemicals and energy, making this method an economically and an eco-friendly feasible process 13 (FELIPUCI, et al., 2020; IOELOVICH, 2016; AGBOR, et al., 2011). Physical pretreatments consist in use techniques to reduce the size of the biomass particles, such as knife milling or ball milling, however the physical pretreatment is known to demand more energy in the process, but is faster and don’t generate undesirable compounds (FELIPUCI et al, 2020 and 2021; RASTOGI, SHRIVASTAVA, 2017). The lignocellulosic material is mainly composed of cellulose, hemicellulose and lignin, which are strictly connected, and this connection makes the material resistant to pretreatments. This is responsible for the high recalcitrance of the material. Microorganisms have the capacity to produce specific enzymes that break the lignocellulosic structure and decrease the recalcitrance. Combining methods focusing on decreasing the material recalcitrance and breaking the lignocellulosic structure, then applying enzymatic hydrolysis (carbohydrate hydrolysis), can generate oligomers from cellulose and xylan with more value-added, such as cellooligosaccharides (COS) and xylooligosaccharides (XOS), respectively (FELIPUCI, et al., 2021; SINDHU, BINOD, PANDEY, 2016; RASTOGI, SHRIVASTAVA, 2017). COS are oligomers formed by β-1,4 linked D-glucose units with a few numbers of glucose. COS are not digestible and can show prebiotic effect, stimulating in vitro growth of healthy microorganisms, such as Lactobacillus and Bifidobacterium strains (ADEBOLA, et al., 2014; VÁSQUEZ, et al., 2000; CAO, GUO, HUA, 2020; PALANIAPPAN, 2021). XOS are sugar oligomers made up of xylose units, they are also considered healthy due to their capacity to prevent disorders in human nutrition, such as gastrointestinal disorders and infection, obesity control and better nutrient absorption (KUMAR, et al., 2021; MANO, 2017). This thesis aimed to debate the methods involved in COS and XOS production, testing chemicals, physical and biological pretreatments, and the combination of them, considering the biotechnological and economics aspects. OBJECTIVES The objective of this study was to produce COS and XOS using sugarcane bagasse and cotton waste via biological and physical pretreatment, and via chemical, physical and biological pretreatment, respectively. A combination of pretreatment was evaluated and the pretreated material was characterized. 14 Specific Objectives - Evaluate the structural changes of sugarcane bagasse due to biological pretreatment with white-rot fungi; - Evaluate the physical pretreatment effects in the sugarcane bagasse after the biological pretreatment; - Evaluate the production of COS and XOS via enzymatic hydrolysis of a combination of pretreated sugarcane bagasse; - Evaluate the structural changes in cotton biomass after physical, chemical and biological pretreatment and a combination of them; - Evaluate the production of COS via enzymatic hydrolysis using cotton residue after ball milling, alkaline and biological pretreatment. Thesis format and organization This thesis was organized into chapters, which were designed considering independent publications such as book chapter and papers. The organization was as follows: General introduction and objectives; Chapter 1 – book chapter: Biotechnological Aspects of Microbial Pretreatment of Lignocellulosic Biomass (DOI: https://doi.org/10.1007/978-981-15-9593-6_6), published in Biorefineries: A Step Towards Renewable and Clean Energy, published in Springer, reuse authorization license number 5947030195987 by Spring Nature. This chapter presents a bibliography review about the biological pretreatment in lignocellulosic biomasses. In the chapter was discussed the biotechnological aspects of this technique and presented the microorganisms involved in the process, as well as the enzymes and possibilities of the biological pretreatment in the industry; Chapter 2 – article: Efficient production of cellooligosaccharides and xylooligosaccharides by combined biological pretreatment and enzymatic hydrolysis process (DOI: https://doi.org/10.1007/s13399- 024-05703-1), published in Biomass Conversion and Biorefinery, from Springer, reuse authorization license number 5945500157455 by Spring Nature. Chapter 2 is an article about biological pretreatment, involving the production of COS and XOS using two kinds of physical pretreatment, such as knife mill and ball mill, evaluating different charges of enzymes. Also, were evaluated the changes in the lignocellulosic biomass structure after biological pretreatment and the crystallinity index after knife and ball 15 milling. Chapter 3 - article: Cellooligosaccharides production from cotton residue using biological pretreatment, alkali and milling approaches. The third chapter is an article about COS production from cotton residue involving chemical, physical and biological pretreatments. With the focus on the COS production, a waste with high cellulose content was chosen. The cotton residue was also analyzed by x-ray, FTIR-ATR and scanning electron microscope to evaluate the modifications in the biomass before and after the pretreatments; General conclusions. . 16 REFERENCES ADEBOLA, Oluwakemi Obasola; CORCORAN, Olivia; MORGAN, Winston A. Synbiotics: the impact of potential prebiotics inulin, lactulose and lactobionic acid on the survival and growth of lactobacilli probiotics. Journal of functional foods, v. 10, p. 75-84, 2014. AGBOR, V. B. et al. Biomass pretreatment: fundamentals toward application. Biotechnology advances, v. 29, n. 6, p. 675-685, 2011. CAO, Rou et al. Investigation on decolorization kinetics and thermodynamics of lignocellulosic xylooligosaccharides by highly selective adsorption with Amberlite XAD-16N. Food chemistry, v. 310, p. 125934, 2020. FELIPUCI, Jefferson Poles et al. Biological pretreatment improved subsequent xylan chemical solubilization. Biomass Conversion and Biorefinery, p. 1-8, 2021. FELIPUCI, Jefferson Poles et al. Biotechnological aspects of microbial pretreatment of lignocellulosic biomass. Biorefineries: a step towards renewable and clean energy, p. 121-150, 2020. FORSAN, Carolina Froes et al. Xylooligosaccharide production from sugarcane bagasse and leaf using Aspergillus versicolor endoxylanase and diluted acid. Biomass Conversion and Biorefinery, p. 1-16, 2021. IOELOVICH, M. Ya. Models of supramolecular structure and properties of cellulose. Polymer Science Series A, v. 58, n. 6, p. 925-943, 2016. KUMAR, Vishal et al. Developing a sustainable bioprocess for the cleaner production of xylooligosaccharides: An approach towards lignocellulosic waste management. Journal of Cleaner Production, v. 316, p. 128332, 2021. MANO, Mario Cezar Rodrigues et al. Oligosaccharide biotechnology: an approach of prebiotic revolution on the industry. Applied microbiology and biotechnology, v. 102, p. 17-37, 2018. QASEEM, Mirza Faisal; SHAHEEN, Humaira; WU, Ai-Min. Cell wall hemicellulose for sustainable industrial utilization. Renewable and Sustainable Energy Reviews, v. 144, p. 110996, 2021. RASTOGI, Meenal; SHRIVASTAVA, Smriti. Recent advances in second generation bioethanol production: An insight to pretreatment, saccharification and fermentation processes. Renewable and Sustainable Energy Reviews, v. 80, p. 330-340, 2017. SCAPINI, Thamarys et al. Hydrothermal pretreatment of lignocellulosic biomass for hemicellulose recovery. Bioresource Technology, v. 342, p. 126033, 2021. SHINDU, R.; BINOD, P.; PANDEY, A. Biological pretreatment of lignocellulosic biomass - An overview. Bioresource Technology, v. 199, p. 76-82, 2016. 17 VÁZQUEZ, M. J. et al. Xylooligosaccharides: manufacture and applications. Trends in Food Science & Technology, v. 11, n. 11, p. 387-393, 2000. 18 CHAPTER I: BIOTECHNOLOGICAL ASPECTS OF MICROBIAL PRETREATMENT OF LIGNOCELLULOSIC BIOMASS 19 ABSTRACT In several areas, products are obtained from lignocellulosic biomass, such as bioethanol and personal items. Notwithstanding, it features high recalcitrance, hence its use often demands pretreatment and hydrolysis stages to reach bio-based final products. Industrially, the most common method is the chemical pretreatment which, as the name implies, involves chemical components with potential environmental risks. This procedure is responsible to increase biomass accessibility and to enhance polysaccharides achieving in subsequent stages. Biological pretreatment presents a new perspective to replace or cooperate with its chemical counterpart, once microorganisms can modify the lignocellulosic structure and facilitate accessibility to macromolecules of interest. According to the above, this chapter covers the potential of biological pretreatment as well as the mechanisms of microbial degradation, their enzymes, and the impacts on the economy worldwide. Keywords: sugarcane bagasse; fungus; microorganism; biological pretreatment; degradation 20 1.1 INTRODUCTION Recent technological, social and environmental changes have brought new needs in both science and industry for developing alternative technologies that make it possible to achieve similar products, than those obtained from petroleum sources (RUAN et al. 2019). Since the last years of the 19th century, the world energy matrix has been based on fossil fuels (British Petroleum 2019). Among the possibilities to replace oil, biomass has become the most important resource, able to generate several products by different routes, with the great advantage of being environmentally friendly (GUEDES et al. 2019). In this perspective, bio-based products are currently part of everyday life, with applications in sectors such as engines, packaging, medicines, and many others. With or without slight treatment/modifications, vegetal biomass like crops, vegetable oils, forest, agricultural waste, and also the municipal and industrial ones are used to produce bioproducts (SOROKINA et al. 2017; ROSALES-CALDERON and ARANTES 2019). However, turning vegetal biomass into bioproducts may become a challenge, since the raw material needs to be undergone to different types of stages during the conversion process until reaching suitable yields (HOLWERDA et al. 2019). Pretreatment has a huge importance in the steps of value- added products generated from biomass systems, where complex structure presented in plants must be conditioned for subsequent stages (ANTUNES et al. 2019). The most used methods of biomass pretreatments, such as chemical and physical procedures, have in common the demand for plenty of chemical reagents and/or energy inputs in its process. Such chemicals are widely used in industries to separate biomass components in order to manufacture all kinds of (bio-) products, but in consequence, those reagents are found polluters for the environment. Nowadays, facing an economic and global warm crisis, it is essential and recommended looking for alternatives to low-cost, less oil-dependent, and non-polluting manufacturing methods. Biological pretreatment of biomass is already known as an option to conventional methods used in industries. This method does not generate toxic and inhibitory compounds and need low quantity of chemical and energy input, which makes it an economically and eco-friendly feasible process. Biological pretreatment also can be used before a chemical or physical pretreatment: the biological stage can provide a better decrease of the recalcitrance while the chemical stage provides the 21 separation of the macromolecules. This combination can reduce the costs and chemicals in the whole process (SHINDU et al. 2016; SINGH et al. 2018; AGBOR et al. 2011, FELIPUCI 2020). In this chapter will be discussed how biological pretreatment works, including the enzymes and microorganisms involved in the biomass structure modification. Moreover, the benefits and disadvantages of this method are discussed, as well as value-added and commodity products, mainly on large scale. 1.2 BIOLOGICAL PRETREATMENT Biological pretreatment of lignocellulosic biomass became a fundamental research topic since it is clear that a near-term economy will depend on the supply of biomass to produce bioproducts and bioenergy. It is related to the use of microorganisms, aiming to degrade or modify vegetal biomass structure employing their special enzymatic complexes (AGBOR 2011; SINDHU et al. 2016). Among the vast variety of species in the world, fungi and bacteria are well known to produce specific enzymes for lignocellulose deconstruction, called cellulases, hemicellulases, and ligninases. These enzymes are capable to degrade natural macromolecules found in the plant cell wall, such as cellulose, hemicelluloses, and lignin. Cellulose and hemicelluloses, for instance, are hydrolyzed into smaller molecules (the monomeric sugars) (SHARMA et al. 2019). Among the numerous enzymes produced by fungi that degrade cellulose, hemicellulose and lignin, the most studied are: endo-glucanases, exo-glucanases and β-glucosidases that hydrolyze cellulose; endo-xylanases, β-xylosidases, acetyl xylan esterases and others that degrade xylan and laccases, manganese-peroxidases and lignin-peroxidases that degrade lignin (PAMIDIPATI and AHMED 2019; GAUTAM et al. 2019; MALGAS et al. 2019). The species of fungi that degrade lignin are known as white rot. The ones that depolymerize cellulose and hemicelluloses are named brown rot because the wood degraded takes a brownish appearance, due to the loss of polysaccharides (cellulose and hemicellulose) remaining high amounts of lignin (HATAKKA and HAMMEL 2011). Biological pretreatment does not generate toxic compound (degradation products, inhibitors) during its process and it is ecologically promising, which is an advantage comparing to other usual methods. Moreover, results can be optimized when the 22 strains are pre-selected (SINDHU et al. 2016; VAN KUIJK et al. 2015). In the biodegradation, variable microbial communities is important to the quality of the final results due to its vast amount of enzymes. However, in addition to the microorganism itself, biomass composition, temperature, humidity, pH, aeration rate, incubation time and biomass particle size are elements that can also affect the result and the quality of the pretreatment (SINDHU et al. 2016; FANG et al. 2012; LI et al. 2012; IQBAL et al. 2013; FATOKUN et al. 2016). Usually, biological pretreatment needs long-time requirements (10-14 days), space, and careful growth conditions to work. In industrial scale it may be less attractive but the biological pretreatment can be used together with chemicals and physical pretreatment. The potential of delignification by microorganisms combining with chemical and physical methods is inviting because of the complete degradation of lignocellulosic biomass components, mainly lignin, that can take a long time to reach significant results (AGBOR et al. 2011; HATAKKA 1994; HATAKKA et al. 1993). Recalcitrance is the capacity of a biomass resist to a pretreatment or to enzyme action. The quantity and organization of the components into the cell wall such as cellulose crystallinity are factors that may change the recalcitrance level of biomass (NAIDU et al. 2018; MELATI et al., 2019; PARK et al. 2010). Lignin is suggested to has a crucial hole in the recalcitrance due to its resistance against pathogens and insects, and its removal influences the access to the polysaccharides (SHIMIZU et al., 2020; SCHMATZ et al., 2020; ZHAO et al. 2012; PHITSUWAN et al. 2013) High recalcitrance is a challenge in the search for better methods of macromolecules isolation from biomass. Accordingly, different pretreatment methods have been developed, aspiring to circumvent this problem in order to separate its components. One method to work around the recalcitrance problem is to select varieties with low lignin content (BRIENZO et al., 2015), or delignify biomass decreasing lignin content, considering that lignin is a barrier in carbohydrate extraction (SHIMIZU et al., 2020; BRIENZO et al., 2017). An usual pretreatment focuses on improving the formation or capability to form fermentable sugars by hydrolysis; avoid by-product formation that may prevent subsequent processes and be a good cost- benefit ratio method (MELATI et al., 2019). Thus, biological pretreatment is an option to replace or co-work with other methods of pretreatment by attending such ideal requirements. 23 Other way to degrade lignocellulosic biomass is using co-culture, which use more than one microorganism. This method is based in to use fungus or/and bacteria to degrade the lignocellulosic biomass. However, competition between microorganisms for the substrate is not recommended, and it can be used one after other. This technique is useful due to both microorganisms encompass large quantities of enzymes, which can complete degrade the lignocellulosic material. This process can be used in different areas such as agronomy (degrade pesticides) and industry (carpet decolorization) (YOON et al. 2014; SARIWATI et al. 2017; WANG et al. 2017; SARIWATI et al. 2017; KUMARI and NARAIAN 2016). 1.2.1 Lignocellulosic Biomass structure Lignocellulosic biomass englobes all organic matter directly from plant sources. It is the largest source of carbohydrates in nature, with a great variety, abundance and availability, involving wood, agro-industrial waste, municipal waste, and plants. What draws attention to these materials is that they are renewable resources with energy potential. This presents them as possible substitutes for fossil fuels, generating sustainable energy through bioethanol and co-generation of electric energy (by a burning process) (NANDA et al. 2015). Consequently, interest in research, both in scientific and industrial fields, grows constantly (BILGILI et al. 2017; MAO et al. 2015; ASLAN 2016; TOKLU 2017; SHARMA et al. 2019). One of the most used lignocellulosic biomass is the sugarcane bagasse (Saccharum spp) Currently, the bagasse is used in the production of electrical and thermal energy through its combustion in high-pressure boilers in plants (FERNANDES et al. 2018). Another application aims to obtain second-generation ethanol (cellulosic ethanol), serving as an alternative to replace fossil fuels. Lignocellulosic biomass is also used in the production of clothing, artificial skin, paper, and other products in common use (MIZUHASHI et al. 2015; KIM et al. 2014). More specifically, in biotechnology and biomass conversion, it is possible to produce briquettes, carbon adsorbents, and biofilms. The production of these items depends on the treatment that those biomasses will be undergone. For separation of each macromolecule, there is one or a series of treatments to be based on biological routes. The main characteristic of vegetal biomass is its lignocellulosic structure existing into the cell wall, presented in all plant forms. Its composition is mainly cellulose, 24 hemicelluloses, and lignin, with less quantities pectins, proteins, and extractives (NAIDU et al. 2018). Quantities of each component change according to biomass and soil types, geographic localization, and other factors (De VASCONCELOS, 2015). The three main components (cellulose, hemicelluloses, and lignin) in the cell wall are organized in a way that recalcitrance is increased, making its separation harder in biotechnological processes. Cellulose and hemicelluloses are strongly connected by hydrogen bonds. Hemicelluloses can be located between cellulose fibers, while lignin is connected to the carbohydrates forming a complex interaction network (SCHMATZ et al. 2020; BUSSE-WICHER 2016). Cellulose is the major macromolecule in the plant cell wall (Fig 1). The quantity varies according to biomass type: cotton presented around 95% and sugarcane bagasse showed 25 to 45% (NAIDU et al. 2018). It is also considered most abundant organic polymer found on the planet Cellulose is an arrangement constituted by cellobiose unities (glucose dimers) joined by β-1,4 glycosidic chains. In the cellulose structure, there are amorphous regions which are organized regions demined crystalline and non-crystalline zones (IOELOVICH 2016). Cellulose is widely sought in the industry as raw material for common use products, such as varnish, films, paper, among others. Due to several industrial interests, cellulose isolation from biomass is widely studied. Cellulose can be separated from other carbohydrates by alkaline treatment, or broken by acid treatment. In the case of alkaline treatment, ester linkages break down, resulting in structural modification of the cell wall and facilitating separation from hemicelluloses (GALLETTI and ANTONETTI 2012). 25 Figure 1.1 - Schematic representation of lignocellulosic biomass emphasizing the cellulose macromolecule Source: JASMANIA and THIELEMANS 2018 Hemicelluloses, different from cellulose, are composed of more than one monosaccharide: pentoses, hexoses and uronic acids. In pentoses group is found xylose and arabinose; in hexoses group is found mannose, glucose and galactose and in uronic acids is found glucuronic and galacturonic acids. Those monosaccharides can also be subdivided into three main groups: xyloglucans, xylans, and mannans, that are formed by subunits of mannose. Many of the OH groups at C2 and C3 of xylose units are substituted with O-acetyl groups. The monosaccharides are connected by β and  glycosidic bonds and can have between 80-200 units. Hemicelluloses have amorphous characteristics and a lower degree of polymerization than cellulose. It makes up 15 to 35% of lignocellulosic biomass and it is associated to the integrity of the plant cell wall, having great importance in its shape and resistance. Hemicelluloses correspond to one-third of all renewable carbon on the planet. Hemicellulose has been studied for several applications, with a feature for oligomers such as xylooligosaccharides and manooligosaccharides (De FREITAS et al., 2019; CHIYANZU et al., 2014). Lignin is a biomass macromolecule composed of phenylpropane units of p- hydroxyphenyl (H), syringyl (S) and guaiacyl (G). This polyphenolic structure is 26 organized irregularly and has an amorphous structure. Depending on species, lignin comprehends between 10 to 20% of lignocellulosic biomass, being the third most abundant macromolecule in the plant cell wall. For plants, lignin helps in protection against insects and fungi and also contributes to growth development and mechanical strength. This protection is one of the reasons to the biosynthesis, once infections, metabolic stress and disturbances in cell wall structure are starters to the plant initiate the process (VANHOLME et al. 2010). It is arranged mainly on the secondary wall, making it rigid and waterproof. The most common linkages in lignin macromolecule is the β-O-4, however there are others kind of linkages more resistant to chemical degradation, such as β-5, β- β, 5-5, 5-O-4 and β-1 linkages. Lignin organization is linked with hemicelluloses, together with its irregular structure and a gigantic number of possibilities for connections between its forming units, which suggests that there is a low chance of existing two similar lignin molecules (RALPH et al. 2004). This favors the increasing recalcitrance of its biomass (SCHMATZ et al., 2020). Lignin is an obstacle for a process dedicated to macromolecule separations as it remains as residual content/contaminant (FELIPUCI, 2020). 1.2.2 Microorganisms in Biological pretreatments Microorganisms are considered of key function in biological pretreatments of lignocellulosic biomass. Degradation capacity of microorganisms is widely known, mainly because of the degradative potential of its enzymes, which are produced during its growth. Biological pretreatment technology has generated results in several areas involving biotechnology, bioremediation, biopulping among others. The most common microorganisms applied in biological pretreatment are white-rot, brown-rot, and soft-rot fungi, besides bacteria. These microorganisms are capable to consume all components in lignocellulosic biomass, mainly lignin, and the capacity to mineralize lignin into carbon dioxide and water. Brown-rot fungi are known to degrade polysaccharides more efficiently, and only slightly modifies the lignin, while white-rot fungi can degrade lignin with more facility (KIRK and MOORE 1972; KIRK and HIGHLEY 1973). Holocelluloses/lignin ratio presented in biomass after degradation can be used to measure the fungal effect on the biomass decomposition. The effect on the biomass components can be classified at different ratios: Class 1 (corresponds to brown decomposition agents): ratio less than one; Class 2: whose process has a low 27 amount of residual lignin; Class 3: holocelluloses content is two to five times higher than lignin content; Both classes 2 and 3 correspond to white decomposition agents (TROJANOWSKI 2001). 1.2.2.1 White-Rot Fungi Industrially white-rot fungi are well known as lignin consumers, found in Basidiomycota phylum. Those comprehend over than 90 % of all Basidiomycetes that rot woods. (RILEY et al 2014). This phylum has been studied in several areas, including medicine (MADHANRAJ et al. 2019), agriculture (DUPLESSIS et al. 2011), and forestry (MARTIN et al. 2008). This phylum also includes mushrooms (MORIN et al. 2012), and pathogens of plants, animals, and other fungi (DUPLESSIS et al. 2011; DAWSON 2007). White-rot fungi have great potential to degrade lignocellulosic biomass (Fig 2). Although those fungi also can degrade polysaccharides, they are known as a well specific lignin degrader (RUDAKIYA and GUPTE 2017). Syringil (S) units of lignin usually are preferred instead of guaiacyl (G) units, due to its less resistance to degradation. In certain conditions, white-rot fungi are lignin-selective depending on several factors, like cultivation time, temperature, wood species, and other variables (HATAKKA and HAMMEL 2011; HAKALA et al. 2004). The degradation ability of these fungi has been quite studied not only in lignocellulosic biomass researches, but also in other areas, such as bioremediation, food, pharma, and other industries. These abilities allows the fungi grow in restrictive conditions, such as lignocellulosic wastes In the last decade, several studies focused on these group showed results to degrade pesticides (KAUR et al. 2016; GOUMA et al. 2019), to increase productivity, efficiency, and quality of several products (KUSHWAHA et al. 2018) and applied in pulp and paper industry (SINGH 2018). 28 Figure 1.2 - Scanning electron micrographs of beech wood degradation by white-rot fungi after 120 days Source: BARI et al, 2018. A, C, and E: Pleurotus ostreatus; B, D, and F Trametes versicolor. A and B shows cross-sections (bar 20 µm): the arrows point cell walls already degraded and arrowheads point colonization of hyphae in the cell lumina; C and D show radial sections (bar 100 µm): the arrows point a entire decomposition of ray parenchyma and arrowheads point deconstruction of cell walls and vessels; E and F show tangential sections (bar 100 µm): the arrows point the separation of ray wall with vessels lumina, while arrowhead point disintegration of woody structure. 1.2.2.2. Brown-Rot Fungi 29 Brown-rot fungi are also found in the Basidiomycota group, representing nearly 7 % of this phylum (HATAKKA and HAMMEL 2011; GOODELl 2003). An examples of families that concern to brown-rot fungi are: Adustoporiaceae, Postiaceae Auriporiaceae and others (LIU et al, 2022) Evolutionarily, most of this group are derived from white-rot fungi, probably by losing of decay capability and biodegradative mechanisms (HIBBETT and THORN 2001). Otherwise, white-rot and brown-rot classification are discussed, since new genetic studies suggest continuum rather than a dichotomy between these two groups. In this case, authors suggest that the “white- rot fungi” term would be restricted to fungi that consume all the cell wall macromolecules through activity of lignin-degrading peroxidases (RILEY et al. 2014). The brown color of brown-rot fungi is due to residual lignin left after degradation. It is caused by fungi enzymatic arsenal that degrades polysaccharides. Hemicellulose degradation is faster and polysaccharide depolymerization involves oxidative components and hydrolytic enzymes (HATAKKA and HAMMEl 2011). Degradation capacity is widely known in the bio-pulping area. Bio-pulping is a process where wood chips are treated by microorganisms to improve quality and make stronger paper produced. This method reduces toxicity and pitch content (GUPTA 2019). Using some species of brown-rot fungi with worms to degrade paper mill sludge is a useful strategy to enhance cellulose decomposition (NEGI and SUTHAR 2018). 1.2.2.3. Bacteria Bacteria are known to produce cellulolytic, hemicellulolytic, and ligninolytic enzymes that can also be used in biological pretreatment (SHARMA et al. 2019). An advantage in comparison to fungal pretreatment is that some bacteria can grow faster than fungi besides degrade lignin into small particles. (HATAKKA 2005; KURAKAKE et al. 2007). Examples of wood degrading bacteria can be: Betaproteobacteria, Gammaproteobacteria, and Acidobacteria (JOHNTON et al, 2016). Although bacteria can properly degrade lignocellulosic biomass, its sole use as biological pretreatment has not proved efficient. However, it can improve the enzymatic digestion of lignocellulose after applying another pretreatment, such as physicochemical method (ZHUO et al. 2018). Co-culture using bacteria and/or fungi can degrade lignocellulosic biomass almost completely due to high enzymatic activity. 30 Selecting the best strains, that can produce necessary enzymes is essential for an efficient biological pretreatment in order to produce biofuels and bioproducts (SHARMA et al. 2019). 1.2.3 Enzymes Involved in Biological Pretreatment The effectiveness of a biological pretreatment depends on enzymes ability to address biochemical and physical barriers to hydrolysis. Therefore, a mix of enzymes can co-work to increase biomass access by expanding small pores and open the cell wall matrix (AMIN et al. 2017). Lignocellulose degradation by microorganisms is mainly accomplished by a system of extracellular enzymes that hydrolyze and oxidize the biomass component (Fig 3). Hydrolases (cellulases and hemicellulases) are produced by hydrolytic system to degrade polysaccharides and oxidative catalytic system to degrades lignin by the production of ligninases (SAJITH et al. 2016). Figure 1.3 - Simplified representation of lignocellulolytic enzymes and their action mode Source: SAJITH et al. 2016. 31 1.2.3.1 Cellulases Cellulases are glycosyl hydrolases (GHs) produced by microorganisms while they grow on lignocellulosic materials. They hydrolyze cellulose into shorter chain polysaccharides by breaking down β-1,4-glycosidic bonds. In their structure, they usually have a catalytic domain at the N- terminal and a carbohydrate-binding module at the C- terminal. The first one cleaves the glycosidic linkage and the second one destiny the catalytic domain to the polysaccharide substrate (JAYASEKARA and RATNAYAKE 2019; OBENG et al. 2017). Three main enzymes comprise cellulases enzyme system, endoglucanases (endo-β-1,4-D-glucanases; EC 3.2.1.4), exoglucanases (exo-β-1,4-D-glucanases; EC 3.2.1.91) and glucosidases (β-D-glucoside glucanhydrolases, EC 3.2.1.21). These enzymes are categorized as per their structure and function, however their collaborative work is essential for complete hydrolysis of the complex cellulose fibers (SAJITH et al. 2016). Endoglucanases generate oligosaccharides with free chain ends by hydrolyzing internal β-1,4-glycosidic bonds and acting randomly on amorphous areas of cellulose. These enzymes can convert cellodextrin (intermediate product of cellulose hydrolysis) into cellobiose and glucose (SINGH et al. 2016). Endoglucanases has rapid dissociation, can reduce chain length and viscosity by acting on cellulose but exhibit no activity against crystalline cellulose such as avicel (AKAMINE et al. 2018; OBENG et al. 2017; SAJITH et al. 2016). Exoglucanases acts on the crystalline region of cellulose and release cellobiose as product from reducing (EC 3.2.1.91) or non-reducing ends (EC 3.2.1.176). The oligosaccharide chain portion that each enzyme attacks are related to its classification. However the actions of the enzymes are unidirectional in a long-chain oligomer (OBENG et al. 2017; SINGH et al. 2016). These enzymes are more active against crystalline cellulose substrates such as avicel and cellooligosaccharides but do not hydrolyze soluble resultants of cellulose like carboxymethyl cellulose (JAYASEKARA and RATNAYAKE 2019; SAJITH et al. 2016). β-glucosidases present rigid structure with an active site that favors disaccharides entry, however, they also can hydrolyze low degree of polymerization soluble cellodextrins. These enzymes act on cellobiose to complete the hydrolysis 32 process of cellulose. As result, glucose with a free hydroxyl group at C4 from the non- reducing end of oligosaccharides are released (OBENG et al. 2017; SAJITH et al. 2016). Retention and reversion are catalytic mechanisms that lead to successful cellulose hydrolysis. This is performed by two catalytic amino acid residues of the enzymes, a proton donor and a nucleophile. Both of them stereochemicaly modifies the anomeric carbon configuration, facilitating enzymatic cleavage of the glycosidic bonds (GARVEY et al. 2013). Cellulolytic enzyme multisystem can suffer inhibition by its products. For this reason, β-glucosidases and exoglucanases are essential to alleviate exo- and endoglucanases, respectively, from feedback inhibition. In the same way, β- glucosidase is also inhibited by glucose, therefore is necessary a search for glucose tolerant β-glucosidases (OBENG et al. 2017). Complementary action of these cellulases is crucial for efficient hydrolysis in order to obtain glucose residues, which can be used for several applications such as the production of biofuel and chemicals. Among microorganisms, fungi are responsible for approximately 80% of cellulose hydrolysis and therefore, considered great cellulase producers (SINGH et al. 2016). 1.2.3.2 Hemicellulases Efficient hemicellulose hydrolysis of lignocellulosic biomass improves hydrolysis yield and consequently reduces enzyme costs and dosages, which makes crucial the use of hemicellulases. They are most often glycoside hydrolases and are usually produced by microorganisms together with cellulases. The hemicellulose backbone of a lignocellulosic biomass can be composed by different polysaccharides, depending on the source (SINDHU et al. 2016; SINGH et al. 2016). Mannan and xylan are the most common hemicelluloses found in nature. Xylan is the main hemicellulose in lignocellulosic biomass from agriculture residues, comprised of xylose units in the backbone chain that are usually linked to acetyl and ferulic groups, arabinofuranosyl or glucuronic acid residues. Therefore, multiple enzymes are necessary to decompose xylan, including endoxylanase (EC 3.2.1.8), β- xylosidase (EC 3.2.1.37) that acht on the main chain of xylan. The enzymes that work on the pending grups are α-arabinofuranosidase (EC 3.2.1.55), and α-glucoronidases (EC 3.2.1.139) (ÁBREGO, CHEN & WAN, 2017). In addition, acetyl xylan esterases 33 (EC 3.1.1.72), ferulic acid esterases (EC 3.1.1.73), and p-coumaric acid esterases (EC 3.1.1.x) are also requested for the complete deconstruction of xylan (CHADHA et al. 2019). Hemicellulases structures are consisted by a catalytic domain to perform enzyme functions. They can be glycosyl hydrolases that cleaves glycosidic bonds or can be carbohydrate esterases that hydrolyze ester bonds, between xylan and acetic acid or ferulic acid substitutions (JUTURU and WU, 2013). Xylanases hydrolyze β-1,4 linkages in xylan backbone chain, producing xylooligosaccharides. Most of them belong to glycoside hydrolase (GH) families 10 and 11, however enzymes that are exclusively active on D-xylose-containing substrates, known as “true xylanases”, are only on family 11 (TYAGI, PATIL and GUPTA, 2019). β- xylosidases hydrolyze a low degree of polymerization xylooligomers, produced by xylan hydrolysis, into xylose. Xylanases action is inhibited by xylooligomers produced in the hydrolysis, therefore β-xylosidases action removes end-product inhibition increasing the efficiency of xylanases (CHADHA, RAI and MAHAJAN, 2019). β-mannanases hydrolyze mannan-based hemicelluloses. As result, short β-1,4- mannooligomers are released that can be hydrolyzed into mannose by β- mannosidases. Arabinofuranosidases catalyze the removal of arabinosyl substituents and facilitate an increase in access points of xylanase to xylan Both β-mannanases and arabinofuranosidases are required for mannan or arabinofuranosyl containing hemicelluloses (TERRONE et al. 2020). The α-1,2-glycosidic bond can be broken down by α-D-glucuronidases releasing glucuronic acid from the xylan chain (CHADHA, RAI and MAHAJAN, 2019; SINGH et al. 2016). Acetyl xylan esterases are enzymes responsible to remove acetyl groups linked to β-D-xylopyranosyl residues by hydrolyzing the ester bonds. The accessibility of enzymes that break the backbone by steric hindrance can be interfered by acetyl side- groups, therefore their removal makes the xylanases action easier. Ferulic acid esterases and p-coumaric acid esterases also catalyze ester bonds on xylan. The first enzymes are recognized to break down ester linkages between ferulic acid and arabinose substitutions on xylan, and the second acts on the bond between arabinose and p-coumaric acid (CHADHA, RAI and MAHAJAN 2019; BAJPAI 2014). Due to the complex chemical structure of hemicelluloses, its hydrolysis into its constituent monomers requires catalytic action of versatile enzymes that work synergistically. Hemicellulolytic enzymes can be produced by different fungi and bacteria, however, the source of most commercially important hemicellulases are fungi 34 (MANJU and CHADHA 2011). They have biotechnological potential and several industrial applications, like hemicelluloses hydrolysis of lignocellulosic biomass, improving cellulose saccharification (CHADHA, RAI and MAHAJAN 2019). 1.2.3.3 Ligninases Lignin is one of the main responsible for recalcitrance in lignocellulosic biomass because its complex structure, protecting polysaccharides (SCHMATZ et al. 2020). To break down the lignin structure, microorganisms developed some specific extracellular enzymes based on oxidative reactions. In nature, lignin degradation is important to the biogeochemical carbon cycle (RUIZ-DUEÑAS and MARTÍNES 2009). Those enzymes are also used in the bioremediation process and its action is an important step for lignin removal in industries that work with cellulosic biomass (JHA 2019). Ligninases are, generally, separated in two different types: phenol oxidases and peroxidases. Laccases are an example of phenol oxidases. Lignin degradation by laccases (EC 1.10.3.2) is normally by oxidation of phenolic compounds, yielding quinines and phenoxy radicals. Peroxidases make part of oxidoreductases family. This group of enzymes catalyzes lignin depolymerization utilizing H2O2 (SAJITH et al. 2016). Laccase enzymes are observed in plants, insects, bacteria, and fungi, mainly in the white-rot group. In fungi, these enzymes are involved not just in lignin degradation but also in sporulation, pigmentation of the fungus, detoxification, and fruiting body (CLUTTERBUCK 1990; THURSTON 1994). The molecular weight of laccase is around 50 to 100 kDa and they are classified as multicopper oxidases, which can be monomeric, dimeric, or tetrameric. Laccase use molecular oxygen to oxidize phenolic rings to phenolic radicals. Laccase can cleave Cα–Cβ cleavage, aryl-alkyl cleavage, and Cα-oxidation. Products may be submitted through non-enzymatic reaction, like polymerization, hydration, or dismutation, or a second enzyme-catalyzed oxidation (MADHAVI and LELE 2009; SAJITH et al. 2016). With a redox mediators present, laccases can also catalyze the breakdown of non-phenolic lignin structures, and cleave β-O-4 linkages. Lignin peroxidase (EC 1.11.10.14) is considered one of the key enzymes in plant cell wall degradation due to its ability to oxidize non-phenol lignin structures. This reaction can cleavage Cα–Cβ bonds, mediating ring-opening reactions. Lignin 35 peroxidases are oxidized by hydrogen peroxide, and, this catalysis results in the creation of intermediate radicals such as phenoxy and veratryl alcohol (WONG 2009; RUIZ-DUEÑAS and MARTÍNES 2009). Lignin peroxidase and laccase are considered “partners” enzymes in certain conditions, due to substrate provided by lignin peroxidase after lignin degradation (BOOMINATHAN and REDDY 1992). Manganese peroxidase (EC 1.11.1.13) attacks both phenolic and non-phenolic lignin units. This enzyme works as a mediator in enzymatic activity, once it is converted from Mn2+ into Mn3+. Several monomeric phenols are oxidized by Mn3+ cation, including dyes and phenolic lignin model compounds (DATTA et al. 2017). 1.2.4. Enzymatic Hydrolysis of Biological Pretreated Material In a biorefinery system, lignocellulosic biomass hydrolysis is an essential phase in the whole process, since through hydrolysis intermediate products are obtained by breaking up of macromolecules existent in pretreated biomass (BICHOT et al. 2018; POCAN et al. 2018). The intermediate denomination is because these products will be used at a subsequent stage of conversion, the main intermediate products are monomers such as hexoses and pentoses coming from cellulose and hemicelluloses (LOOW et al. 2016). Hydrolysis or saccharification can be performed by acid, enzymatic or combined procedures, among the aforementioned, the biological process is possibly the most researched in the last years (POCAN et al. 2018). Hydrolysis by biological routes shows benefits associated to mild temperature in operation, high ratio (quantitative) between obtained product and precursors (monomers), minimal corrosion problems and in enzymatic hydrolysis does not produce inhibitory chemicals that can modify enzymes activities (AMEZCUA-ALLIERI et al. 2017; JAHNAVI et al. 2017). The key to the biological hydrolysis of pretreated lignocellulosic biomass is the hydrolytic enzymes; cellulose saccharification happens by deed of cellulolytic enzymes (cellulases), and hemicelluloses splitting befalls by action of hemicellulolytic enzymes (hemicellulases) (BHARDWAJ et al. 2019; BARBOSA et al. 2020). These enzymes can be synthesized mainly by fungi, bacteria, yeast, or algae through its controlled growth in solid or submersed fermentations (DOTSENKO et al. 2018; ARUWAJOYE et al. 2020). Instead of producing hydrolytic enzymes, there is the alternative to purchase commercial enzymes prepared by different industries dedicated to 36 synthesize and purify enzymatic cocktails that act according to specific conditions in hydrolysis (FLORES-GÓMEZ et al. 2018). Table 1.1 shows a summary of some characteristics related to hydrolytic enzymes, their mode of action, product formation, and inhibitory aspects. 37 Table 1.1 - Properties of cellulases and hemicellulases action on lignocellulosic biomass Macromolecule Enzyme EC / Synonym Act on Product Compounds that cause inhibition Refere nces Cellulose Endoglycana ses 3.2.1.4 / Carboxylmethylcellulases (CMCases) -(1→4) bonds at non- crystalline sections Reducing and non-reducing new parts Cellobiose (MURP MURPHY et al. 2013; PARK et al. 2019) Exoglycanas es 3.2.1.91 / Cellobiohydrolases (CBHs) or Avicelases -(1→4) bonds at cellulose ends and new reducing parts and non-reducing parts Cellobiose and other glycooligomers Glycose (FRY 2003; VIANNA BERNARDi et al. 2019) - glycosidases 3.2.1.21 / Cellobiases -(1→4) bonds at cellobiose Glycose Glycose, mannose and galactose (TETE R et al. 2014; HSIEH et al. 2014) Xylan Endoxylanas es 3.2.1.8 -(1→4) bonds at xylan backbone Xylobiose and other xylooligomers Xylobiose and Xylotriose (PUCH ARt et al. 2018; FU et al. 2019) - xylosidases 3.2.1.37 -(1→4) bonds at xylobiose Xylose Xylose (YEOM AN et al. 2010; BOSETTO et al. 2016) - arabinofuranosidases 3.2.1.55 -(1→2), -(1→3) and -(1→5) bonds at xylose- arabinose linkages Xylose and Arabinose Glycose and Galactose (NUMA N and BHOSLE 2006; YEOMAN et al. 2010; MALGAS et al. 2019) - glucuronidases 3.2.1.139 -(1→2) bonds at glucuronic acid-xylose linkages Methyl glucuronic acid and xylose -- (DASH NYAM et al. 2018; MALGAS et al. 2019) - galactosidases 3.2.1.23 / Lactases -(1→4) bonds at galactose- xylose linkages Galactose and Xylose Galactose (HUSAI N 2010; KHOSRAVI et al. 2015; MALGAS et al. 2019) acetyl xylan esterases 3.1.1.6 Acetyl groups located at xylan branches Acetyl groups *Organophosphate compounds (MONT ORO-GARCÍA et al. 2011; PAWAR et al. 2016; 38 RAZEQ et al. 2018) ferulic acid esterases 3.1.1.73 Ester bonds in arabinose- ferulic acid linkages Arabinose, Ferulic acid and p- coumaric acid (phenolic acids**) -- (SZWA JGiER et al. 2010; LOPES et al. 2018) p-coumaric acid esterases 3.1.1.73 Ester bonds in arabinose-p- coumaric acid linkages Arabinose and p-coumaric acid (phenolic acid**) -- (LOPE S et al. 2018) Linear mannan Endomanna nases 3.2.178 -(1→4) bonds at mannan backbone Non- reducing new parts and mannobiose and mannotriose Glycose and Galactose (VRIES et al. 2005; DA CRUZ 2013; LOPES et al. 2018) - mannosidases 3.2.1.25 Non- reducing new parts and other mannosaccharides Mannose Sucrose (MCCA BE et al. 1990; DA CRUZ 2013; LOPES et al. 2018) - galactosidases 3.2.1.22 -(1→6) bonds at galactose- mannose linkages Galactose and Mannose *Ag2+ and Hg+ (Da CRUZ 2013; SIRISHA et al. 2015; LOPES et al. 2018) Galactan Endogalacta nases 3.2.1.89 -(1→3) bonds at galactanan backbone Galacto- disaccharide Galacto- trisaccharide and Galcto- tetrasaccharide -- (GONZ ÁLEZ-AYÓN et al. 2019) - galactosidases 3.2.1.23 / Lactases -(1→4) bonds at galactose- xylose linkages Galactose and Xylose Galactose (HUSAI N 2010; KHOSRAVI et al. 2015; MALGAS et al. 2019) Source: Authors *Product did not generate in pretreatment and/or hydrolysis of lignocellulosic biomass; ** Cause inhibition in most of hydrolytic enzymes 39 Finally, it should be taken into account that hydrolytic enzymes can suffer deactivation by temperature, pH, reaction time, stirring intensity, enzymatic loads, and mixing modes (BALAN 2014; HU et al. 2016; SINGHVI and GOKHALE 2019). Substrate characteristics and modifications over the enzymatic hydrolysis can increase the material recalcitrance (WALLACE et al. 2016). Therefore, it is recommended to develop new researches with new conditions that exploit novel tolerance levels for increasing pretreatment and hydrolysis yields. 1.2.5 Mechanisms of Cell Wall Degradation by Microorganisms During periods of fungal growth, cell wall undergoes structural modifications that allow access to inside components (RILEY et al. 2014). Although degrading enzymes are known and studied, degradation can occur in a different manner according to situations: chemical structure and composition of the cell wall are different among woody materials (or non-wood) and enzymatic arsenals of microorganisms are different among them (Fig 4). These factors determine the degradation level of the material and make it difficult to fully understand how biomass is consumed and how the degradation process occurs. Thus enzymes involved in the degradation process must be suitable to each substrate. Furthermore, it is important to evaluate which microorganism and its respective strain are most adequate for each kind of substrate. Degradation efficiency by microorganisms depends, in many cases, on the chemical structure of molecules and on the presence of efficient enzymes in degrading compounds, which are specific for most substrates (PEREIRA and FREITAS 2012). Biomass chemical structure can influence the metabolism of the microorganisms, especially regarding rates and extent of biodegradation. In the case of catabolic enzymes that have low specificity for its substrate, xenobiotics with a chemical structure similar to natural compounds can be recognized by an active enzyme system and, consequently, used by microorganisms as a source of nutrients and energy (PEREIRA and FREITAS 2012). 40 Figure 1.4 - Scanning electron microscope images on the surface of the Oil palm Empty Fruit Bunch Source: ARBAAIN et al 2019. (a) Untreated; (b) biologicaly pretreated using Schizophyllum commune (ENN1); (c) biologicaly pretreated using Phanerochaete chrysosporium. Carbon sources can influence fungi growth, which can affect growing patterns (MANNAA and KIM 2017). Hyphae development allows better colonization of lignocellulosic material and also penetrate easily to plant cell walls than bacteria, reaching macromolecules unavailable for those microorganisms (PEREIRA and FREITAS 2012). Enzymes are a crucial tool for the degradation of lignocellulosic biomass. Microorganisms release those enzymes which work in a synergistic and independently action, such as peroxidases, laccases, xylanases, and the other enzymes. An example of cell wall degradation is proposed in figure 5 (ZENG et al. 2014). In this degradation proposal, the plant cell wall is degraded by Phanerochaete chrysosporium, which is capable to degrade all components of the lignocellulosic biomass. Fungal hyphae attach inside the cell wall, secreting enzymes. Manganese peroxidases (MnP) oxidize Mn2+ to Mn3+ and break the phenolic and non-phenolic lignin units (DATTA et al. 2017; WONG 2009). Lignin peroxidases (LiP) oxidize non- phenolic structures to mineralized lignin, cleaving Cα–Cβ bonds, mediating ring- 41 opening reactions (WONG 2009; RUIZ-DUEÑAS and MARTÍNES 2009). This process occurs in the secondary cell wall, in which are located structural carbohydrates as well as aromatic backbone. Cellulases hydrolyze β-1,4-glycosidic bonds and act on the microcrystalline region in cellulose chain to break the cellulose into monomers of cellobiose and D-glucose. Cellobiose dehydrogenases co-work with cellulases to break cellulose chains into small saccharides, generating hydroxyl radicals, H2O2, and Fe3+. Figure 1.5 - Proposed process of degradation of the wheat straw cell wall by Phanerochaete chrysosporium Source: ZENG et al. 2014 Although the process of degradation could be different from all microorganisms, the enzymes work similarly but secreted at a different amount, and one characteristic that can be noticed is the variety of the lignocellulosic structure/composition. In wheat lignin degradation using analytical pyrolysis was revealed that Cα–Cβ bonds and free phenolic units are preferred than non-phenolic units by Pleurotus eryngii and Phanerochaete chrysosporium. This preferential is due to the redox potential, that is lower in comparison with the etherified ones, permitting easier oxidation by ligninolytic 42 peroxidases and laccases produced by the fungi. In vitro, applying enzyme in lignocellulosic biomass, P. eryngii is capable to reduce the phenolic content of lignin, evidencing its capacity of modifying a lignocellulosic materials (MARTÍNEZ et al. 2001; CAMARERO et al. 2001). Another example of lignocellulosic biomass deconstruction is with the brown-rot fungi Penicillium echinulatum. In this case, using different carbon sources, was grown wild-type (2HH) and a mutant strain (S1M29). It was realized that the mutant was more capable to produce cellulases and hemicellulases, showing that the variety of microorganisms can differentiate by the quantity of enzymes produced (SCHNEIDER et al. 2016). 1.3. ECONOMIC IMPACTS AND CHALLENGES ON INDUSTRIAL SCALE INVOLVING BIOLOGICAL PRETREATMENT Studies involving biological pretreatments are needed today for several reasons, including environmental friendly process, chemical reduction, and energy savings. There is a growing number of items produced from fossil derivatives such as plastics and tires that are not renewable, in addition to remaining in nature indefinitely. Nevertheless, it is important to mention that a biotechnological route should concern about energy and chemical reagents applied, aiming to be more advantageous than traditional processes. For biofuels, specifically, greenhouse gases bring concern and it is on the part of governments. Gas derived from fossil is already being replaced by biofuels, which draws attention to new processes of production and ways to reduce costs. The type of biomass, process complexity, and value of by-product influence the choice of pretreatment (BAJPAI 2016). Despite chemical pretreatments holding the main focus on these procedures, biological pretreatments are able to optimize those processes in several levels, for instance: reduce the water, chemicals, and energy spent, generate less inhibitor and toxic compounds, reduce the costs and improve performance and yield. In the food industry, one of the most worrying problems is waste since all economic classes in society have a certain degree of waste generation (MCCARTHY and LIU 2017). This food that is not used can be turned into energy by the biological or thermochemical process. Biological pretreatment in food waste has advantages in comparison with conventional methods of pretreatment such as low cost and simplicity 43 (PHAM et al 2015). Lignocellulosic biomass products can be a source of material and energy in order to support a more sustainable society. Products of direct consumption or second value-added are already present in human life such as paper, fibers and textiles, nanocellulose, organic acids, furfural, and others (ZAMANI 2015). Food and biofuels are examples where biological pretreatment can be used to improve the productivity and reduce costs. Moreover, the high quantity of products produced from biological pretreatment in lignocellulosic biomass makes them more accessible and, consequently, cheapest. Biological pretreatment can be economical, however, it is hard to speculate how much can impact and change the human lifestyle. The extensive number of products that can be produced with lignocellulosic biomass after a biological pretreatment makes harder this count, considering the production cost and sell value of each one. Products individually can be speculated is considering the cost. An example, the xylan extraction using biological pretreatment before chemical (H2O2) pretreatment reduced the need for the chemical reagent to reach the same results, which means less cost in the process (FELIPUCI 2020). On the other hand, the production of fermentable sugar by biological pretreatment of corn stover using posterior enzymatic hydrolysis showed to be more expensive (1.41 $/kg) than steam explosion (0.43 $/kg), dilute sulfuric acid (0.42 $/kg) and ammonia fiber explosion (0.65 $/kg) methods, anyway, the prices of all pretreatments depends from several factors (BARAl and SHAH 2017). In this case, there was no need of detoxification using biological pretreatment. However, this method investigated required reactors, mainly due to long pretreatment time. Biological pretreatment could considerer an option of process outside not using any reactor, but face other problems such as contamination. Although the advantages of an experimental scale, the use of biological pretreatment in the industry is still a challenge. Recent studies showed the potential of microorganisms in biofuels productions using biological pretreatment (YAHMED et al 2017; ZABED et al 2019). However, it is a common view of all the difficulties involved in biological pretreatment on a large scale. Microorganism utilization in biotechnological processes requires certain precautions, which needs to add one or more steps in the process: contamination and sterilization of growth site are some examples. Furthermore, microorganism growth is slow, while sugars are fundamental as an energy source (VASCO-CORREA et al. 2016; UMMALYMA 2019). An option to 44 improve the process and pass through those problems is the genetic engineering as well as co-culture of suitable microbial consortium (SHARMA et al, 2019). 1.4. CONCLUDING REMARKS Biological pretreatment has several advantages over traditional biomass separation methods. Application of microorganisms and their enzymes, in addition to enhancing the breakdown of lignocellulosic structure, makes the process cheaper and less aggressive to nature. An important advantage is no by-products generation, improving the fermentable sugars production by enzymatic hydrolysis of cellulose, with appreciable cost-benefit, among other benefits. Microorganisms present great potential for industrial use. Employment of microorganisms in pretreatments, or just their enzymes, can provide a reduction of energy and chemical reagents consumption in the separation process of lignocellulosic biomass macromolecules. Microorganism co-cultivation is a valid technique option with biotechnological potential, once the enzymes produced by the microorganisms can complement each other, achieving a greater degree of degradation. Mechanism degradation of plant cell wall depends on the microorganism in question and, mainly, on its enzyme production and action on lignocellulosic biomass. Even though the use of the micro in the industrial-scale requires greater cultivation assistance, it still offers important advantages: there are cost reduction and yield improvement for the biorefinery area and also less chemical residues in the environment. 45 REFERENCES ÁBREGO, U; CHEN, Z.; WAN, C. Consolidated bioprocessing systems for cellulosic biofuel production. In: LI, Y.; GE, X. (Ed.) Advances in Bioenergy. 2017. v. 2, p. 143- 182. AGBOR, V. B. et al. Biomass pretreatment: fundamentals toward application. Biotechnology advances, v. 29, n. 6, p. 675-685, 2011. AMEZCUA-ALLIERI, M. A.; SÁNCHEZ DURÁN, T.; ABURTO, J. Study of Chemical and Enzymatic Hydrolysis of Cellulosic Material to Obtain Fermentable Sugars. Journal of Chemistry, v. 2017, p. 1-9, 2017. AMIN, F. R. et al. Pretreament methods of lignocellulosic biomass for anaerobic digestion. AMB Express, v. 7, p. 72, 2017 ANTUNES, F. A. F. et al. Overcoming challenges in lignocellulosic biomass pretreatment for second-generation (2G) sugar production: emerging role of nano, biotechnological and promising approaches. 3 Biotech, v. 9, n. 230, p. 1-17, 2019. ARUWAJOYE, G. S.; FALOYE, F. D.; KANA, E. G. Process Optimisation of Enzymatic Saccharification of Soaking Assisted and Thermal Pretreated Cassava Peels Waste for Bioethanol Production. Waste and Biomass Valorization, v. 11, p. 2409-2420, 2020. ASLAN, A. The casual relationship between biomass energy use and economic growth in the United States. Renewable and Sustainable Energy Reviews, v. 57, p. 362-366, 2016. BAJPAI, P. Future Perspectives. In: BAJPAI, P. (Ed.) Pretreatment of Lignocellulosic Biomass for Biofuel Production. Singapore: Springer, p 77-81, 2016 BAJPAI, P. Microbial xylanolytic systems and their properties. In: BAJPAI, P. (Ed.) Xylanolytic enzymes. Academic Press, p. 19-36, 2014. BALAN, V. Current Challenges in Commercially Producing Biofuels from Lignocellulosic Biomass. International Scholarly Research Notices Biotechnology, v. 2014, p. 1-31, 2014. BARBOSA, F. C.; SILVELLO, M. A.; GOLDBECK, R. Cellulase and oxidative enzymes: new approaches, challenges and perspectives on cellulose degradation for bioethanol production. Biotechnology Letters, v. 42, n. 6, p.75-884, 2020. BHARDWAJ, N.; KUMAR, B.; VERMA, P. A detailed overview of xylanases: an emerging biomolecule for current and future prospective. Bioresources and Bioprocessing, v. 6, n. 40, p. 1-36, 2019. 46 BICHOT, A. et al. Understanding biomass recalcitrance in grasses for their efficient utilization as biorefinery feedstock. Reviews in Environmental Science and Bio/Technology, v. 17, p. 707-748, 2018. BILGILI, F. et al. Can biomass energy be an efficient policy tool for sustainable development? Renewable and Sustainable Energy Reviews, v. 71, p. 830-845, 2017. BOOMINATHAN, K.; REDDY, C. A. Fungal degradation of lignin: biotechnological applications. Handbook of applied mycology, v. 4, p. 763-822, 1992. BOSETTO, A. et al. Research Progress Concerning Fungal and Bacterial β- Xylosidases. Applied biochemistry and biotechnology, v. 178, p. 766-795, 2016. BRIENZO, M. et al. Influence of pretreatment severity on structural changes, lignin content and enzymatic hydrolysis of sugarcane bagasse samples. Renewable energy, v. 104, p. 271-280, 2017. BRIENZO, M. et al. Relationship between physicochemical properties and enzymatic hydrolysis of sugarcane bagasse varieties for bioethanol production. New biotechnology, v. 32, n. 2, p. 253-262, 2015. BRITISH PETROLEUM. BP Statistical Review of World Energy. London: Pureprint Group Limited, 2019. BUSSE-WICHER, M. et al. Xylan decoration patterns and the plant secondary cell wall molecular architecture. Biochemical Society Transactions, v. 44, n. 1, p. 74- 78, 2016. CAMARERO, S. et al. Compositional changes of wheat lignin by a fungal peroxidase analyzed by pyrolysis-GC-MS. Journal of Analytical and Applied Pyrolysis, v. 58, p. 413-423, 2001. CHADHA, B. S.; RAI, R.; MAHAJAN, C. Hemicellulases for lignocellulosic-based bioeconomy. In: PANDEY, A. et al. (Ed.) Biofuels: Alternative feedstocks and conversion processes for the production of liquid and gaseous biofuels. Academic Press, 2019. p. 427-445. CHIYANZU, I. Application of endo-β-1, 4, d-mannanase and cellulase for the release of mannooligosaccharides from steam-pretreated spent coffee ground. Applied biochemistry and biotechnology, v. 172, n. 7, p. 3538-3557, 2014. CLUTTERBUCK, A. J. The genetics of conidiophore pigmentation in Aspergillus nidulans. Journal of General Microbiology, v. 136, n. 9, p. 1731-1738, 1990. DA CRUZ, A. Mannan-Degrading Enzyme System. In: POLIZELI, M. de L.; RAI, M. (Ed.) Fungal Enzymes. Boca Ratón: CRC Press, p. 233-257, 2013. DASHNYAM, P. et al. β-Glucuronidases of opportunistic bacteria are the major contributors to xenobiotic-induced toxicity in the gut. Scientific Reports, v. 8, p. 1-12, 2018. 47 DATTA, R. et al. Enzymatic degradation of lignin in soil: a review. Sustainability, v. 9, n. 7, p. 1163, 2017. DAWSON, J. R.; THOMAS, L. Malassezia globosa and restricta: breakthrough understanding of the etiology and treatment of dandruff and seborrheic dermatitis through whole-genome analysis. Journal of Investigative Dermatology Symposium Proceedings, v. 12, n. 2, p. 15-19, 2007. DE FREITAS, C.; CARMONA, E.; BRIENZO, M. Xylooligosaccharides production process from lignocellulosic biomass and bioactive effects. Bioactive Carbohydrates and Dietary Fibre, v. 18, p. 100184, 2019. DE VASCONCELOS, J. N. Ethanol Fermentation. In: SANTOS, F.; CALDAS, C.; BORÉM, A. (Ed.) Sugarcane: Agricultural Production, Bioenergy and Ethanol. Academic Press, 2015. p. 311-340. DOTSENKO, A. S. et al. Enzymatic Hydrolysis of Cellulose Using Mixes of Mutant Forms of Cellulases from Penicillium verruculosum. Moscow University Chemistry Bulletin, v. 73, p. 58-62, 2018. DUPLESSIS, S. et al. Obligate biotrophy features unraveled by the genomic analysis of rust fungi. Proceedings of the National Academy of Sciences, v. 108, n. 22, p. 9166-9171, 2011. FANG, G. R.; LI, J. J.; CHENG, X.; CUI, Z. J. Performance and spatial succession of a full-scale anaerobic plant treating high-concentration cassava bioethanol wastewater. Journal of Microbiology and Biotechnology, v. 22, p. 1148-1154, 2012. FATOKUN, E. N.; NWODO, U. U.; OKOH, A. I. Classical optimization of cellulase and xylanase production by a marine Streptomyces species. Applied Sciences, v. 6, n. 10, p. 286, 2016. FELIPUCI, J. P. et al. Biotechnological aspects of microbial pretreatment of lignocellulosic biomass. Biorefineries: a step towards renewable and clean energy, p. 121-150, 2020 FELIPUCI, J. P. Xylan extraction from microbiologically pretreated sugarcane bagasse. 2020. 66f. Dissertation (Master in Biological Sciences (Applied Microbiology)) - São Paulo State University/UNESP, Rio Claro, 2020. FERNANDES, E. S. Effect of granulometry on acid pretreatment, accessibility, exposed lignin surface and enzymatic saccharification of sugarcane bagasse. 2018. Dissertation - São Paulo State University/UNESP, 2018. FLORES-GÓMEZ, C. A. et al. Conversion of lignocellulosic agave residues into liquid biofuels using an AFEXTM-based biorefinery. Biotechnology for Biofuels, v. 11, n. 7, p. 1-18, 2018. FRY, S. C. Postharvest Physiology: Ripening. In: THOMAS BBT-E of APS (Ed.) https://doi.org/10.1016/j.bcdf.2019.100184 48 Encyclopedia of Applied Plant Sciences. Oxford: Elsevier, 2003. p. 794-807. FU, L. H. et al. Purification and characterization of an endo-xylanase from Trichoderma sp., with xylobiose as the main product from xylan hydrolysis. World Journal of Microbiology and Biotechnology, v. 35, p. 171, 2019. GALLETTI, A. M. R.; ANTONETTI, C. Biomass pretreatment: separation of cellulose, hemicellulose and lignin-existing technologies and perspectives. In: ARESTA, M.; DIBENEDETTO, A.; DUMEIGNIL, F. Biorefinery: From Biomass to Chemicals and Fuels. Berlin: De Gruyter, 2012. p. 101-111. GARVEY, M. et al. Cellulases for biomass degradation: comparing recombinant cellulase expression platforms. Trends in Biotechnology, v. 31, n. 10, p. 581-593, 2013. GAUTAM, R. L. et al. Basic Mechanism of Lignocellulose Mycodegradation. In: NAIRAN, R. (Ed.) Mycodegradation of Lignocelluloses. Switzerland: Springer Cham, 2019. p. 1-22. (Series Fungal Biology). GONZÁLEZ-AYÓN, M. A., et al. Enzyme-catalyzed production of potato galactan- oligosaccharides and its optimization by response surface methodology. Materials (Basel), v. 12, p. 1-15, 2019. GOODELL, B. Brown-rot fungal degradation of wood: our evolving view. In: GOODELL, B.; NICHOLAS, D. D.; SCHULTZ, T. P. (Ed.) Wood Deterioration and Preservation: advances in our changing world. Washington: American Chemical Society, 2003. p 97-118. (ACS Symposium Series). GOUMA, S. et al. Studies on Pesticides Mixture Degradation by White Rot Fungi. Journal of Ecological Engineering, v. 20, n. 2, 2019. GUEDES, F. et al. Climate-energy-water nexus in Brazilian oil refineries. International Journal of Greenhouse Gas Control, v. 90, p. 1-11, 2019. GUPTA, P. A Review on Advancement of Pulp and Paper Industry. Journal of Emerging Technologies and Innovative Research, v. 6, n. 6, p. 351-356, 2019. HAKALA, T. K. et al. Evaluation of novel wood-rotting polypores and corticioid fungi for the decay and biopulping of Norway spruce (Picea abies) wood. Enzyme and Microbial Technology, v. 34, p. 255-263, 2004. . HATAKKA, A. Biodegradation of lignin. In: Biopolymers Online. Wiley‑VCH Verlag GmbH & Co., 2005. Available via . Accessed 08 Jun 2020. HATAKKA, A. I.; VARESA, T.; LUNN, T. K. Production of multiple lignin peroxidases by the whiterot fungus Phlebia ochraceofulva. Enzyme and Microbial Technology, v. 15, n. 8, p. 664-669, 1993. http://dx.doi.org/10.12911%2F22998993%2F94918 https://onlinelibrary.wiley.com/doi/abs/10.1002/3527600035.bpol1005.%20Accessed%2008%20Jun%202020 https://onlinelibrary.wiley.com/doi/abs/10.1002/3527600035.bpol1005.%20Accessed%2008%20Jun%202020 49 HATAKKA, A. Lignin-modifying enzymes from selected white-rot fungi: production and role from in lignin degradation. FEMS Microbiology Reviews, v. 13, n. 2-3, p. 125-135, 1994. HATAKKA, A.; HAMMEL, K. E. Fungal biodegradation of lignocelluloses. In: HOFRICHTER, M. (Ed.) Industrial applications. Berlin: Springer, 2011. p 319-340. (The Mycota, v. 10). HIBBETT, D. S.; THORN, R. G. Basidiomycota: Homobasidiomycetes. In: MCLAUGHLIN, D. J.; MCLAUGHLIN, E. G.; LEMKE, P. A. (Ed.) Systematics and evolution. Berlin, Heidelberg: Springer, 2001. p. 121-168 (The Mycota, v. 7). HOLWERDA, E. K. et al. Multiple levers for overcoming the recalcitrance of lignocellulosic biomass. Biotechnology for Biofuels, v. 12, n. 1, p. 1-12, 2019. HU, B. et al. Optimization and Scale-Up of Enzymatic Hydrolysis of Wood Pulp for Cellulosic Sugar Production. BioResources, v. 11, n. 3, p.7242-7257, 2016. HUSAIN, Q. Beta Galactosidases and Their Potential Applications: A Review. Critical Reviews in Biotechnology, v. 20, p. 41-62, 2010. IOELOVICH, M. Y. Models of supramolecular structure and properties of cellulose. Polymer Science, v. 58, n. 6, p. 925-943, 2016. IQBAL, H. M. N. et al. Media optimization for hyper-production of carboxymethyl cellulase using proximally analyzed agro-industrial residue with Trichoderma harzianum under SSF. International Journal of Agricultural Sciences and Veterinary Medicine, v. 4, n. 2, p. 47-55, 2013. JAHNAVI, G. et al. Status of availability of lignocellulosic feed stocks in India: Biotechnological strategies involved in the production of Bioethanol. Renewable and Sustainable Energy Reviews, v. 73, p. 798-820, 2017. JAYASEKARA, S.; RATNAYAKE, R. Microbial cellulases: an overview and applications. In: PASCUAL, A. R. (Ed.) Cellulose. London: IntechOpen, 2019. p. 83- 104. JHA, H. Fungal Diversity and Enzymes Involved in Lignin Degradation. In: NAIRAN, R. (Ed.) Mycodegradation of Lignocelluloses. Switzerland: Springer Cham, 2019. p. 35-49. (Fungal Biology Series). JOHNSTON, Sarah R.; BODDY, Lynne; WEIGHTMAN, Andrew J. Bacteria in decomposing wood and their interactions with wood-decay fungi. FEMS Microbiology Ecology, v. 92, n. 11, p. fiw179, 2016.. JUTURU, V.; WU, J. C. Insight into microbial hemicellulases other than xylanases: a review. Journal of Chemical Technology and Biotechnology, v. 88, n. 3, p. 353- 363, 2013. 50 KAUR, H.; KAPOOR, S.; KAUR, G. Application of ligninolytic potentials of a white-rot fungus Ganoderma lucidum for degradation of lindane. Environmental monitoring and assessment, v. 188, n. 10, p. 588, 2016. . KHOSRAVI, C. et al. Sugar Catabolism in Aspergillus and Other Fungi Related to the Utilization of Plant Biomass. In: SARIASLANI, S.; GADD, G. (Ed.) Advances in Applied Microbiology. Amsterdam: Academic Press, 2015. P. 1-28. KIM, J. et al. Disposable chemical sensors and biosensors made on cellulose paper. Nanotechnology, v. 25, n. 9, p. 092001, 2014. KIRK, T. K.; HIGHLEY, T. L. Quantitative changes in structural components of conifer woods during decay by white-and brown-rot fungi. Phytopathology, v. 63, n. 11, p. 1338-1342, 1973. KIRK, T. K.; MOORE, W. E. Removing lignin from wood with white-rot fungi and digestibility of resulting wood. Wood and Fiber Science, v. 4, n. 2, p. 72-79, 1975. KUMARI, S.; NARAIAN, R. Decolorization of synthetic brilliant green carpet industry dye through fungal co-culture technology. Journal of Environmental Management, v. 180, p. 172-179, 2016. KURAKAKE, M.; IDE, N.; KOMAKI, T. Biological pretreatment with two bacterial strains for enzymatic hydrolysis of office paper. Current microbiology, v. 54, n. 6, p. 424-428, 2007. KUSHWAHA, A. et al. Laccase from white rot fungi having significant role in food, pharma, and other industries. In: BHARATI, S. L.; CHAURASIA, P. K. (Ed.) Research Advancements in Pharmaceutical, Nutritional, and Industrial Enzymology. IGI Global, 2018. p. 253-277. LI, P. P.; WANG, X. J.; CUI, Z. J. Survival and performance of two cellulose- degrading microbial systems inoculated into wheat straw-amended soil. Journal of Microbiology and Biotechnology, v. 22, p. 126-132, 2012. LIU, Shun et al. Systematic classification and phylogenetic relationships of the brown-rot fungi within the Polyporales. Fungal diversity, v. 118, n. 1, p. 1-94, 2023. LOOW, Y-L. et al. Typical conversion of lignocellulosic biomass into reducing sugars using dilute acid hydrolysis and alkaline pretreatment. Cellulose, v. 23, p. 1491- 1520, 2016. LOPES, A. M.; FERREIRA FILHO, E. X.; MOREIRA, L. R. S. An update on enzymatic cocktails for lignocellulose breakdown. Journal of Applied Microbiology, v. 125, p. 632-645, 2018. MADHANRAJ, R. et al. Evaluation of anti-microbial and anti-haemolytic activity of edible basidiomycetes mushroom fungi. Journal of Drug Delivery and Therapeutics, v. 9, n. 1, p. 132-135, 2019. MADHAVI, V.; LELE, S. S. Laccase: properties and applications. BioResources, v. 4, n. 4, p. 1694-1717, 2009. https://doi.org/10.1016/j.jenvman.2016.04.060 https://doi.org/10.22270/jddt.v9i1.2277 51 MALGAS, S. A mini review of xylanolytic enzymes with regards to their synergistic interactions during hetero-xylan degradation. World Journal of Microbiology and Biotechnology, v. 35, p. 187, 2019. MANJU, S.; CHADHA, B. S. Production of Hemicellulolytic enzymes for hydrolysis of lignocellulosic biomass. In: PANDEY, et al. (Ed.) Biofuels: Alternative feedstocks and conversion processes. Academic Press, 2011. p. 203-228. MANNAA, M.; KIM, K. D. Influence of temperature and water activity on deleterious fungi and mycotoxin production during grain storage. Mycobiology, v. 45, n. 4, p. 240-254, 2017. MAO, G. Past, current and future of biomass energy research: a bibliometric analysis. Renewable and Sustainable Energy Reviews, v. 52, p. 1823-1833, 2015. MARTIN, F. et al. The genome of Laccaria bicolor provides insights into mycorrhizal symbiosis. Nature, v. 452, p. 88-92, 2008. MARTÍNEZ, A. T. et al. Studies on wheat lignin degradation by Pleurotus species using analytical pyrolysis. Journal of Analytical and Applied pyrolysis, v. 58, p. 401-411, 2001. MCCABE, N. R.; BILITER, W.; DAWSON, G. Preferential Inhibition of Lysosomal Beta-Mannosidase by Sucrose. Enzyme, v. 43, p. 137-145, 1990. MCCARTHY, B.; LIU, H. B. Food waste and the ‘green’ consumer. Australasian Marketing Journal, v. 25, n. 2, p. 126-132, 2017. MELATI, R. B. et al. Key factors affecting the recalcitrance and conversion process of biomass. BioEnergy Research, v. 12, n. 1, p. 1-20, 2019. MIZUHASHI, H. et al. Investigation of comfort of uniform shirt made of cellulose considering environmental load. In: SAEED, K.; HOMENDA, W. (Ed.) Computer Information Systems and Industrial Management. Springer Cham, 2015. P. 527- 538. (Lecture Notes in Computer Science, v. 9339). MONTORO-G