Food Waste Conversion (Methods and Protocols in Food Science) [1st ed. 2023] 1071633023, 9781071633021

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Methods and Protocols in Food Science

Cristobal N. Aguilar Gonzalez Ricardo Gómez-García Mohammed Kuddus  Editors

Food Waste Conversion

METHODS

AND

PROTOCOLS

IN

Series Editor Anderson S. Sant’Ana University of Campinas Campinas, Brazil

For further volumes: http://www.springer.com/series/16556

FOOD SCIENCE

Methods and Protocols in Food Science series is devoted to the publication of research protocols and methodologies in all fields of food science. Volumes and chapters will be organized by field and presented in such way that the readers will be able to reproduce the experiments in a step-by-step style. Each protocol will be characterized by a brief introductory section, followed by a short aims section, in which the precise purpose of the protocol will be clarified.

Food Waste Conversion Edited by

Cristobal N. Aguilar Gonzalez Facultad de Ciencias Quimicas, Universidad Autonoma de Coahuila, Saltillo, Coahuila, Mexico

Ricardo Gómez-García Universidade Católica Portuguesa, CBQF – Centro de Biotecnologia e Química Fina – Laboratório Associado, Escola Superior de Biotecnologia, Porto, Portugal

Mohammed Kuddus Department of Biochemistry, College of Medicine, University of Hail, Hail, Saudi Arabia

Editors Cristobal N. Aguilar Gonzalez Facultad de Ciencias Quimicas Universidad Autonoma de Coahuila Saltillo, Coahuila, Mexico

Ricardo Go´mez-Garcı´a Universidade Cato´lica Portuguesa CBQF – Centro de Biotecnologia e Quı´mica Fina – Laborato´rio Associado, Escola Superior de Biotecnologia Porto, Portugal

Mohammed Kuddus Department of Biochemistry College of Medicine University of Hail Hail, Saudi Arabia

ISSN 2662-950X ISSN 2662-9518 (electronic) Methods and Protocols in Food Science ISBN 978-1-0716-3302-1 ISBN 978-1-0716-3303-8 (eBook) https://doi.org/10.1007/978-1-0716-3303-8 © The Editor(s) (if applicable) and The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023 This work is subject to copyright. All rights are solely and exclusively licensed by the Publisher, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. The publisher, the authors, and the editors are safe to assume that the advice and information in this book are believed to be true and accurate at the date of publication. Neither the publisher nor the authors or the editors give a warranty, expressed or implied, with respect to the material contained herein or for any errors or omissions that may have been made. The publisher remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. This Humana imprint is published by the registered company Springer Science+Business Media, LLC, part of Springer Nature. The registered company address is: 1 New York Plaza, New York, NY 10004, U.S.A.

Preface to the Series Methods and Protocols in Food Science series is devoted to the publication of research protocols and methodologies in all fields of food science. The series is unique as it includes protocols developed, validated, and used by food and related scientists as well as theoretical basis are provided for each protocol. Aspects related to improvements in the protocols, adaptations, and further developments in the protocols may also be approached. Methods and Protocols in Food Science series aims to bring the most recent developments in research protocols in the field as well as very well established methods. As such the series targets undergraduate, graduate, and researchers in the field of food science and correlated areas. The protocols documented in the series will be highly useful for scientific inquiries in the field of food sciences, presented in such way that the readers will be able to reproduce the experiments in a step-by-step style. Each protocol is characterized by a brief introductory section, followed by a short aims section, in which the precise purpose of the protocol is clarified. Then, an in-depth list of materials and reagents required for employing the protocol is presented, followed by a comprehensive and step-by-step procedures on how to perform that experiment. The next section brings the do’s and don’ts when carrying out the protocol, followed by the main pitfalls faced and how to troubleshoot them. Finally, template results are presented and their meaning/conclusions addressed. The Methods and Protocols in Food Science series fills an important gap, addressing a common complain of food scientists, regarding the difficulties in repeating experiments detailed in scientific papers. With this, the series has a potential to become a reference material in food science laboratories of research centers and universities throughout the world. Campinas, Brazil

Anderson S. Sant’Ana

v

Preface Food loss and waste has been identified as one of the major problems to fulfill the food crisis worldwide and to the development of sustainable food systems. Almost one-third of the food worldwide production is lost or wasted each year, happening in the entire food value chain, either during production, post-harvest, distribution, processing as well as in retailers, and final consumers. There are detrimental effects of food waste on the environment, including the production of greenhouse gases emissions and the dispersal of potential infectious diseases. The food loss and waste could be avoided or recycled in alternative circumstances. Although there are both social and scientific efforts to resolve this global problem, the truth is that much remains to be done. The impact of food loss and waste can be reduced by using innovative multidisciplinary approaches such as integrated food design, valorization, circular bioeconomy, or processing of food waste into value-added products. The lack of knowledge of scientific progress and the missing sustainable technologies within food processes or processing contributes significantly to slow progress of food waste management. In this perspective, the aim of this book is to offer an updated review regarding protocols for food waste conversion through the use of novel food processing tools to food manufacturing processes or to assess the sustainability of new food products with the objective of food waste minimization or the re-valorization of food residues with the purpose to give a comprehensive introduction into methods and procedures all related to the food waste conversion. This book includes a comprehensive reference in the most progressive field of relevant technology to the conversion of food waste, so this book will be combined in the most understandable way of well-established protocols and procedures and will be of interest to professionals, scientists, and academics related to food science and food biotechnology. This book covers 14 chapters that provide an updated knowledge of food processing, food waste management, and production of value-added products from food waste. It will help to reduce food waste and to cope food crisis problems in the world. The chapters also highlight the production of bioactive compounds from agro-industrial waste by using various (bio)technologies. In conclusion, this book is an updated reference in the most progressive field of food waste management and processing by using state-of-the-art technology that will be useful for the food scientists and academics involved in food waste management. Therefore, this book offers scientific knowledge of interest for readers of the food fields, including chemistry, processing, bioengineering, and biotechnology. In the last, but not least, we would like to thank all the authors who have eagerly contributed their chapters in this book. We also express our sincere gratitude to Springer Nature for providing this opportunity. Saltillo, Coahuila, Mexico Porto, Portugal Hail, Saudi Arabia

Cristobal N. Aguilar Gonzalez Ricardo Gomez-Garcı´a Mohammed Kuddus

vii

Contents Preface to the Series . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Preface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Contributors. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

PART I

SUMMARY AND LITERATURE REVIEW ON VALUE-ADDED BIOACTIVE COMPOUNDS FROM FOOD WASTE VALORIZATION

1 Bioactive Compounds from Food and Its By-products: Current Applications and Future Perspectives . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Bianca Rodrigues de Albuquerque, Ru´bia Carvalho Gomes Correˆa, Shirley de Lima Sampaio, and Lillian Barros 2 Food Waste Management Method Through 3R Concept . . . . . . . . . . . . . . . . . . . . Anna Ilina´, R. Ramos-Gonza´lez, R. Arredondo-Valde´s, C. Barrera-Martı´nez, E. Laredo-Alcala´, Patricia M. Albarracin, G. M. Alvarez, and J. L. Martı´nez-Herna´ndez

PART II

v vii xi

3

43

PHYSICO-CHEMICAL AND THERMOCHEMICAL TREATMENTS FOR FOOD WASTE EXPLOITATION

3 Microencapsulation of Bioactive Compounds from Agro-industrial Waste. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 55 ´ ngeles Va´zquez-Nu´n ˜ ez, Mariela R. Michel, Marı´a de los A Mayra Aguilar-Za´rate, Jorge E. Wong-Paz, and Pedro Aguilar-Za´rate 4 Sustainable Extraction of Flavonoids from Agricultural Biomass and Agro-industrial By-products: Natural Deep Eutectic Solvent Synthesis, Extraction, and Chromatographic Analysis. . . . . . . . . . . . . . . . . . . . . . . . 67 Luis Alfonso Jime´nez-Ortega, Josue´ D. Mota-Morales, Laura A. Contreras-Angulo, and Jose´ Basilio Heredia 5 Protocol for the Extraction of Lignin from Brewer’s Spent Grain Using Deep Eutectic Solvents. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 79 Ana C. Cassoni, Patrı´cia Costa, and Manuela Pintado 6 Protocol for Antioxidant Dietary Fiber Determination: Structural Characterization and Quantification . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 85 Taˆnia Braganc¸a Ribeiro, Maria Emilia Brassesco, Diva Santos, and Manuela Pintado 7 Use of Ultrasound Technology for Food Waste Breakdown . . . . . . . . . . . . . . . . . . 97 Martina de la Rosa-Herna´ndez, M. Carmen Gutie´rrez-Sa´nchez, ˜ iz-Ma´rquez, Abigail Reyes-Munguı´a, and Jorge E. Wong-Paz Diana B. Mun 8 Integrated Biorefinery Strategy for Orange Juice By-product Valorization: A Sustainable Protocol to Obtain Bioactive Compounds . . . . . . . . . 113 Ana A. Vilas-Boas, Ricardo Go mez-Garcı´a, De´bora A. Campos, Marta Correia, and Manuela Pintado

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9 Energy Integration of the Hydrothermal Pretreatment of Food Waste in Terms of a Sustainable Biorefinery . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 125 Iosvany Lopez-Sandin, Rosa M. Rodrı´guez-Jasso, and He´ctor A. Ruiz

PART III 10

11

12

13

14

FOOD WASTE AS A CARBON SOURCE FOR FUNGI BASED PROCESSES: BIOACTIVES OBTENTION AND RELEASEMENT

Solid-State Fermentation as Strategy for Food Waste Transformation . . . . . . . . . Israel Bautista-Herna´ndez, Monica L. Cha´vez-Gonza´lez, Arturo Siller Sa´nchez, Karen N. Ramı´rez Guzma´n, Cristian Torres Leo n, Pedro Aguilar Za´rate, Cristobal N. Aguilar Gonzalez, and Deepak Kumar Verma Protocol for the Solid-State Fermentation-Assisted Extraction (SSFAE) of Bioactives from Tomato Waste: The Case of Carotenoids . . . . . . . . . Juanita Y. Mendez-Carmona, Karen N. Ramı´rez-Guzman, Juan A. Ascacio-Valdes, Leonardo Sepulveda, Jose´ Sandoval, and Cristobal N. Aguilar Gonzalez Protocol for the Production of Trichoderma Spores for Use as a Biological Control Agent Through the Revalorization of Agro-industrial Waste . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . ˜ a-Mendoza, Monica L. Chavez-Gonza´lez, Salvador A. Saldan and Cristobal N. Aguilar Gonzalez Submerged Fermentation as a Strategy for the Valorization of Fish By-products to Obtain High-Protein Meals . . . . . . . . . . . . . . . . . . . . . . . . . ˜ o-Hernandez, Anna Marı´a Polanı´a, Alexis Garcı´a, Liliana London Germa´n Bolivar, and Cristina Ramı´rez Monitoring Methods for Anaerobic Digestion of Food Waste: Physicochemical and Molecular Analysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Mario Alberto Yaverino-Gutierrez, Juan Gerardo Flores-Iga, Martha Ine´s Velez-Mercado, Aldo Sosa-Herrera, ˜ o, Marı´a de las Mercedes Esparza-Perusquia, Miriam P. Lue´vanos Escaren Ayerim Y. Herna´ndez Almanza, Fernando Herna´ndez Tera´n, Javier Ulises Herna´ndez-Beltra´n, and Nagamani Balagurusamy

Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Contributors CRISTOBAL N. AGUILAR GONZALEZ • Bioprocesses and Bioproducts Research Group, Food Research Department, School of Chemistry, Universidad Autonoma de Coahuila, Saltillo, Mexico MAYRA AGUILAR-ZA´RATE • Facultad de Ciencias Quı´micas-CIEP, Universidad Autonoma de San Luis Potosı´, San Luis Potosı´, San Luis Potosı´, C. P, Mexico PEDRO AGUILAR-ZA´RATE • Departamento de Ingenierı´as, Tecnologico Nacional de Me´xico/ Instituto Tecnologico de Ciudad Valles, Ciudad Valles, San Luis Potosı´, C. P, Mexico PATRICIA M. ALBARRACIN • CEDIA, FRT, UTN, Tucuma´n, Argentina G. M. ALVAREZ • Instituto Cubano de las Investigaciones de los Derivados de la can˜a de azu´car, La Habana, Cuba R. ARREDONDO-VALDE´S • Department of NanoBioscience, School of Chemistry, Universidad Autonoma de Coahuila, Saltillo, Mexico JUAN A. ASCACIO-VALDES • Bioprocesses and Bioproducts Research Group. Food Research Department, School of Chemistry, Universidad Autonoma de Coahuila, Saltillo, Mexico NAGAMANI BALAGURUSAMY • Laboratorio de Biorremediacion, Facultad de Ciencias Biologicas, Ciudad Universitaria de la Universidad Autonoma de Coahuila, Torreon, Coahuila, Mexico C. BARRERA-MARTI´NEZ • Conservation of Biodiversity and Ecology of Coahuila Research Center (CICBEC), Universidad Autonoma de Coahuila, Saltillo, Mexico LILLIAN BARROS • Centro de Investigac¸a˜o de Montanha (CIMO), Instituto Polite´cnico de Braganc¸a, Campus de Santa Apolonia, Braganc¸a, Portugal; Laboratorio Associado para a Sustentabilidade e Tecnologia em Regio˜es de Montanha (SusTEC), Instituto Polite´cnico de Braganc¸a, Braganc¸a, Portugal ISRAEL BAUTISTA-HERNA´NDEZ • Bioprocesses and Bioproducts Research Group, Food Research Department, School of Chemistry, Universidad Autonoma de Coahuila, Saltillo, Mexico; Nanobioscience Research Group. Food Research Department, School of Chemistry, Universidad Autonoma de Coahuila, Saltillo, Mexico GERMA´N BOLIVAR • Faculty of Natural and Exact Sciences, Biology Department, MIBIA Group, Universidad del Valle, Cali, Valle del Cauca, Colombia MARI´A EMILIA BRASSESCO • Universidade Catolica Portuguesa, CBQF – Centro de Biotecnologia e Quı´mica Fina – Laboratorio Associado, Escola Superior de Biotecnologia, Porto, Portugal ANA C. CASSONI • Universidade Catolica Portuguesa, CBQF – Centro de Biotecnologia e Quı´mica Fina – Laboratorio Associado, Escola Superior de Biotecnologia, Porto, Portugal MO´NICA L. CHA´VEZ-GONZA´LEZ • Bioprocesses and Bioproducts Research Group, Food Research Department, School of Chemistry, Universidad Autonoma de Coahuila, Saltillo, Mexico; Nanobioscience Research Group. Food Research Department, School of Chemistry, Universidad Autonoma de Coahuila, Saltillo, Mexico LAURA A. CONTRERAS-ANGULO • Centro de Investigacion en Alimentacion y Desarrollo, A.C, Culiaca´n, Sinaloa, Mexico RU´BIA CARVALHO GOMES CORREˆA • Centro de Investigac¸a˜o de Montanha (CIMO), Instituto Polite´cnico de Braganc¸a, Campus de Santa Apolonia, Braganc¸a, Portugal; Laboratorio Associado para a Sustentabilidade e Tecnologia em Regio˜es de Montanha (SusTEC),

xi

xii

Contributors

Instituto Polite´cnico de Braganc¸a, Braganc¸a, Portugal; Postgraduate Program in Clean Technologies, Cesumar University – UNICESUMAR, Maringa, Parana, Brazil; Cesumar Institute of Science, Technology and Innovation – ICETI, Maringa, Parana´, Brazil PATRI´CIA COSTA • Universidade Catolica Portuguesa, CBQF – Centro de Biotecnologia e Quı´mica Fina – Laboratorio Associado, Escola Superior de Biotecnologia, Porto, Portugal BIANCA RODRIGUES DE ALBUQUERQUE • Centro de Investigac¸a˜o de Montanha (CIMO), Instituto Polite´cnico de Braganc¸a, Campus de Santa Apolonia, Braganc¸a, Portugal; Laboratorio Associado para a Sustentabilidade e Tecnologia em Regio˜es de Montanha (SusTEC), Instituto Polite´cnico de Braganc¸a, Braganc¸a, Portugal; REQUIMTE/LAQV, Department of Chemical Sciences, Faculty of Pharmacy, University of Porto, Porto, Portugal MARTINA DE LA ROSA-HERNA´NDEZ • Faculty of Professional Studies Huasteca Zone, Autonomous University of San Luis Potosı´, Ciudad Valles, San Luis Potosı´, Mexico MARI´A DE LAS MERCEDES ESPARZA-PERUSQUIA • Facultad de Medicina, Departamento de Bioquı´mica, Universidad Nacional Autonoma de Me´xico, Ciudad de Me´xico, Mexico SHIRLEY DE LIMA SAMPAIO • Centro de Investigac¸a˜o de Montanha (CIMO), Instituto Polite´ cnico de Braganc¸a, Campus de Santa Apolonia, Braganc¸a, Portugal; Laboratorio Associado para a Sustentabilidade e Tecnologia em Regio˜es de Montanha (SusTEC), Instituto Polite´cnico de Braganc¸a, Braganc¸a, Portugal ´ NGELES VA´ZQUEZ-NU´N˜EZ • Facultad de Estudios Profesionales Zona MARI´A DE LOS A Huasteca, Universidad Autonoma de San Luis Potosı´, Ciudad Valles, San Luis Potosı´, C. P, Mexico JUAN GERARDO FLORES-IGA • Laboratorio de Biorremediacion, Facultad de Ciencias Biologicas, Ciudad Universitaria de la Universidad Autonoma de Coahuila, Torreon, Coahuila, Mexico ALEXIS GARCI´A • Faculty of Engineering, School of Food Engineering, Universidad del Valle, Tulua´, Valle del Cauca, Colombia RICARDO GO´MEZ-GARCI´A • Universidade Catolica Portuguesa, CBQF – Centro de Biotecnologia e Quı´mica Fina – Laboratorio Associado, Escola Superior de Biotecnologia, Porto, Portugal M. CARMEN GUTIE´RREZ-SA´NCHEZ • Faculty of Professional Studies Huasteca Zone, Autonomous University of San Luis Potosı´, Ciudad Valles, San Luis Potosı´, Mexico JOSE´ BASILIO HEREDIA • Centro de Investigacion en Alimentacion y Desarrollo, A.C, Culiaca´n, Sinaloa, Mexico AYERIM Y. HERNA´NDEZ ALMANZA • Laboratorio de Biorremediacion, Facultad de Ciencias Biologicas, Ciudad Universitaria de la Universidad Autonoma de Coahuila, Torreon, Coahuila, Mexico JAVIER ULISES HERNA´NDEZ-BELTRA´N • Laboratorio de Biorremediacion, Facultad de Ciencias Biologicas, Ciudad Universitaria de la Universidad Autonoma de Coahuila, Torreon, Coahuila, Mexico ANNA ILINA´ • Department of NanoBioscience, School of Chemistry, Universidad Autonoma de Coahuila, Saltillo, Mexico LUIS ALFONSO JIME´NEZ-ORTEGA • Centro de Investigacion en Alimentacion y Desarrollo, A. C, Culiaca´n, Sinaloa, Mexico E. LAREDO-ALCALA´ • Conservation of Biodiversity and Ecology of Coahuila Research Center (CICBEC), Universidad Autonoma de Coahuila, Saltillo, Mexico CRISTIAN TORRES LEO´N • Reaserch Center and Ethnobiological Garden (CIJE), Universidad Autonoma de Coahuila, Viesca, Coahuila, Mexico

Contributors

xiii

LILIANA LONDON˜O-HERNANDEZ • BIOTICS Group, School of Basic Sciences, Technology and Engineering, Universidad Nacional Abierta y a Distancia – UNAD Colombia, Bogota´, Colombia IOSVANY LO´PEZ-SANDIN • Biorefinery Group, Food Research Department, School of Chemistry, Autonomous University of Coahuila, Saltillo, Coahuila, Mexico MIRIAM P. LUE´VANOS ESCAREN˜O • Laboratorio de Biorremediacion, Facultad de Ciencias Biologicas, Ciudad Universitaria de la Universidad Autonoma de Coahuila, Torreon, Coahuila, Mexico J. L. MARTI´NEZ-HERNA´NDEZ • Department of NanoBioscience, School of Chemistry, Universidad Autonoma de Coahuila, Saltillo, Mexico JUANITA Y. MENDEZ-CARMONA • Bioprocesses and Bioproducts Research Group. Food Research Department, School of Chemistry, Universidad Autonoma de Coahuila, Saltillo, Mexico MARIELA R. MICHEL • Departamento de Ingenierı´as, Tecnologico Nacional de Me´xico/ Instituto Tecnologico de Ciudad Valles, Ciudad Valles, San Luis Potosı´, C. P, Mexico JOSUE´ D. MOTA-MORALES • Centro de Fı´sica Aplicada y Tecnologı´a Avanzada (CFATA), Universidad Nacional Autonoma de Me´xico (UNAM), Quere´taro, QRO, Mexico DIANA B. MUN˜IZ-MA´RQUEZ • Faculty of Professional Studies Huasteca Zone, Autonomous University of San Luis Potosı´, Ciudad Valles, San Luis Potosı´, Mexico MANUELA PINTADO • Universidade Catolica Portuguesa, CBQF – Centro de Biotecnologia e Quı´mica Fina – Laboratorio Associado, Escola Superior de Biotecnologia, Porto, Portugal ANNA MARI´A POLANI´A • Faculty of Engineering, School of Food Engineering, Universidad del Valle, Tulua´, Valle del Cauca, Colombia CRISTINA RAMI´REZ • Faculty of Natural and Exact Sciences, Biology Department, MIBIA Group, Universidad del Valle, Cali, Valle del Cauca, Colombia KAREN N. RAMI´REZ GUZMA´N • Center for Interdisciplinary Studies and Research, Universidad Autonoma de Coahuila, Saltillo, Me´xico R. RAMOS-GONZA´LEZ • Department of NanoBioscience, School of Chemistry, Universidad Autonoma de Coahuila, Saltillo, Mexico ABIGAIL REYES-MUNGUI´A • Faculty of Professional Studies Huasteca Zone, Autonomous University of San Luis Potosı´, Ciudad Valles, San Luis Potosı´, Mexico TAˆNIA BRAGANC¸A RIBEIRO • Universidade Catolica Portuguesa, CBQF – Centro de Biotecnologia e Quı´mica Fina – Laboratorio Associado, Escola Superior de Biotecnologia, Porto, Portugal ROSA M. RODRI´GUEZ-JASSO • Biorefinery Group, Food Research Department, School of Chemistry, Autonomous University of Coahuila, Saltillo, Coahuila, Mexico HE´CTOR A. RUIZ • Biorefinery Group, Food Research Department, School of Chemistry, Autonomous University of Coahuila, Saltillo, Coahuila, Mexico SALVADOR A. SALDAN˜A-MENDOZA • Bioprocesses and Bioproducts Research Group, Food Research Department, School of Chemistry, Universidad Autonoma de Coahuila, Saltillo, Mexico ARTURO SILLER SA´NCHEZ • Bioprocesses and Bioproducts Research Group, Food Research Department, School of Chemistry, Universidad Autonoma de Coahuila, Saltillo, Mexico; Nanobioscience Research Group. Food Research Department, School of Chemistry, Universidad Autonoma de Coahuila, Saltillo, Mexico JOSE´ SANDOVAL • Bioprocesses and Bioproducts Research Group. Food Research Department, School of Chemistry, Universidad Autonoma de Coahuila, Saltillo, Mexico

xiv

Contributors

DIVA SANTOS • Universidade Catolica Portuguesa, CBQF – Centro de Biotecnologia e Quı´mica Fina – Laboratorio Associado, Escola Superior de Biotecnologia, Porto, Portugal LEONARDO SEPULVEDA • Bioprocesses and Bioproducts Research Group. Food Research Department, School of Chemistry, Universidad Autonoma de Coahuila, Saltillo, Mexico ALDO SOSA-HERRERA • Laboratorio de Biorremediacion, Facultad de Ciencias Biologicas, Ciudad Universitaria de la Universidad Autonoma de Coahuila, Torreon, Coahuila, Mexico FERNANDO HERNA´NDEZ TERA´N • Laboratorio de Biorremediacion, Facultad de Ciencias Biologicas, Ciudad Universitaria de la Universidad Autonoma de Coahuila, Torreon, Coahuila, Mexico MARTHA INE´S VELEZ-MERCADO • Laboratorio de Biorremediacion, Facultad de Ciencias Biologicas, Ciudad Universitaria de la Universidad Autonoma de Coahuila, Torreon, Coahuila, Mexico DEEPAK KUMAR VERMA • Agricultural and Food Engineering Department, Indian Institute of Technology Kharagpur, Kharagpur, India JORGE E. WONG-PAZ • Departamento de Ingenierı´as, Tecnologico Nacional de Me´xico/ Instituto Tecnologico de Ciudad Valles, Ciudad Valles, San Luis Potosı´, C. P, Mexico; Faculty of Professional Studies Huasteca Zone, Autonomous University of San Luis Potosı´, Ciudad Valles, San Luis Potosı´, Mexico MARIO ALBERTO YAVERINO-GUTIERREZ • Laboratorio de Biorremediacion, Facultad de Ciencias Biologicas, Ciudad Universitaria de la Universidad Autonoma de Coahuila, Torreon, Coahuila, Mexico PEDRO AGUILAR ZA´RATE • Engineering Department, Tecnologico Nacional de Me´xico/ I. T. de Ciudad Valles, Ciudad Valles, San Luis Potosı´, Mexico

Part I Summary and Literature Review on Value-Added Bioactive Compounds from Food Waste Valorization

Chapter 1 Bioactive Compounds from Food and Its By-products: Current Applications and Future Perspectives Bianca Rodrigues de Albuquerque, Ru´bia Carvalho Gomes Correˆa, Shirley de Lima Sampaio, and Lillian Barros Abstract Foods are important for the supply of essential nutrients to the body, but they can also be sources of bioactive molecules, such as phenolic compounds, alkaloids, glucosinolates, and terpenes, which can exert beneficial bioactivities for the proper functioning of the body. In addition, these molecules have other interesting technological properties for the development of new products in several areas of the industry. On the other hand, food production and processing are responsible for generating a high volume of by-products and bio-residues with no commercial value and high impact on the environment. Nonetheless, food by-products can be a source of bioactive compounds of interest, which has increased the number of studies aimed at obtaining bioactive compounds with high added value from agro-industry residues. Thus, the development of green extraction technologies to obtain this compound efficiently and sustainably has become essential for the incorporation of these recovered bioactive molecules in the industry. This chapter aims to present the main bioactive compounds present in foods and their by-products, as well as to summarize the main pharmaceutical and food applications of these compounds and to introduce the new green technologies applied to the recovery of these compounds. Key words Phytochemicals, Nutraceuticals, Antioxidant, Natural food additives, Cosmetic ingredients, Circular economy

1

Introduction Albeit food production is sufficient to feed the global population, millions of people still endure hunger and undernourishment worldwide. According to the Food and Agriculture Organization (FAO), more than one-third of all food produced for human consumption is lost along the food chain, generating economic losses, waste of natural resources, and environmental damage, as a substantial part of this biomass is destinated to sanitary landfills and waste incineration [1]. Fruits and vegetables correspond to 45% of the total food waste volume [1]. Of the 1.4 million metric tons of fruits processed by year worldwide, 25–30% is transformed into

Cristobal N. Aguilar Gonzalez et al. (eds.), Food Waste Conversion, Methods and Protocols in Food Science, https://doi.org/10.1007/978-1-0716-3303-8_1, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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4

Bianca Rodrigues de Albuquerque et al.

waste or by-products. For instance, the generation of bio-residues from citrus fruit production and processing can achieve 60%, whereas for grapes, it can correspond to 20–40% [2, 3]. The discarded biomass is composed of peels, seeds, roots, and pomace, fruit parts that commonly concentrate higher amounts of bioactive compounds than the corresponding pulps [2]. Such materials could be used to improve human nutrition, particularly those rich in vitamins, minerals, fiber, oils, and bioactive compounds (BCs) with functional actions. Table 1 presents some examples of food production and their by-products rich in BC. Food bioactive compounds (FBCs) are being prospected for the treatment and prevention of manifold human ailments. These molecules hold diverse acting paths, among which are as follows: (1) interaction with DNA, proteins, and other molecules generating several desired organism outcomes, (2) free radical inactivation capabilities, and (3) positive transformation on the configuration and metabolic performance of the gut microbiota [4]. FBCs consist of a magnificent pool of chemicals for the obtainment of novel medicines, nutraceuticals, food ingredients, and food packing, meeting consumers growing interest in health promotion and natural and green food products, besides a sustainable food system [5]. Given this context, this work aimed at providing an up-to-date critical review of nutritional, therapeutical, technological, and environmental aspects involved in the exploitation of BC and bioactivities from food and its by-products. Recent evidence on the use of FBC in the treatment of diseases and the development of new drugs has been compiled and discussed. However, special attention was devoted to efforts aiming at the following: (1) applications of food and food waste-derived BC in foodstuff and food packaging and (2) valorization of food residues as sources of FBC, highlighting trends and perspectives of exploiting residual biomass within the upcycling concept. For this purpose, scientific articles and patents published during the last 3 years (2019 to now) were prioritized.

2

Bioactive Compounds in Foods and Food By-products

2.1 Phenolic Compounds

Phenolic compounds (PCs) are secondary metabolites synthesized by two metabolism pathways of plants, namely, by shikimic and melanic acid. This class of BC can be found in diverse matrices, such as vegetables, grains, fruits, fungi, and algae, among others. More than 8000 PC have been identified in nature, and, due to the diversity of these phytochemicals, they can be subclassified in different ways. Usually, PCs are divided into flavonoids and non-flavonoids [6, 7]. Flavonoids are compounds formed by 15 carbon units, 2 benzene rings, and a heterocyclic pyran ring. This group of PCs includes flavanols (main compounds, kaempferol, myricetin, and

Wine

70

55.85

3.1

98

Grape Vines vinifera L.

Guava Psidium guajava L.

Olive Olea europaea L.

Onion Allium cepa L. nd

Olive oil

Juice and nectar

Coffee beans

Juice and cider

Coffee Coffea spp. 10

Apple Malus spp.

Banana flour and chips

81

Foods

Main derivative products

Banana Musa spp. 50,000

Annual production (million tons)

5

50

8

15–20

50

35–50

25–30

Amount of by-product (%)

Skin

[71]

[69, 70]

References

Flavonoid: Quercetin

(continued)

[77]

[76]

[75] PAs: cinnamaldehyde caffeic, cinnamic, gallic, and vanillic acids Flavonoids: Kaempferol, catechin, and quercitrin Tannins and saponins

Flavonoids: anthocyanins, flavan-3- [74] ols, kaempferol, quercetin, myricetin Stilbene: resveratrol

PA: chlorogenic, caffeic, ferulic and [72, 73] coumaric acids, caffeine

Flavonoids: Kaempferol, isoquercetin, quercetin, myricetin, naringenin PA: Ferulic, cinnamic, sinapic, p-coumaric acids

Flavonoids: catechin, epicatechin, procyanidin B2, and hyperin PA: caffeic and chlorogenic acids

Main bioactive compounds

Olive biomass, mill Flavonoids: Hydroxytyrosol, water, leaf, and tyrosol, and oleuropein seeds

Peels and seeds

Skins, stems, and seeds

Coffee pulp

Peels

Pomace

Main by-products

Table 1 Global annual production of important crops, their main derivative products, by-products, and identified FBC, reported in the past 3 years Bioactive Compounds from Food and Its By-products: Current Applications. . . 5

1949.3

680

Sugarcane Saccharum officinarum L.

Rice Oryza spp.

nd not determined, PA phenolic acid.

130 Tomato Lycopersicon esculentum Mill.

368

Annual production (million tons)

Potato Solanum tuberosum L.

Foods

Table 1 (continued)

Pulp, sauce, and peeled tomato

Rice milling

Alcohol and sugar

French fries, chips, hash browns, puree, and frozen food

Main derivative products

8

40

30

2–4

Amount of by-product (%)

Pomace and peels

Rice bran

Bagasse

Peels

Main by-products

[78]

References

[80]

[81]

PAs: caffeic and ferulic phenolic acids PA-triterpene: γ-oryzanol Terpenes: lycopene and β-carotene

[3, 79] PAs: 3-O-caffeoyquinic, 5-Ocafferoylquinic, and 3-O-feruloyl quinic acids Flavonoids: isoorientin Saponins

PAs: Chlorogenic and caffeic acids

Main bioactive compounds

6 Bianca Rodrigues de Albuquerque et al.

Bioactive Compounds from Food and Its By-products: Current Applications. . .

7

quercetin), flavones (main compounds, luteolin and apigenin), isoflavones (main compounds, daidzein and genistein), flavanols or flavan-3-ols (main compounds, catechin and epicatechins), flavanones (main compounds, naringenin and hesperidin), and anthocyanins (main compounds, cyanidin, delphinidin, petunidin, malvidin, pelargonidin, and peonidin). In turn, non-flavonoids are phenolic acids (PAs), stilbenes, lignans, and tannins, among others. PAs are characterized by a phenol ring, benzoic, cinnamic, caffeic, vanillic, gallic, p-coumaric, and ferulic acids being the most common examples. Stilbenes are formed by 14 carbon atoms and 2 benzene rings linked by a double bond. Resveratrol is the most investigated stilbene [6]. Lignans are phytoestrogenic compounds structurally constituted by the combination of two phenylpropanoids. Secoisolariciresinol, matairesinol, medioresinol, pinoresinol, lariciresinol, and syringaresinol are some examples of lignans [7]. Tannins comprise a group of complex PC, better classified into condensed tannins and hydrolysable tannins. Simply put, condensed tannins, also known as proanthocyanidins, are those formed by the condensation of at least two flavan-3-ols monomers. Hydrolyzable tannins are complex polyesters formed by phenolic acids and sugar molecules, which, when subjected to acidic conditions or enzymatic action, undergo hydrolysis, releasing phenolic acids. The most common hydrolysable tannins are those derived from gallic acid (gallotannins) and ellagic acid (ellagitannins) [7]. PCs are extensively investigated due to their high incidence in natural sources and their multiple bioactivities. Many of these compounds have more than two bioactivities, including antioxidant, anti-inflammatory, antiproliferative, and antimicrobial, as shown in Table 2. Bioactive properties of PC have been explored for the alternative and adjuvant treatments for some diseases and in the development of functional foods and cosmetic ingredients, among others [6, 7]. 2.2

Terpenoids

Terpenes are secondary metabolites produced in plants and some microorganisms. Some compounds belonging to this class have been applied in medicinal treatments. Terpenes can be classified according to the number of isoprenoids (C5) presented in their structure in hemiterpenes (C5), monoterpenes (C10), sesquiterpenes (C15), diterpene (C20), sesterterpenes (C25), triterpenes (C30), and tetraterpenes (C40) [8]. Several terpenes have displayed important biological activities in manifold models, such as antiinflammatory, anticancer, antioxidant, and antiviral effects, as presented in Tables 2 and 3 [8, 9]. Artemisinin, for instance, has been used as an antimalarial drug, while paclitaxel, a diterpenoid obtained from bark and wood of Pacific yew, has been applied as an anticancer drug [9]. In the case of tetraterpenes, namely, carotenoids, the major investigated biological effect has been cardiovascular protection [10].

Phenolic compounds

Brazilian quinoa (BRS Piabiru) Flavonoids: quercetin and (Chenopodium quinoa Willd) kaempferol derivatives

Bambara groundnut [Vigna subterranean (L.) Verdc.] seed and hulls

Antioxidant activity

Artichoke (Cynara scolymus L.) PAs: 1-O-, 3-O-, 4-O-, and floral stems 5-O-caffeoylquimic acids Flavonoids: luteolin and apigenin derivatives Tannins: procyanidin dimer

Antioxidant activity

Antioxidant activity Tyrosine inhibition

Anti-inflammatory activity Antibacterial activity

Antibacterial activity

PAs: 5-O-caffeoylquinic and 4-O-caffeoylquinic acids Flavonoids: epicatechin, phloretin-2-Oxyloglucoside, phlorizin

Apple (Malus domestica Borkn.) pomace

Bioactivities

Main FBC

Source

TBARS OxHLIA

DPPH Tyrosinase inhibition activity

[83]

[84]

IC50 = 0.87 mg/mL IC50 = 0.45 mg/mL IC50 = 764 ug/mL IC50 Δt60 = 59 ug/mL

[82]

ABTS DPPH FRAP Albumin denaturation assay Staphylococcus aureus

IC50 = 185.62–2019.7 μg/mL IC50 = 588.09–1440.24 μg/mL IC50 = 184.58–491.68 μg/mL Inhibition >98% at 1 mg/mL concentration MIC = 1–1.5 mg/mL; MBC = 1.5–2.0 mg/mL Enterococcus faecium MIC = 1–1.5 mg/mL; MBC = 1.5–2.0 mg/mL Streptococcus agalactiae MIC = 1–1.5 mg/mL; MBC = 1.5–2.0 mg/mL Salmonella typhimurium MIC = 1–1.5 mg/mL; MBC = 1.5–2.0 mg/mL Escherichia coli MIC = 1–1.5 mg/mL; MBC = 1–1.5 mg/mL Candida albicans MIC = 1 mg/mL; MBC = 1–2 mg/mL

References [70]

Outcomes

Propionibacterium acnes MIC = 2.5 mg/mL

Type of assay

Table 2 Recent reports on in vitro evaluation of biological properties of phytochemicals found in food and food by-products

Antioxidant activity Anti-inflammatory activity

PC

Flavanone: neohesperidin and hesperetin

Antioxidant activity Flavonoids: cyanidin derivatives, peonidin derivatives, quercetin derivatives, myricetin derivatives Terpenes: ursolic and oleanolic acids

Antidiabetic activity Flavonoids: peonidin-3-Oacetylglucoside, quercetin-3O-glucuronide and isorhamnetin-3-O-glucoside, and catechin

Cinnamon [Cinnamomum cassia (L.) J. Presl] leaf residue

Citrus peels

Cranberry (Vaccinium macrocarpon Ait.)

Grape (Vitis vinifera L. var. MErlot) pomace

Anti-ageing activity

Antiproliferative activity

PAs: 4-O-coffeoylquinic and p-coumaroyl hexoside Flavonoids: (iso)liquiritigeninO-hexoside, apigenin-Ohexoside, myricetin, and ellagic acid derivatives

Camu camu [Myrciaria dubia (Kunth) McVaugh] peels

34.17% inhibition 64.93% inhibition

GI50 = 180, 304, 238, and 279 ug/mL, respectively

[88]

# salivary α-amylase activity α-Amylase inhibition

(continued)

[87]

IC50 = 3.47 × 10-4 μg/mL

[86]

[85]

[24]

DPPH

Saccharomyces cerevisiae " CLS, # ROS production chronological lifespan (CLS)

DPPH Albumin denaturation assay

HeLa, NCI-H460, HepG2, and MCF-7 cells

Main FBC

GI50 = 300, 278, and 258 μg/ mL respectively MIC = 20 mg/mL MIC = 20 mg/mL MIC = 20 mg/mL MIC = 20 mg/mL MIC = 10 mg/mL MIC = 10 mg/mL MIC = 10 mg/mL MIC = 10 mg/mL

MCF-7, HeLa, HepG2 Escherichia coli Klebsiella pneumoniae Morganella morganii Proteus mirabilis Pseudomonas aeruginosa Enterococcus faecalis Listeria monocytogenes MRSA

Antibacterial activity

Staphylococcus aureus

MIC = 1 mg/mL; MBC = 2 mg/ [25] mL Bacillus cereus MIC = 2 mg/mL; MBC = 4 mg/mL Listeria monocytogenes MIC = 2 mg/mL; MBC = 4 mg/mL Escherichia coli MIC = 1 mg/mL; MBC = 2 mg/mL Salmonella typhimurium MIC = 2 mg/mL; MBC = 4 mg/mL Enterobacter cloacae MIC = 2 mg/mL; MBC = 4 mg/mL

Flavonoids: Quercetin derivatives, epicatechin, and B-type (epi)catechin

Kiwi (Actinidia deliciosa cv. Hayward) peels

[90]

[89]

IC50 = 2.07 and 0.82 μg/mL IC50 = 299 μg/mL

TBARS, OxHLIA NO inhibition

IC50 = 204 μg/mL; IC50 Δt60 = 42 μg/mL

References

Outcomes

Type of assay

TBARS, OxHLIA

Antibacterial activity

Antioxidant activity Anti-inflammatory activity Antiproliferative activity

Bioactivities

Antioxidant activity Juc¸ara (Euterpe edulis Martius) PAs: Ferulic, caffeic acid peels Flavonoids: Taxifolin, apigenin, kaempferol, hispidulin, quercetin, and cyanidin derivatives

Jaboticaba [Myrciaria Flavonoids: Quercitrin and Jaboticaba (Vell.) Berg] peels anthocyanins Tannins: Bis-HHDP-glucose, trisgalloyl-HHPD-glucose, galloyl-bis-HHDP-glucose, pentagalloyl glucose

Source

Table 2 (continued)

TBARS, OxHLIA NCI-H460, HepG2, MCF-7, and HeLa cells

Antioxidant activity Antiproliferative activity

Potato (Solanum tuberosum L.) Anthocyanins, phenolic acids peels

# ROS production

MIC = 0.5 mg/mL; MFC = 1 mg/mL MIC = 1 mg/mL; MFC = 2 mg/mL MIC = 0.5 mg/mL; MFC = 1 mg/mL MIC = 1 mg/mL; MFC = 2 mg/mL MIC = 1 mg/mL; MBC = 2 mg/mL GI50 = 180, 304, 238, and 279 μg/mL, respectively

Staphylococcus aureus

[91]

[22]

(continued)

IC50 = 26–230 ug/mL and IC50 [92] Δt60 = 16–294 μg/mL GI50 = 69–281, 79–365, 51–315, and 49–346 μg/mL

MIC = 0.11 mg/mL; MBC = 0.38 mg/mL Staphylococcus epidermis MIC = 0.02 mg/mL; MBC = 1.12 mg/mL Micrococcus kristinae MIC = 0.1 mg/mL; MBC = 0.28 mg/mL Enterococcus faecalis MIC = 3 mg/mL Escherichia coli MIC = 0.38 mg/mL; MBC = 0.75 mg/mL Pseudomonas aeruginosa MIC = 1.5 mg/mL

Antibacterial activity

Lignans: Isohydroxymatairesinol and punicatannin C Flavonoids: Phloretin, quercetin, indolamine PA: Coutaric acid

Mouse dermal fibroblasts treated with epinephrine

Pomegranate (Punica granatum L.) husk

Antioxidant, antiinflammatory, and antiaging activity

PA: Hydroxytyrosol PC: Oleuropein

HeLa, NCI-H460, HepG2, and MCF-7 cells

Penicillium aurantiogriseum Trichoderma viride

Aspergillus versicolor

Aspergillus Niger

Aspergillus ochraceus

Olive (Olea europaea L.) oil

Antiproliferative activity

Antifungal activity

Antioxidant activity

PAs: Gallic and caffeic acids Stilbene: Resveratrol

Terpene: Ganoresinoid A

Sugarcane (Saccharum officinarum L.) bagasse

Ganoderma resinaceum Boud.

Type of assay

Outcomes

SH-SY5Y cells treated with H2O2 LPS-induced BV-2 microglia cells

IC50 = 5.41 μM

92.27 cell viability

[94]

IC50 = 4.86, 3.21, 1.76, and 1.36 [93] respectively.

[21]

References

nd not declared, PA phenolic acid, PCs phenolic compounds, ABTS 2,2′-azino-bis(3-ethylbenzothiazoline-6-sulfonic acid), DPPH 2,2-diphenyl-1-picrylhydrazyl, FRAP ferric reducing antioxidant power, CUPRAC cupric ion reducing antioxidant capacity assay, TBARS thiobarbituric acid reactive substances, OxHLIA oxidative hemolysis inhibition assay, NO nitrogen oxide, NCI-H460 small cell lung cancer cells, HepG2 human liver cancer cell line, MCF-7 human breast cancer cell line, HeLa human cervical cancer cell line, SH-SY5Y cloned subline of a neuroblastoma cell line, IC50 half maximal inhibitory concentration, GI50 concentration causing 50% cell growth inhibition, MIC minimum inhibitory concentration, MBC minimum bactericidal concentration, MFC minimum fungicidal concentration, MRSA methicillin-resistant Staphylococcus aureus, LPS lipopolysaccharide, # decrease, " increase

Antioxidant and antiaging activities Anti-inflammatory activity

DPPH, ABTS, FRAP, and CUPRAC

DPPH 45.65% inhibition Tyrosinase enzyme 8.95% inhibition inhibition test α-Glucosidase inhibition α-Glucosidase activity 77.12% inhibition assay α-Amylase inhibition α-Amylase activity assay 70.94% inhibition

Antioxidant activity L-tyrosine inhibition

Alkaloids: Caulerpin and caulerpenin PC: Phenols, flavonoids, and tannins Terpenes: Sesquiterpenoids, diterpenoids, sitosterol Saponins

Sea grape [Caulerpa racemosa (Forssk) J.Agardh]

Bioactivities

Main FBC

Source

Table 2 (continued)

Type 2 diabetes Clinical trial

Obesity

Antioxidant and antihyperglycemic Antioxidant and antidiabetic activities Anti-obesity

Antioxidant and antiglycemic activities

Phenolic compounds

PAs: Ellagic and gallic acids. Tannins: Punicalagin and punicalin

Onion (Allium cepa L.) peel

Pomegranate (Punica granatum L.) peel

Coffee (Coffea PA: Chlorogenic acid arabica L.) pulp Alkaloids: Caffeine and trigonelline Diterpenes

Jabuticaba [Myrciaria Jaboticaba

Flavonoids: Cyanidin-3-Oglucoside

Type 2 diabetes Animal model

Antidiabetic activity

Flavonoids: Peonidin-3-Oacetylglucoside, quercetin3-O-glucuronide and isorhamnetin-3-Oglucoside, and catechin

Grape (Vines vinifera L. var. Merlot) residues

Metabolic syndrome

[88]

[32]

[96]

#starch hydrolysis and glucose levels

# blood glucose levels, malondialdehyde concentration # triglycerides, LDL-C, "HDL-C

(continued)

[97]

#triglycerides and starch absorption

[95]

# TG, LDL-C, and TC,

Animal model

[82]

#LDL-C, TG, atherogenic index of plasma

[73] # weight body, abdominal fat, blood pressure, TG; " glucose tolerance, microbiota gut

References

Outcomes

Animal model

Type 2 diabetes Animal model

Type 2 diabetes Clinical trial

Type 2 diabetes Animal model

Antioxidant activity

Antidiabetic activity

PAs: 1-O-, 3-O-, 4-O-, and 5-O-caffeoylquimic acids Flavonoids: Luteolin and apigenin derivatives Tannins: Procyanidin dimer

Artichoke (Cynara scolymus L.) floral stems

Assay model

Disorder

Bioactivities

Flavonoids: Anthocyanins Blueberries (Vaccinium virgatum Ait. and Vaccinium corymbosum L.)

Main FBC

Source

Table 3 Applications of bioactives from food and food by-products for the treatment of common diseases, reported in the past 3 years

Bioactive Compounds from Food and Its By-products: Current Applications. . . 13

Aging Antioxidant activity, problems tyrosinase inhibition, α-glucosidase and α-amylase inhibition

Alkaloids: Caulerpin and caulerpenin PC: Phenols, flavonoids, and tannins Terpenes: Sesquiterpenoids, diterpenoids, sitosterol Saponins

Sea grapes [Caulerpa racemosa (Forssk) J. Agardh]

Animal model

Assay model

Clinical trial

Flavonoids

Date palm (Phoenix dactylifera L.)

Hypoglycemic activity CD

Hypolipidemic activity Atherosclerotic Animal model

Flavonoid: Hesperidin

Animal model

Citrus (Citrus spp.) fruit

Aging skin

Antiaging activity

Olive (Olea PA: Hydroxytyrosol europaea L.) oil PC: Oleuropein

Animal model

Ultraviolet Animal model (UV)induced skin wrinkle formation

Antiaging activity

Stilbene: Resveratrol Flavonoids: Anthocyanins

Metabolic syndrome

Disorder

Grape (Vitis labrusca L. cv. Baile) peel

Bioactivities

Antioxidant, antidiabetic, and anti-inflammatory activities

Main FBC

Purple grumixama Flavonoids: Cyanidin-3-Oglucoside, delphinidin (Eugenia quercetin, and catechin brasiliensis derivatives Lam.) Tannins: Ellagic acid derivatives

(Vell.) Berg] peels

Source

Table 3 (continued) References

[21]

# blood glucose, TC; " coactivator PGC-1α (regulator of energy metabolism)

# LDL-C, "HDL-C, # TC

[100]

# TC, plaque formation, [99] adipose deposition, macrophage infiltration into atherosclerotic lesion

# NF-κβ pathway regulation [22] " Nrf2 expression

[98]

# skin thickening, " Nrf2 expression, " HO-1 expression, # MMP-1, # MMP-9

# body weight, TG, " insulin [31] sensitivity

Outcomes

14 Bianca Rodrigues de Albuquerque et al.

PD

AD

AD

Estrogenic effect

Hypolipidemic, antioxidant and anti-inflammatory

Neuroprotective action Neuroprotective action

Terpene: Zeaxanthin

Neuroprotective Pas: Chlorogenic acid action Flavonoids: Catechin, epicatechin Tannins: Procyanidins, B-type procyanidins, and epicatechin Neuroprotective action

Isoflavones

Flavonoids: Anthocyanins, flavanols, phenolic acids Tannins: Proanthocyanins

Anthocyanins and other unidentified phenolic compounds

Flavonoids: Flavones, vitexin, nobiletin, ellagic acid Terpenes: Squalene, geranyl isovalerate

Soybean [Glycine max (L.) Merrill]

Wolfberry (Lycium barbarum L.)

Avocado (Persea americana Mill.) peel

Dried grapes (raisin) (Vitis vinifera L.)

Mulberry (Morus alba Linn.)

Pomegranate (Punica granatum L.) juice and seed

PD

Clinical trial

CD

Animal model

Animal model

Animal model

Animal model

Clinical trial

Clinical trial

Animal model

CD

CD

Hypocholesterolemia activity

Flavonoids: Anthocyanins

Purple-black barberry (Berberis integerrima Bunge)

CD

Hypocholesterolemia activity

Pecan nut [Carya Flavonoids, saponin, and illinoinensis alkaloids (Wangenh.) K. Koch]

[101]

[104]

[105]

[37]

"HDL-C, #vascular age

# lipid peroxidation

"spatial memory, # malondialdehyde

(continued)

" tyrosine hydroxylase level, [107] striatal dopamine, # malondialdehyde, proinflammatory cytokines level

" memory, # amyloid burden [106]

[103]

# coronary heart disease, myocardial infarction, cardiovascular disease death

#TG, TC, LDL-C, [102] sd-LDL-C, non-HDL-C, and TC/HDL-C

#LDL-C, TC, " HDL-C

Bioactive Compounds from Food and Its By-products: Current Applications. . . 15

Colon cancer

Antiproliferative activity

Anticancer activity

Terpene: Lutein

Sulforaphane

Flavonoids: Cyanidin derivatives, peonidin derivatives, quercetin derivatives, myricetin derivatives Terpenes: Ursolic and oleanolic acids

nd

Raw broccoli (Brassica oleracea L. var. italica) sprouts

Cranberry (Vaccinium macrocarpon ait.)

Akebia trifoliata Triterpenoids (Thunb.) Koidz. Pericarp

Colon cancer

Neuroprotective action

Stilbene: Resveratrol

Anti-inflammatory activity

Neuroprotective action

Animal model

Animal model

Animal model

Animal model

Animal model

Assay model

Colon Animal model inflammation

PD

AD

AD

nd

Neuroprotective action

PAs: 3-O-caffeoyquinic, 5-Ocafferoylquinic, and 3-Oferuloylquinic acids Flavonoids: Isoorientin Saponins

Disorder

Sugarcane (Saccharum officinarum L.) top

Bioactivities

Main FBC

Source

Table 3 (continued)

[108]

[109]

[27]

[87]

[110]

" dopamine levels, acetylcholinesterase, and tyrosine hydroxylase activities # number of aberrant crypt foci; " percentage of Bifidobacterium and clostridium XVIa # Proinflammation, tumor incidence, multiplicity, and burden

" weight body and antiinflammatory activity, # NO synthesis

[79]

"spatial learning, memory, number of neurons

Protection against memory loss and tau pathology, # amyloid burden

References

Outcomes

16 Bianca Rodrigues de Albuquerque et al.

Antioxidant and anticancer activities Antioxidant activity

Antioxidant activity

PA-terpene: γ-oryzanol

Flavonoid: Anthocyanins

Flavonoids

Glucosinolate glucoraphanin, sulforaphane

Rice (Oryza spp.) bran oil

Grape (Vines vinifera L var. Merlot) residues

Brazilian pepper (Schinus terebinthifolius Raddi)

Broccoli (Brassica oleracea L. var. italica)

Antimicrobial activity

Colon cancer

Anticancer activity

PAs: Benzoic and cinnamic acids Flavonoids: Anthocyanins, flavan-3-ols Tannins: Proanthocyanidins

Sorghum [Sorghum bicolor (L.) Moench] bran

Ileostomy patients

Nd

Nd

Skin cancer induced by UV

Lung cancer

Anticancer activity

Terpene: Limonene

Mandarin (Citrus reticulata Blanco cv Dancy) peels

Colon cancer

Anticancer activity

PAs: Hydroxycinnamic acids, benzoic acids Flavonoids: Flavonols and flavan-3-ols Terpenes: Triterpenic acids

Jujube (Ziziphus jujuba mill.)

Lung metastasis in colorectal tumor

Anticancer and antimetastatic activities

Flavonoid: Tricin

Grains bran

Clinical trial

Eukaryotic model organism— Saccharomyces cerevisiae

Animal model— Caenorhabditis elegans

Animal model

Animal model

Animal model

Animal model

Animal model

[111]

[114]

[115]

[116]

#skin lesions

" longevity

"protection against oxidant agent (H2O2)

(continued)

#Escherichia coli K12 growth [117]

[113]

Inhibition of proliferation and induced apoptosis in cancer cell

#tumor growth, " apoptosis [112] of malignant cells

#disease activity index, [26] spleen weight, tumor numbers; " colon length, hematological parameters

# tumor growth and lung metastasis

Bioactive Compounds from Food and Its By-products: Current Applications. . . 17

Ulcerative colitis

nd

SARS-CoV-2

Disorder

Animal model

Clinical trial

Animal model

Assay model

References

[119]

# lesions, oxidative stress, [120] and inflammatory process

" antioxidant activity in the plasma

# viral load, lung injury, and [118] pulmonary pathology

Outcomes

nd not determined, PA phenolic acid, PCs phenolic compounds, CDs cardiovascular diseases, AD Alzheimer’s disease, PD Parkinson’s disease, TC total cholesterol, TG triglycerides, HDL-C high-density lipoprotein cholesterol, LDL-C low-density lipoprotein cholesterol, NO nitrogen oxide

Anti-inflammatory activity

Mulberry (Morus Phenolic acids, flavonoids, macroura Miq.) anthocyanins

Antiviral activity Antioxidant

Sulforaphane

Cruciferous vegetables

Bioactivities

Black rice (Oryza Flavonoids, anthocyanins sativa L. indica) Verene and Artemide varieties

Main FBC

Source

Table 3 (continued)

18 Bianca Rodrigues de Albuquerque et al.

Bioactive Compounds from Food and Its By-products: Current Applications. . .

19

2.3 Alkaloids and Xanthine Alkaloids

Alkaloids are nitrogenous compounds present in plants and some fungi. They can be classified as true alkaloids, which are those derived from amino acids and with at least one nitrogen atom in the heterocyclic ring; protoalkaloids, which do not contain nitrogen in the heterocyclic rings; and pseudoalkaloids that are terpene or steroid derivatives. In addition, they can be subdivided according to their structure into pyrrole, pyrrolidine, pyrrolizidine, pyridine, piperidine, tropane, quinoline, isoquinoline, aporphine, norlupinane, indole, indolizidine, terpenoid alkaloid, imidazole, purine, and steroid [11]. Xanthine alkaloids are compound derivatives of purine alkaloids. The most studied xanthines are caffeine and theobromine, both derivatives from purine alkaloids that show in their chemical structure nitrogen atoms, which contribute to their bioactivity, such as anti-asthmatic and antibacterial activities, besides neurological protection [12].

2.4

Saponins

Saponin is a class of secondary metabolites found in more than 500 plant species; however, the main source of this compound class is the bark of the Quillaja saponaria Molina tree. Saponins can present bioactivities, such as anti-inflammatory, antiviral, and anticancer activities, among others. Furthermore, these compounds have been exploited as natural surfactants and stabilizers of emulsion systems. This physicochemical property is related to the molecular structure of saponins, which consists of a hydrophobic triterpenoid or steroid (aglycone) skeleton and one or more hydrophilic oligosaccharides. In this way, the classification of saponins is performed according to the aglycone part of the molecular structure and according to the number of similar sugar chains [13].

2.5

Glucosinolates

Glucosinolates are compounds of the secondary metabolism of plants that present in their structure sulfur molecules. More than 130 glucosinolates have been identified in nature, most of them found in plants belonging to the Brassicaceae family. In plants, these compounds are responsible for the unique aroma and flavor and exert defense mechanisms against microbes and herbivores. The dietary ingestion of glucosinolates may provide beneficial outcomes, such as anti-inflammatory, antimicrobial, antioxidant, and cholinesterase inhibitor activities [14]. When plant tissue is damaged, glucosinolates undergo the action of an enzyme, namely, a β-thioglucosidase (myrosinase), which converts them into nitriles, thiocyanates, and isothiocyanates. In the human and animal body, these compounds are converted into sulforaphane, a chemical capable of exerting antimicrobial, antioxidant, anti-inflammatory, and antiproliferative activities, the reason why a diet rich in vegetable crucifers (major glucosinolate sources) is indicated for health maintenance [15].

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Bianca Rodrigues de Albuquerque et al.

Bioactivities of Natural Compounds from Food and Food By-products Foods and food by-products can be a source of diverse chemical compounds that can exert bioactivities in a system (such as a food matrix) and/or body, such as shown in Tables 2 and 3. Next, the main bioactivities carried out by FBC will be addressed.

3.1 Antioxidant Activity

Oxidative processes can be initiated from free radicals present in a system or body, these being known as reactive oxygen substances (ROS), such as oxygen derivatives, namely, peroxyl (•OOH), hydroxyl (•OH), hydrogen peroxide (H2O2), and superoxide anion (O2-). Oxidative processes in a system, such as a food matrix, can lead to the degradation of macromolecules, occasionally leading to the degradation of the product and making it unsuitable for consumption. In the human body, the oxidative process can also attack macromolecules, such as lipids and protein, causing cellular and DNA damage. These harmful actions to health can lead to the development of several chronic diseases, such as diabetes, cardiovascular diseases, and cancer, among others, as well as the triggering of neurodegenerative diseases [6, 7]. BC found in foods and their by-products, such as those shown in Table 2, may be able to scavenge oxidant species, preventing and delaying the oxidative process. In light of this, several natural compounds obtained from food have been studied as alternatives for the treatment of diseases. The following will present some effects of FBC on diseases caused by oxidant and chronic inflammatory processes.

3.2 Antiinflammatory Activity

Inflammatory processes where there is an increase in the production of antibodies are fundamental for the fight against infections and the recovery of the organism, leading to the reestablishment of homeostasis. Notwithstanding, the prolongation of the inflammation process can lead to the installation of a low—however chronic—proinflammatory state. This state is correlated with the emergence of chronic ailments such as diabetes, cancer, cardiovascular, and neurological diseases. To minimize inflammatory processes, several drugs can be used: (1) glucocorticoids, that is, those that act on the inhibition of prostaglandins and proteins involved in inflammatory processes, and (2) nonsteroid drugs, which are those that act through the enzyme cyclooxygenase. However, these drugs can have several significant side effects for systems and organs. In order to develop new drugs with less adverse effects, natural compounds, such as polyphenols, terpenes, alkaloids, and saponins, have been studied concerning their ability to mitigate inflammatory processes [16]. Table 2 presents recent reports on promissory antiinflammatory activity of food and food by-product extracts.

Bioactive Compounds from Food and Its By-products: Current Applications. . .

21

3.3 Antiaging Activity

Aging is an unavoidable event affected by genetic, environmental factors, and lifestyle. Indirect evidence suggest that manifold BC can display antiaging effects, albeit direct evidence in this field is still needed [17]. The aging process also occurs in the skin and can be characterized by a reduction in the production of important biochemical elements, such as collagen, elastin, and hyaluronic acid, which makes the skin sensitive, damaged, wrinkled, and sagging. Normally, skin aging occurs naturally; however, cellular aging can be accelerated by environmental conditions, such as exposure to an ultraviolet radiation process (photoaging), pollution, and toxic agents (extrinsic factors). Some daily habits can slow down the process of these skin degradation processes, such as stress management and a balanced diet that includes BC, especially those with an antioxidant character. In addition, some BC can benefit collagen synthesis via the activation of TGF-β/Smad [18, 19]. The antiaging activity of FBC is closely linked to the two aforementioned bioactivities, i.e., antioxidant and antiinflammatory activities, but also to the specific interaction of such compounds with aging-related enzymes, acting as epigenetic modulators [20]. As the aging process is the main risk factor for the development of chronic diseases, strategies to minimize the speed of cellular aging can delay or eliminate the progress of diseases, and, in this sense, the balance of ROS is linked to greater longevity [21]. In addition to the substantially investigated resveratrol, epicatechin, quercetin, and curcumin, other FBC have been reported to act as antiaging agents (Tables 2 and 3). For example, the synergistic effect of the antioxidant and anti-inflammatory activities of olive oil promoted an antiaging action on mouse dermal fibroblasts treated with epinephrine [22].

3.4 Antiproliferative Activity

The mechanism of action of BC on malignant cells is not fully unraveled. Nonetheless, it is known that their antioxidant and anti-inflammatory activities have a relation with the antiproliferative effects verified for food compounds. Extracts obtained from fruit by-products, namely, kiwi, jabuticaba, and camu camu peels have shown antiproliferative activity toward tumor cells [23–25] (Tables 2 and 3). Moreover, terpene-rich extracts of mandarin peel were able to reduce tumors caused by lung cancer [1]. Likewise, the dietary consumption of jujube fruit, a great source of phenolic acids, flavonoids, and terpenes, has decreased colon cancer activity and decreased the number of tumors in mice [26]. Finally, the daily intake of food sources of sulforaphane has been correlated with the reduction of cancer [27].

3.5 Antimicrobial Activity

Bacterial resistance to medicinal drugs has been a momentous issue over the past years. In this context, it is imperative to detect novel and efficient alternatives. Plant matrices are potential sources of powerful bactericidal agents that are already being incorporated in novel commercial drugs with efficient bactericidal effects and, not

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less important, with reduced side effects and hepatotoxicity [28]. Phenolic compounds have shown antibacterial activity against various bacteria and fungi [26]; for example, the phenolic extract of kiwi peel, composed of quercetin derivatives, epicatechin, and B-type (epi)catechin, showed antibacterial and bactericidal activity against Staphylococcus aureus, Bacillus cereus, Listeria monocytogenes, Escherichia coli, Salmonella typhimurium, and Enterobacter cloacae at low concentrations, besides antifungal and fungicidal effects against Aspergillus ochraceus, A. niger, A. versicolor, and Penicillium aurantiogriseum [25]. Other evidence on the antimicrobial potential of FBC is displayed in Table 2.

4

Applications of FBC

4.1 Pharmaceutical Products

FBCs are of great importance for the development of nutraceuticals and new drugs. In the last decades, the use of phytochemicals has been increasing to promote good health and the prevention of various diseases [6, 7, 29]. In the following topics, applications of FBC for the treatment of the main pathologies currently harming world population will be addressed.

4.1.1 Metabolic Syndrome and Diabetes

Metabolic syndrome (MS) is a pathological condition that occurs by oxidative stress and chronic low-grade inflammation in the body, characterized by at least four pathological diagnoses, namely, abdominal obesity, decreased ability to process glucose, dyslipidemia, and high blood pressure. MS patients should follow a diet rich in fruits and vegetables to improve gut conditions. Several studies have indicated that FBC may attenuate MS. The ingestion of catechin and caffeine, for instance, can facilitate weight loss, act in the activation of enzymes, and improve gut microbiota [30, 31]. Table 3 shows recent evidence on the positive effects of FBC in MS patients. Type 2 diabetes (T2DM) is one of the most severe and common chronic ailments worldwide. The global diabetic population is expected to exceed 400 million in 2030. In T2DM, high levels of glucose induce the production of reactive oxygen species, also causing metabolic stress, initiating the inflammatory process, and decreasing the antioxidant response of the immune defense system [32]. Sundry FBCs have displayed potential as antidiabetic agents, for example, the intake of flavonoid-rich foods, especially flavanols, flavanols, and anthocyanins, was associated with a lower risk of developing T2DM [33]. In a study applying an in vivo model, diabetic rats daily fed a bread made with onion peel extract (1–3%) or onion powder (7%) had a significant decrease in blood glucose levels, besides improvements in body weight and oxidant activity after 8 weeks of experiment [32].

Bioactive Compounds from Food and Its By-products: Current Applications. . .

23

4.1.2 Cardiovascular Diseases

Cardiovascular diseases (CVDs) are the main cause of premature deaths worldwide. The risks of developing CVD may be related to hereditary factors and metabolic syndrome. However, an unhealthy lifestyle, including low frequency or lack of physical exercise, high stress levels, and poor eating habits, can trigger inflammatory and oxidative processes that are important factors in the emergence and evolution of CVDs. A balanced diet, including fruits and vegetables rich in antioxidant compounds, is recommended to reduce the cardiovascular risk. In this sense, FBC have been evaluated as therapeutic agents in the prevention and treatment of heart diseases [8]. Pharmaceutical products based on plant terpenes have been patented in the last 10 years [8], due to their role in CVD prevention. The daily ingestion of other FBC can also have a positive effect on cardiovascular health. Aali et al. [34], in a study conducted with 404 women, verified that the ingestion of lignin and stilbenes had an effect on body mass index, whereas waist circumference decreased with the intake of beverages containing phenolic acids. Moreover, stilbene intake impacted total cholesterol, likewise other polyphenol ingestion influenced triglyceride levels. Other in vivo studies and clinical trials are shown in Table 3.

4.1.3 Neurological Diseases

The rate of neurological diseases has been increasing worldwide, being a global health problem that mainly affects the elderly population (> 60 years old). It is estimated that by 2050 more than 80 million people will be diagnosed with some neurological disease [35]. Acute neurological diseases are caused by a punctual condition causing permanent damage, whereas chronic neurological diseases are caused by hereditary factors or can be developed due to the frequent exposure to heavy metals, toxic substances, and high level of stress, among other conditions that modify the normal functioning of proteins. This class of chronic diseases that tends to be aggravated over the life are denominated as neurodegenerative [7, 35, 36]. Among the neurodegenerative diseases, Alzheimer’s disease (AD), Parkinson’s disease (PD), and amyotrophic lateral sclerosis are the most common in the elderly population [35, 37]. A balanced diet rich in FBC has been recommended for the prevention/ delay of neurodegenerative diseases. Some FBCs exert neuroprotective activity, acting to inhibit and stop oxidant and inflammatory processes at a neuronal level [35, 36]. Some promising studies that address the use of FBC in the treatment of neurodegenerative diseases are presented in Table 3. Gol et al. found that a daily dose of 6 g of dried grapes (raisin) over a period of 60 days increased the antioxidant activity in the blood of AD-induced rats, consequently decreasing the level of malondialdehyde [37]. Another in vivo study showed that the polyphenol-rich grape seed extract had interaction and disintegration capacity of

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paired helical filaments caused by the abnormal unfolding of tau proteins, pathological markers of AD [38]. Many authors have investigated the therapeutic potential of anthocyanins on neurological diseases. This class of flavonoids has shown the ability to improve cognitive functions and memory by protecting neurons and inhibiting the deposition of amyloid beta peptides, also acting as antioxidant and anti-inflammatory agents [35]. Zhang et al. verified that the daily intake of foods rich in flavonoids, such as tea, apples, berries, oranges, and red wine, reduced the risk of death in patients diagnosed with PD [36]. 4.1.4

Cancer

4.2 Cosmetic Products

Cancer is a global health problem that affects more than 18 million people a year, and unfortunately, trends indicate that this figure could rise beyond 23 million new cases of by 2030. Traditional cancer treatments include surgery and radiation treatment, followed by chemotherapy treatment with antimetabolic substances and DNA-interactive agents. Due to the complexity of each cancer case, sometimes the combination of these treatments cannot achieve the cure, and at times, they cause severe side effects, affecting patients’ life quality [39]. FBCs are promising agents for the prevention and treatment of malignant tumors (Tables 2 and 3). The bioactive extracts obtained from camu camu and kiwi peels, for instance, were able to inhibit the proliferation of cervical carcinoma cells (HeLa), lung carcinoma (NCI-H460), breast carcinoma (MCF-7), and hepatocellular carcinoma (HepG2) [24, 25]. Also, in vivo studies (Table 3) have shown promising results regarding alternative treatments based on FBC. For instance, the regular intake of broccoli, a food rich in sulforaphane, can contribute to the intestinal flora balance, thus favoring the growth of bacteria that offer protection against colon carcinoma [27]. Traditionally, cosmetic products are formulated with chemical substances of synthetic origin. However, some of these compounds have been banned due to the uncertainty regarding their safety when associated to prolonged exposure. At the same time, consumers’ preference has been shifting toward products with “natural” claims [40]. As a result, the cosmetic industry has been looking for alternatives for the development of products based on natural ingredients, such as FBC [40, 41]. Next, a few examples of the application of food-derived ingredients into cosmetic formulations are presented. Coffee silverskin has been applied as a promising anti-wrinkle and antiaging agent due to the presence of chlorogenic acid and caffeine in its composition, compounds that can exert UVB absorption properties [42]. Natural colorants obtained from foods, such as anthocyanins, quercetin, curcumin, and carotenoids, are alternatives to the synthetic dyes commonly applied in cosmetics, which are often associated with allergies [43]. Likewise, essential oils rich in terpenes have been used to produce fragrances. Lemon,

Bioactive Compounds from Food and Its By-products: Current Applications. . .

25

tangerine, bergamot, orange, lime, and grapefruit peels have complex terpene compositions that impart unique citrus and floral aromas [44]. Vegetable oils, such as palm, olive, soy, sunflower, and safflower, can act as emulsifiers in cosmetics. Finally, saponins from food and food waste can be exploited as biosurfactants [40]. Another important factor concerning the application of plantbased cosmetic ingredients is related to the increasing consumer’s tendency to seek products classified as vegan. In this sense, a new challenge arises for the cosmetic industry to replace ingredients derived from animals [40]. 4.3 Food Products and Food Packaging

Processed food products can contain additives to ensure quality, both sensory and microbiological, throughout their shelf life. Food additives can help to maintain or improve food freshness, taste, texture, or appearance. Some food additives have been in use for centuries for preservation—such as salt (in meats such as bacon or dried fish), sugar (in marmalade), or sulfur dioxide (in wine). However, many of the additives commonly used by the food industry are of synthetic and artificial origin, and several of them have been banned due to uncertainties about their safety [3]. Furthermore, consumers, who are aware and in search of healthier habits, are looking for products free of synthetic/artificial ingredients. In this sense, the food industry faces the challenge of replacing the traditional additives with plant-based ones [3, 45]. FBCs are promising natural food additives, as presented in Table 4. Anthocyanins, flavonoids that present a range of attractive colors between pink-red-purple-blue, carotenoids with a yellow-orange-red color, and other compounds such as curcumin (golden-yellow color), are promising natural colorants for the replacement of synthetic dyes [45]. For instance, an anthocyanin-rich extract obtained from jabuticaba peel was applied to macarons. The product showed attractive and stable color throughout shelf life [46]. Likewise, anthocyanins extracted from sohiong remained stable throughout the industrial manufacturing of yoghurt, hard candy, and syrup [47]. Tomato powder rich in lycopene also showed coloring property when added to yoghurt, also improving the antioxidant capacity of the dairy product [48]. Furthermore, due to their antioxidant, antibacterial, and antifungal properties, FBC can act as preservative agents in food products. This is the case of a beetroot extract, which showed multifunctionality when applied in cooked pork processing, inhibiting lipid oxidation and the growth of Listeria monocytogenes during storage [49]. The terpene-rich grapefruit peel extract showed the ability to increase the antioxidant activity of mustard oil and consequently increase the shelf life of the product [50]. FBCs have also been used as natural additives in packing systems, such as active-intelligent food packing. Sundry studies have focused on the development of smart packaging that indicates the

Anthocyanins

Anthocyanins

Anthocyanins

Anthocyanins

Carotenoids

Lycopene

Epicatechin, catechin, procyanidins Functional B1 and B2, protocatechuic, ingredient p-OH-benzoic, gallic acids

Phenolic compounds

Phenolic compounds

Sohiong (Prunus nepalensis L.)

Red and purple potatoes (Solanum tuberosum L.)

Fig (Ficus carica L.) peels

Blackthorn (Prunus spinosa L.) epicarp

Mandarin (Citrus reticulata L.) peels

Tomato (Lycopersicon esculentum L.) peel powder

Cacao (Theobroma cacao L.) hulls

Cinnamon [Cinnamomum cassia (L.) J.Presl] leaf residue

Grape (Vitis vinifera L.) seed powder

Functional ingredient

Functional ingredient

Natural colorant/ functional ingredient

Natural colorant

Natural colorant Natural colorant

Natural colorant

Natural colorant

Natural colorant

Anthocyanins

Jabuticaba [Myrciaria Jaboticaba (Vell.) Berg] peels

Purple color and antioxidant activity

Stable color and sensorial acceptance

Stable color and high sensorial acceptance

Attractive and stable color

Outcomes

Balady bread

Yogurt

Biscuits

Yoghurt

Antioxidant activity

Antioxidant activity and albumin denaturation inhibition

Antioxidant activity

Intense red color and antioxidant activity

Bake and bread Development of yellow color

Icing and Beijinho

Soft drink

Yoghurt, syrup, and hard candy

Macarons

Technological propriety Product

FBC

Source

Food products

Table 4 FBC as ingredients in food products and packaging, reported in the last 3 years

[57]

[85]

[56]

[48]

[122]

[121]

[92]

[47]

[46]

References

26 Bianca Rodrigues de Albuquerque et al.

Phenolic compounds and glucosinolates

Flavonoids and triterpenes

Phenolic compounds

Naringin, trimethoxybenzoic acid, ferulic acid, epicatechin, caffeic acid, vanillic acid, and chlorogenic acid

Broccoli sprout (Brassica oleracea L. var. italica)

Red propolis

Beetroot (Beta vulgaris L.)

Grapefruit (Citrus paradisi L. cv. Redblush) peel

nd not declared

Preservative

Proanthocyanidins

nd

CO2 indicator

pH indicator

Anthocyanins

Red cabbage (Brassica oleracea L.)

pH indicator

Natural preservative

Natural preservative

Natural preservative

Natural preservative

Functional food

Turmeric root (Curcuma longa L.) Curcumin Black carrot (Dacus carota L.) Anthocyanins

Anthocyanins

Red barberry (Berberis vulgaris L.)

Food packaging

Anthocyanins and quercetin derivatives

Purple potato (Solanum tuberosum L. var. bora valley) and Tartary buckwheat (Fagopyrum tataricum L.) seed

Increase of antioxidant activity and shelf life extension

Antibacterial activity against listeria monocytogenes and inhibition of lipid oxidation

Inhibition of lipid oxidation and putrefaction

[50]

[49]

[125]

[124]

[123]

Edible bioactive packing

Improvement of the antioxidant and antibacterial activities. The film acts as an oxygen barrier

Smart packing Color change according to pH increasing

[55]

[53]

[52] Smart packing Color change from blue to purple with the increasing of CO2 concentration

Biodegradable Color change in response to fish [51] smart film fillet freshness

Mustard oil

Cooked pork

Tilapia salami

Ricotta cheese Antibacterial activity against listeria sp.

Extruded food Antioxidant activity product

Bioactive Compounds from Food and Its By-products: Current Applications. . . 27

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Bianca Rodrigues de Albuquerque et al.

freshness of food through the incorporation of anthocyanins and curcumin in polymers [51–53]. Essential oils rich in aldehydes, phenolics, and oxygenated terpenoids are excellent natural compounds to integrate food packaging, mainly because of their hydrophobicity and strong antimicrobial properties [54]. Furthermore, the antioxidant and antimicrobial actions of the FBC added to the film may help in food preservation [55]. Other examples of FBCs incorporated in food packaging are shown in Table 4. Another interesting and promising aspect of the application of FBC is the development of functional ingredients to produce functional food products (see Table 4). In this sense, the food by-products and residues have been investigated as potential FBC. For example, encapsulated extracts of cocoa hulls were used to confer antioxidant activity to traditional cookies [56], as well as grape seed powder added to bread [57]. Saponins obtained from edible oil wastes, namely, soy and camellia oil cake, were used to remove pesticides from fruits and vegetables. The results indicated that saponins can be a more efficient biodegradable reseller alternative than conventional detergents [58].

5 Green Approaches for the Recovery of BC from Food Industry By-products and Residues Agri-food processing produces massive amounts of nonedible residues and by-products, which are effortlessly accessible elements for upcycling active molecules. In this sense, polyphenols are especially valued given their opulence in plant-based residues and the market interest of their biological capabilities (e.g., natural antioxidant property) as part of drug, nutraceutical, and cosmetical formulations [59]. The extraction approaches for the recovery of FBC are in permanent evolution. For instance, the adoption of alternative solvents like ionic liquids and deep eutectic solvents (DESs) has been proven to minimize the environmental impact. Moreover, the integration of green techniques, as the combination of ultrasound with SFE processes, can be a viable means to increase the extraction kinetics, boot the extraction yield, and reduce the extraction time (and thus operational costs), while also preserving the final product’s quality [60]. Table 5 shows other combinations of extraction processes or the employment of various technologies in parallel aiming the recovery of interest BC from food residues. Another approach that could increment extraction efficiency is the combination of an enzymatic treatment at high pressure with the supercritical fluid extraction (SFE) or with pressurized liquid extraction (PLE) [61, 62]. Likewise, pulsed electric field (PEF), a nonthermal technology usually used for food conservation,

Functional foods with upgraded Chlorogenic, protocatechuic, enhanced nutritional and caffeic, ferulic, and coumaric acids, caffeine, rutin, quercetin, health-promoting effects. Natural ingredients such as kaempferol, and quercetin preservatives, antioxidants, dyes, natural flavors, and prebiotics for the food and pharmaceutical industries

Coffee (Coffea spp.) skin, pulp, parchment, silverskin

Grape (Vitis vinifera L.) pomace Tannins, anthocyanins, resveratrol, gallic and syringic (skins, pulp, seeds, stems), acids, hesperidin, p-coumaric grape marc (skins and pulp), and winery lees and dihydroxybenzoic acids, lignans, stilbenes

Polyphenols with widely proven beneficial effects on human health

Cocoa (Theobroma cacao L.) bean shells (CBS) and pod husks (CPH)

(continued)

MAE; ohmic heating (OH); high- [126, 129] Development of novel and voltage electrical discharges functional foods. Fortification (HVED); optimized solidand enhancement in meat liquid extraction (SLE); products, protection in olive pressurized hot water extraction and vegetable oils. Applications (PHWE); SFE SC-CO2; and to animal feed, nutraceuticals, dietary supplements, and integrated pressure-driven cosmetics membrane-based technologies

[72, 128] Ultrasound (UEA), pulsed electric field (PEF), pressurized liquid (PLE) subcritical and supercritical fluid (SWE and SFE) technologies. Fermentation using Penicillium purpurogenum and Saccharomyces cerevisiae

Obtainment of antioxidants and Ultrasound (UEA), microwave [127] nutraceuticals. CPH extracts as (MAE), pulsed electric field ingredients of functional (PEF), pressurized liquid (PLE) cosmetic products subcritical and supercritical fluid (SFE and SWE) technologies

Aqueous ethanol extraction or by [126] Production of micro- and alkaline treatment nanomaterials, particles, and fibers such as nanospheres for the protection of BC, drug delivery system, Pickering emulsion stabilization, and even controlled release fertilizer

Prolamins, which are the endosperm storage proteins of cereal grains

References

Cereal by-products such as gluten meals (GM), dried distillers grains with solubles (DDGS) and brewer’s spent grain (BSG)

Recovery approach

Targeted upcycling/application

Target bioactive compound(s)

Food by-product/residue

Table 5 Current approaches for the recovery of FBC from important plant-based by-products and residues, reported in the past 3 years

Bioactive Compounds from Food and Its By-products: Current Applications. . . 29

Target bioactive compound(s)

Recovery approach

References

Functional food additives, Valuable BC such as tyrosol, preserving agents, besides hydroxytyrosol, oleocanthal, cosmeceutical products oleuropein, ligstroside, squalene, fatty acids, quercetin, triterpene acid, and carotenoids

[135] The shell as a biomass for ethanol A biorefinery approach using Polyphenols as stilbene transintegrated extraction processes production and source of resveratrol, flavonoids as by PLE, SFE, and SWE. The BC. Skins extracted with this proanthocyanidins, isoflavones solvent (ethanol) produced ethanol for the recovery of daidzein, and genistein, besides from peanut shells would be the natural antioxidants for food luteolin and carotene main extractor industry. Peanut meal can furnish protein and polysaccharides to enrich foodstuff

Olive (Olea europaea L) oil pomace, skin, and waste waters

Peanut (Arachis hypogaea L.) skin, shell and peanut meal

Enzymatic method, UEA, MAE, PLE, SFE, and HVED; infrared-assisted extraction (IR-AE) and membrane separation techniques

SFE, MAE, UAE, and more recently, ultrasound-assisted natural deep eutectic solvent (NADES)

Antioxidant compounds such as Packaging, fat replacer, and carotenoids, besides gallic acid, antioxidant and antimicrobial mangiferin, ellagic acid, applications are the most kaempferol, quercetin explored

Mango (Mangifera indica L.) peel, bagasse, and kernel

[61, 62]

[133, 134]

[132]

Natural additives such as PLE and UAE both employing preservatives, antioxidants, and ethanol and water (3:1, wt.) as nutraceuticals the solvent

Antioxidant polyphenols, hesperidin, narirutin

[131]

Lime (Citrus latifolia Tan.) pomace

Hydrothermal pretreatment optimized for each target compound class

Natural food ingredients such as preservatives, antioxidants, colorants, and nutraceuticals

[130] Obtainment of anthocyanin-based Optimized high-intensity ultrasound (HIUS) extraction: nutraceuticals for the Ultrasound intensity of 3.7 W/ prevention and treatment of cm2 and solvent composition chronical ailments. Pectin-rich biomasses can be used as (water/ethanol) of 50 g/100 g thickener and gelling agents

Targeted upcycling/application

Jabuticaba [Myrciaria Jaboticaba Sugars, organic acids, and (Vell.) Berg] peel antioxidant anthocyanins and phenolic compounds

Jabuticaba [Myrciaria Jaboticaba Anthocyanin-rich extracts and (Vell.) Berg] peel pectin-rich biomass

Food by-product/residue

Table 5 (continued)

30 Bianca Rodrigues de Albuquerque et al.

Obtainment of natural antioxidants and repellents ingredients

[138]

Bioactive phenolics such as rosmarinic acid

Sweet basil (Ocimum basilicum L.) hydrodistilled residue-byproducts

Enzymatic pretreatment

Optimized UAE (maximum yield [137] of 11.15%): Extraction time of 30 min, temperature of 33 °C and power of 400 W

High-quality pectin with great functional and technological features for pharmaceutical or food applications

High-purity pectin

Sunflower (Helianthus annuus L.) head and stem

[136]

UAE; MAE; SFE; enzymeassisted SFE; ultrasound and enzyme (cellulase)-assisted extraction; cellulase magnetic nanobiocatalyst; cloud pointassisted bioactive extraction; surfactant-assisted extraction; solid-state culture

Pomegranate (Punica granatum Conjugated fatty acids like Ingredients to improve food L.) seed and peel linolenic acid, phenolics such as properties and shelf life, gallic acid, and tocopherols preservatives and natural additives in food packing, pharmaceutical applications

Bioactive Compounds from Food and Its By-products: Current Applications. . . 31

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emerges as an optimal alternative for pretreatment before extraction by virtue of its minimal operational time and efficient power consumption, although still in its infancy in the food and nutraceutical industries [63]. Membrane technology is a feasible possibility for BC recovery, as the obtainment of valuable substances from food residues can compensate the cost of compulsory waste treatment procedure. In this regard, membrane separation can be employed in food waste treatment in combination with recovery of FBC, which will drive to a sustainable manufacture of nutraceuticals [64]. Here it is worth commenting that, in spite of all technicalscientific progress archived in past few years, the most appropriate BC extracting approach continues depending on the food matrix, targeted BC, and their structural features and interactions [59].

6 Current and Future Perspectives Regarding the Recovery and Application of BC from Food and Agro-industrial Food Waste The food and pharmaceutical industries have been investing on the research and development of new products containing BC with the aim to replace synthetic ingredients and additives. As a result of this tendency, many patents have been published on the topic in the last years. The following are a few examples of patents that describe and protect the intellectual property of methods for the extraction and recovery of BC, as well as some potential applications. One of the biggest and mutual efforts from the scientific and manufacturing communities has been the utilization of food residues and bio-products as suitable sources of BC. In a patent from 2016, apple peels were applied as a source of flavonoid-rich fractions for the development of cosmetic and nutraceutical formulations, aiming at the prevention and treatment of conditions associated with oxidative stress and/or inflammation, including neurodegenerative diseases [65]. Potato peel waste was also used as a source of phenolic compounds in a patent published in 2017. The patent describes a method for extracting antioxidative phenolic acids that can be applicable to industries such as foodstuff ingredients and cosmetics. Another major interest of the food industry has been the replacement of synthetic color-ants with natural alternatives. In particular, anthocyanin-based colorants have been on the radar with many products already available on the market containing these natural sources of red and purple dyes. In 2017, a patent from Kraft Foods in partnership with the University of Illinois was published describing a method of extracting anthocyanins from colored corn cultivars, producing aqueous-based colorants suitable for application in food products [66]. Similarly, and also in 2017,

Bioactive Compounds from Food and Its By-products: Current Applications. . .

33

another patent describing a method for extracting anthocyanins was published, this time using sorghum as the food matrix. The extracts were also aqueous-based aiming to be applied in a variety of edible products such as foods, nutraceuticals, and pharmaceutical formulations [67]. Mars Food also published a patent on this topic, describing a method for isolating blue anthocyanin fractions from vegetables, fruit juices, and extracts [68].

7

Concluding Remarks Consumers’ environmental awareness is expanding, and the goals of the 2030 Agenda for Sustainable Development comprise the efficient utilization of natural resources and the environmentally friendly management of chemicals and all wastes throughout their life cycle. A huge body of evidence herein discussed shows that agro-industrial food by-product and residues are sustainable sources to obtain high-value BC for food, pharmaceutical, and medical industries, within the concepts of upcycling and circular economy. However, to make food by-products ad residues viable sustainable source of BCs, the selection and optimization of satisfactory eco-friendly extraction approaches that allow the recovery and sustainability of target molecules are imperative. Thus, sustainability and circularity of processes should be major common goals for scientists and industrial sector, reducing postharvest losses, valorizing by-products, and recovering BC. Science has the huge challenge to build up scalable technology for industry to be able to be fast, with great recovery efficiency, short process time, accessibility, and minimum use of organic solvents.

Acknowledgments The authors are grateful to the Foundation for Science and Technology (FCT, Portugal) for financial support through national funds FCT/MCTES (PIDDAC) to CIMO (UIDB/00690/2020 and UIDP/00690/2020) and SusTEC (LA/P/0007/2020). L. Barros thanks the national funding by FCT, PI, through the institutional scientific employment program-contract for her contract. B.R. Albuquerque thanks FCT for her PhD grants (BD/136370/2018). R.C.G. Correˆa is a research grant recipient of Cesumar Institute of Science, Technology and Innovation (ICETI).

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Chapter 2 Food Waste Management Method Through 3R Concept Anna Ilina´, R. Ramos-Gonza´lez, R. Arredondo-Valde´s, C. Barrera-Martı´nez, E. Laredo-Alcala´, Patricia M. Albarracin, G. M. Alvarez, and J. L. Martı´nez-Herna´ndez Abstract Food waste and other organic waste that go to landfills generate greenhouse gas emissions and other types of pollution. It is very important to manage waste treatments to reduce pollution, pests, and environmental impact. The 3R concept comprises three beneficial and attractive actions for the food service industries. Reduce, a better way to reduce waste is not to generate it. Reduction is the first step in managing food waste. It is the most powerful and effective action that companies can take to manage waste. This involves designing technologies and policies to prevent or minimize waste, as the opportunity to save food and labor costs while achieving a positive impact on the environment. By avoiding the generation of waste, money is not spent on raw materials that would otherwise go to waste and, at the same time, on the costs of handling or processing these materials. Reuse, do not leave waste when using what has been achieved. Reusing food waste makes it possible to give value to something that would otherwise go to waste. In food service, the most common reuse opportunities involve the redistribution of overproduced food. Recycle, when waste is generated, look for ways to recycle it instead of leaving it in the landfill. Recycling is defined as the use of inedible waste to process it into new products to avoid wasting potentially useful materials. This activity implies development of research focused on the characterization of inedible waste and development of processes to give added value to the products of their conversion. Key words Food waste, 3R concept, Reduce, Reuse, Recycle

1

Introduction The dramatic expansion of human population (from 8 to 9.7 billion in the next 30 years) [1] next to economic influences has triggered plenty anthropogenic activities that derivate in environmental pollution eventually causing negative impact in human health and life quality [2]. Industrialization, agricultural activities, and waste elimination are the leading cause of water, soil, and air contamination around the world [3]. Within all the industrialization areas, food production and services have also experienced changes. As reported by the FAO (Food and Agriculture Organization), food production

Cristobal N. Aguilar Gonzalez et al. (eds.), Food Waste Conversion, Methods and Protocols in Food Science, https://doi.org/10.1007/978-1-0716-3303-8_2, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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Fig. 1 Worldwide food loss and waste statistic data

of primary crops was 9.4 billion tons (2019), and in agreement with its statistics, there is enough food to feed most of the world population [4]. Unfortunately, more than 800 million people still suffer from hunger, malnutrition, and other food-related problems [5]. However, despite the fact that there is still a huge need for large-scale food production, the consequences of this massive manufacture are reflected in the food waste accumulation and the way in which it is disposed of, since statistically it represents an economic, social, and environmental impact (Fig. 1) [6, 7]. Food waste refers to all the matter that during its preparation or processing was not properly used and becomes a lost, for example, red meat or fish and its bones, eggshells, vegetables and fruit skin, dairy products, coffee, grains, etc. Within this classification 60% of food waste could be avoided. However, in developing countries, food losses are hardly traceable since most of them start at the beginning of the food production cycle, due to economic limitations for adequate infrastructure and protocols to follow. Compared to developed countries, food loss occurs in the final stages of the cycle since there is no effective communication in the supply chain [8]. Hence, prioritizing food loss and waste could greatly help to improve environmental and economic problems that affect us. To accomplish the aforementioned, it is indispensable to fulfill certain strategies that meet the requirements demanded by today’s world. Within these strategies stand the circular economy (CE) in which its main objective is that raw materials and products generated by them remain in the country’s economy and that the waste produced stays to a minimum. This strategy participates in all the stages of the productive life cycle, that is, from idea creation to

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waste management and storage through manufacturing and sale. The implementation of a circular economy brings with it concepts like zero waste, which joins the current sustainability rules [9]. CE is based on an important sustainable approach with high degree of success called 3R. 3R which means reduce, reuse, and recycle is a waste management hierarchy that was first introduced by Russell and Burch in 1959 in their book The Principles of Humane Experimental Technique [10]. As main concepts of this strategy, reduce refers to the decrease of waste to minimize pollution. Reuse involves the repeated use of the waste for the same purpose. Finally, the term recycle indicates that once a waste has finished its useful life, it can be used again as raw material or for a different purpose [11]. The 3R principles have been used in food waste since they bond with the sustainable development that the world society requires promoting active participation of consumers in each activity, i.e., planning cooking purchase efficiently, using leftover meals to feed your pets, or as a potting soil conditioner [12]. Along with household consumers, large companies of different applications can also work under this management scheme since food waste has been used for the development of new products such as detergents from vegetable oils, fertilizers from slaughterhouse waste, and bio-compost from skin and seed of fruits and vegetables [13]. In this sense, for the food industry, reduction is one of the most important activities that companies can apply for residue management developing protocols to prevent the excessive generation of waste and/or reduce it, which leads to great savings in resources and protection of the environment. On the other hand, food waste reuse can be considered an activity that forces the industry to give value to products that would be discarded. Some applied activities in the industries are donation and redistribution, while for food waste recycling, the procedures are to donate all the food waste to compost companies or create their own recycle services that can manage meat, oil, and grease residues [14]. Following these strategies, important goals of the 2030 Agenda for Sustainable Development adopted by all United Nations member states in 2015 could be achieved (Fig. 2) [6]. This chapter attempts to present the current situation on the implementation of the 3R strategy in food waste management focusing on the importance of reducing or eliminating this problem transforming a disadvantage such as food overproduction into an adequate and sustainable plan for this resource use.

2

The 3R Actions Are Applied to Waste Management in the Citrus Industry Citrus fruits are the most abundant crop with around 88 million tons per year, and oranges represent 80% of this production [15]. Because of this huge industrial production of oranges, many

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Fig. 2 3R strategy and circular economy for achieving Sustainable Development Goals

wastes are generated each year. After harvesting, the orange is transported to warehouses, from which it is distributed to stores and food markets and is made available to people and companies. Companies transform it into different products such as essential oils, orange wine, orange vinegar, orange nectar, oranges in syrup, osmotic dehydrated orange, candied oranges, crystallized orange, orange pulp, orange marmalade, and orange juice. The orange processing industry produces up to 34.7 billion tons per year of by-products, such as peel, pulps, wax, and others [16]. Nowadays, a particular trend is studying the reduction and revalorization of by-products, promoting the formation of products with low environmental load [17–19] with a perspective circular economy [20]. Orange agro-industrial residues are complex matrices made up of epidermis or peel, flavedo which is a sublayer, albedo which is a spongy layer and a rich source of flavanones, vascular bundles which are the fine lines found in the orange and contains a thin membrane which is known as the septum, endocarp, or pulp and seeds [21]. To avoid dumping these residues in landfills, the easiest way to use them is to consider them as fuel after drying due to their high content of cellulose and flammable organic compounds. Furthermore, due to their excellent nutritional quality, orange residues have been used in silage systems to produce livestock feed. Silage is a way of conserving this residue, improving its nutritional properties. Technologies for the transformation of waste to compost

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applied as fertilizer in the field are promoted. The mentioned procedures allow the reduction of the generation of waste as an example of the first and second R of the three. Moreover, citric waste can be a valuable source of biologically active compounds, and they are just raw material for many medical and industrial applications. Much of these bioactive compounds are secondary metabolites used by plants to attract pollinating agents and against pathogens and herbivores. They are mainly terpenoids, alkaloids, and phenolic compounds [22]. In addition, it is mainly composed of pectin, limonene, and oil [23]. The development of technologies to obtain value-added products from wastes will help to achieve a sustainable production [24]. Alternatives to the use of citrus wastes have been proposed, such as production of ethanol, phenolic compound recovery, and single cell protein production, among others [25]. Citrus fruits are rich in vitamins A, C, and E, minerals, flavonoids, coumarins, limonoids, carotenoids, pectin, and other chemical compounds. These substances decrease the risk of diseases such as cancer, diabetes, and cardiovascular problems [23], may inactivate free radicals, prevent lipid peroxidation reactions, and avoid oxidative damage of cells [26]. One of the by-products of the citrus industry is wax. In this context, Frutech International Corporation de Mexico S.A. de C.V. is a Mexican enterprise transforming citrus fruits to produce essential oils, folds, and terpenes. From the orange essential oil process, a waxy solid fraction is produced as waste which requires either disposal or further processing. This waste is obtained in the essential oil purification process after decantation when the orange essential oil is standing at a low temperature for a specific time. The process, called winterization, consists of a partial crystallization of the raw material components, followed by separating the waxy solid from the essential oil by decantation [27]. In this process, the highest melting point glycerides and long-chain esters of fatty acids are precipitated at low temperature [28]. Additionally, this waste contains bioactive compounds which can be used as biocides [29]. Commonly, this waste does not have any commercial value. Considering that 1.5% in the production of citrus oil corresponds to orange wax, it is estimated that from 1000 kg of oranges about 5.52 kg of oil are produced and 0.08 kg of wax [15]. Considering the world production of orange, 4000 tons of waxy orange residues were produced in the 2017/2018 period [30]. At the current time, the quantity of processed oranges by Frutech provides high availability of orange wax. The orange processing season is from November to March or April, depending on the fruit quality. From November of 2016 to April of 2017, 1,200,000 tons of fruits were processed, and about 1% of wax (48 tons of wax waste) was produced. Bioactive compounds have been recovered from natural sources using solid–liquid and liquid–liquid extractions [16], cold pressing, hydrodistillation, and steam distillation [15]. However,

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these methods have some disadvantages. New technologies have been used to overcome these problems. One of these alternative technologies is the magnetic solid-phase adsorption [18]. It is a nondestructive and efficient adsorption method, characterized by using a low amount of solvents, short extraction time [31], and low energy expenditure [32]. Frutech is searching for alternatives to revalorize this orange wax waste (OWW) to obtain value-added products with low environmental impact. It has been reported that the OWW is composed of triglycerides with a high melting point which can be used to formulate products such as margarine [33], and some other products could be obtained. Besides resveratrol and limonene, the solid waste contains waxy compounds, such as wax hydrocarbons, wax esters, fatty alcohols, and minor amounts of simple coumarins and furanocoumarins [29]. The bioactive compounds in orange wax and other citrus have been reported to control Cryptotermes brevis termites [29]. The insecticidal activity is related to coumarin and furanocoumarin presence, although these compounds are contained in low concentrations (99% purity. 2. Using a hot plate, keep the NaDES at 40 °C. Once the extraction has started, keep the temperature at 45–50 °C. The ultrasonic probe raises the temperature by itself. 3. Be careful with the ultrasonic probe; it should not be turned on in the air or out of the liquid as it can cause serious damage to it and the operator. 4. It is recommended to dilute (1:5 to 1:10) the extract with mobile phase solution A for better ionization and to avoid saturating the detector.

Sustainable Extraction of Flavonoids from Agricultural Biomass and Agro. . .

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Sustain Chem Eng 2(5):1063–1071. https:// doi.org/10.1021/sc500096j ´ ngeles Ferna´ndez M, 11. Espino M, de los A Gomez FJV, Silva MF (2016) Natural designer solvents for greening analytical chemistry. Trends Analyt Chem 76:126–136. https:// doi.org/10.1016/j.trac.2015.11.006 12. Choi YH, van Spronsen J, Dai Y, Verberne M, Hollmann F, Arends IWCE et al (2011) Are natural deep eutectic solvents the missing link in understanding cellular metabolism and physiology? Plant Physiol 156(4):1701–1705. https://doi.org/10.1104/pp.111.178426 13. Dai Y, Verpoorte R, Choi YH (2014) Natural deep eutectic solvents providing enhanced stability of natural colorants from safflower (Carthamus tinctorius). Food Chem 159: 116–121. https://doi.org/10.1016/j. foodchem.2014.02.155 14. Alam MA, Muhammad G, Khan MN, Mofijur M, Lv Y, Xiong W et al (2021) Choline chloride-based deep eutectic solvents as green extractants for the isolation of phenolic compounds from biomass. J Clean Prod 309: 127445. https://doi.org/10.1016/j.jclepro. 2021.127445 15. Castro VIB, Craveiro R, Silva JM, Reis RL, Paiva AC, Duarte AR (2018) Natural deep eutectic systems as alternative nontoxic cryoprotective agents. Cryobiology 83:15–26. https://doi.org/10.1016/j.cryobiol.2018. 06.010 16. Dai Y, Witkamp G-J, Verpoorte R, Choi YH (2013) Natural deep eutectic solvents as a new extraction media for phenolic metabolites in Carthamus tinctorius L. Anal Chem 85(13): 6272–6278. https://doi.org/10.1021/ ac400432p 17. Dai Y, Varypataki EM, Golovina EA, Jiskoot W, Witkamp G-J, Choi YH et al (2021) Natural deep eutectic solvents in plants and plant cells: in vitro evidence for their possible functions. In: Verpoorte R, Witkamp G-J, Choi YH (eds) Advances in botanical research, Academic Press, vol 2021, pp 159–184 18. Loow Y-L, New EK, Yang GH, Ang LY, Foo LYW, Wu TY (2017) Potential use of deep eutectic solvents to facilitate lignocellulosic biomass utilization and conversion. Cellulose 24(9):3591–3618. https://doi.org/10.1007/ s10570-017-1358-y 19. Scelsi E, Angelini A, Pastore C (2021) Deep eutectic solvents for the valorisation of lignocellulosic biomasses towards fine chemicals.

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22. Panic´ M, Andlar M, Tisˇma M, Rezic´ T, Sˇibalic´ D, Cvjetko Bubalo M et al (2021) Natural deep eutectic solvent as a unique solvent for valorisation of orange peel waste by the integrated biorefinery approach. Waste Manag 120:340–350. https://doi.org/10.1016/j. wasman.2020.11.052 23. Vasyliev G, Lyudmyla K, Hladun K, Skiba M, Vorobyova V (2022) Valorization of tomato pomace: extraction of value-added components by deep eutectic solvents and their application in the formulation of cosmetic emulsions. Biomass Convers Biorefin. https:// doi.org/10.1007/s13399-022-02337-z

Chapter 5 Protocol for the Extraction of Lignin from Brewer’s Spent Grain Using Deep Eutectic Solvents Ana C. Cassoni, Patrı´cia Costa, and Manuela Pintado Abstract The agro-food industry annually produces tons of lignocellulosic waste with an intrinsic high value. Specifically, lignin—an underrated bioresource that is mostly burned for energy production—has a tremendous potential as source of high-value chemicals. A promising lignin valorization requires an efficient extraction process able to recover high-purity lignin with considerable yields. Hence, we describe a protocol to extract lignin from a relevant agro-food residue—brewer’s spent grain—through the use of a deep eutectic solvent system, which has been proven an excellent potential to promote the fractionation of highpurity lignin from lignocellulosic material. The biomass is subjected to a hydrolysis at 120 °C for 5 h, and the lignin is further recovered by precipitation with distilled water. This protocol gives simple and detailed recommendations to effectively recover lignin from brewer’s spent grain contributing to the valorization of this abundant residue. Key words Lignin, Deep eutectic solvents, Brewer’s spent grain

1

Introduction Residues from the agro-food industry arise mainly from food losses, wastes, peels, seeds, pomace, leaves, bagasse, and prunings from fruits, vegetables, and cereals and account for 30% of the produced residues [1–3]. Most of these plant origin residues are mainly composed of lignocellulosic material, which is considered an abundant, cheap, and valuable renewable resource [4]. The carbohydrate fractions (cellulose and hemicellulose) are more studied and valorized when compared to lignin that is still an underestimated biopolymer [4], although the research interest has been consistently increasing over the last decade [5]. Lignin has a threedimensional amorphous structure with aromatic nature composed of three monomers p-hydroxyphenyl (H), guaiacyl (G), and syringyl (S) that derive of monolignols p-coumaryl, coniferyl, and sinapyl alcohol, respectively [6]. The monomers have distinct

Cristobal N. Aguilar Gonzalez et al. (eds.), Food Waste Conversion, Methods and Protocols in Food Science, https://doi.org/10.1007/978-1-0716-3303-8_5, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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arrangements according to the biomass type: softwoods have more G units, hardwoods have equal amounts of S and G units, and grasses present mostly H units [6, 7]. Besides the different arrangements, the amount of lignin also varies according to the type of biomass, with softwood and grasses presenting high and low amounts of lignin, respectively [8]. In order to achieve a promising lignin valorization, the recovery of lignin, specifically with a considerable yield and high purity, is relevant [9]. Some extraction methods are already employed at an industrial level (pulp and paper industry) [10], while other methods are used in a biorefinery context [4, 11]. Most of the studies on lignin from the agro-food residues, from the last decade, use mainly alkaline and organosolv methods [5] which may usually involve the use of toxic solvents, with harsh treatment conditions, in addition to needing combined pretreatments (e.g., autohydrolysis) [11, 12]. In order to overcome some of the problems related to the traditional methods, deep eutectic solvents (DES) have been suggested as a green alternative with the advantage of selectively solubilizing lignin [13]. DES are characterized by having a lower melting point than their components, which are always a hydrogen bond donor (HBD) and a hydrogen bond acceptor (HBA) [14]. DES are considered environmentally friendly due its high biodegradability, nontoxicity, and recyclability, being also easy to prepare, with adjustable properties [9, 15]. Type III DES are the most commonly used, consisting of choline chloride (CC) as HBA [9]. The present protocol proposes the application of a type III DES (lactic acid (LA):CC) to extract lignin from a relevant agrofood residue: brewer’s spent grain (BSG).

2 2.1

Materials Raw Material

(a) Brewer’s spent grain (BSG), kindly provided by Super Bock group (Porto, Portugal). (b) Oven. (c) Mill.

2.2

DES Preparation

(a) Lactic acid (≥ 85%; Sigma-Aldrich, Germany). (b) Choline chloride (98%; Sigma-Aldrich, Germany). (c) Stirring plate.

2.3 Biomass Pretreatment

(a) Raw material (BSG). (b) DES. (c) Heating plate (see Note 1).

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(d) Cloth filter. (e) Distilled water.

3

Lignin Recovery (a) Vacuum filter system (see Note 2). (b) Filter paper. (c) Oven.

4

Methods

4.1 Raw Material Preparation

(a) The BSG was oven dried at 65 °C, till constant weight. (b) The BSG was slightly milled to homogenize particle size. (c) The BSG waste was stored in plastic bags, protected from light and at room temperature, until used.

4.2

DES Preparation

(a) The DES solvent was prepared using LA (≥85%; SigmaAldrich, Germany) and CC (98%; Sigma-Aldrich, Germany) at a molar ratio of 5:1. (b) The LA was added to the CC and the mixture was left under agitation overnight. (c) The DES was considered prepared when the solution was transparent.

4.3 Biomass Pretreatment

(a) The DES solvent was added to the BSG biomass in liquid to solid ratio of 10 (wt.). (b) The mixture was thoroughly agitated until homogenized, so the DES was in contact with the biomass. (c) The hydrolysis was performed at 120 °C for 5 h. (d) The mixture was agitated hourly to ensure that all the biomass is in contact with the DES solvent (see Note 2).

4.4

Lignin Recovery

1. After the pretreatment, the DES liquor was separated from the solid residue using a cloth filter. 2. Distilled water was added to the DES liquor in a ratio of 4:1, and lignin was left to precipitate overnight at 4 °C. 3. The recovery of the precipitated lignin was performed by vacuum filtration using paper filter (see Notes 3 and 4). The precipitated lignin was oven dried at 50 °C until constant weight.

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4. The recovered lignin was weighted, and the yield was determined using the following equation: Lignin Yield ð%Þ =

5

ðmass of precipitated ligninÞ  100 ðmass of lignin in biomassÞ

Notes 1. Depending on the quantities, this step can be performed in an oven or an oil bath that can reach 120 °C. 2. If the plate allows stirring, pretreatment can be performed with constant stirring. 3. The lignin recovery can be performed through centrifugation. 4. It is possible to recycle the DES and use it for further extractions [16].

Acknowledgments This work was supported by National Funds from FCT—Fundac¸˜ao para a Cieˆncia e a Tecnologia—through project UIDB/50016/ 2020. Funding for author Ana C. Cassoni was via a PhD fellowship, administered by Fundac¸˜ao para a Cieˆncia e a Tecnologia, Portugal (ref. SFRH/BD/143198/2019). References 1. FAO (2019) The State of Food and Agriculture 2019. Moving forward on food loss and waste reduction 2. Peters K (2019) Technology of fruits and vegetable processing. ED-Tech Press, Waltham Abbey 3. Saha K, Dasgupta J, Chakraborty S, Antunes FAF, Sikder J, Curcio S, dos Santos JC, Arafat HA, da Silva SS (2017) Optimization of lignin recovery from sugarcane bagasse using ionic liquid aided pretreatment. Cellulose 24:3191– 3207 4. Bajwa DS, Pourhashem G, Ullah AH, Bajwa SG (2019) A concise review of current lignin production, applications, products and their environment impact. Ind Crop Prod 139: 111526 5. Cassoni AC, Costa P, Vasconcelos MW, Pintado M (2022) Systematic review on lignin valorization in the agro-food system: from sources to applications. J Environ Manag 317: 115258

6. Feofilova EP, Mysyakina IS (2016) Lignin: chemical structure, biodegradation, and practical application (a review). Appl Biochem Microbiol 52:573–581 7. Strassberger Z, Tanase S, Rothenberg G (2014) The pros and cons of lignin valorisation in an integrated biorefinery. RSC Adv 4: 25310–25318 8. Abdelaziz OY, Brink DP, Prothmann J, Ravi K, Sun M, Garcı´a-Hidalgo J, Sandahl M, Hulteberg CP, Turner C, Lide´n G, GorwaGrauslund MF (2016) Biological valorization of low molecular weight lignin. Biotechnol Adv 34:1318–1346 9. Hong S, Shen XJ, Xue Z, Sun Z, Yuan TQ (2020) Structure-function relationships of deep eutectic solvents for lignin extraction and chemical transformation. Green Chem 22:7219–7232 10. Gillet S, Aguedo M, Petitjean L, Morais ARC, Da Costa Lopes AM, Łukasik RM, Anastas PT (2017) Lignin transformations for high value

Protocol for the Extraction of Lignin from Brewer’s Spent Grain. . . applications: towards targeted modifications using green chemistry. Green Chem 19:4200– 4233 11. Wang H, Pu Y, Ragauskas A, Yang B (2019) From lignin to valuable products–strategies, challenges, and prospects. Bioresour Technol 271:449–461 12. Yoo CG, Meng X, Pu Y, Ragauskas AJ (2020) The critical role of lignin in lignocellulosic biomass conversion and recent pretreatment strategies: a comprehensive review. Bioresour Technol 301:122784 13. Francisco M, Van Den Bruinhorst A, Kroon MC (2012) New natural and renewable low transition temperature mixtures (LTTMs):

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screening as solvents for lignocellulosic biomass processing. Green Chem 14:2153–2157 14. Abbott AP, Boothby D, Capper G, Davies DL, Rasheed RK (2016) Experimental study of evacuated tube collector/storage system containing paraffin as a PCM. Energy 114:1063– 1072 15. Zhang Q, De Oliveira Vigier K, Royer S, Je´roˆme F (2012) Deep eutectic solvents: syntheses, properties and applications. Chem Soc Rev 41:7108–7146 16. Kumar AK, Sharma S, Shah E, Patel A (2018) Technical assessment of natural deep eutectic solvent (NADES) mediated biorefinery process: a case study. J Mol Liq 260:313–322

Chapter 6 Protocol for Antioxidant Dietary Fiber Determination: Structural Characterization and Quantification Taˆnia Braganc¸a Ribeiro, Marı´a Emilia Brassesco, Diva Santos, and Manuela Pintado Abstract Identifying the dietary fiber’s (DF) chemical composition (i.e., monosaccharides, lignin, uronic acids, phenolics) is essential for a better knowledge of the various health properties of the fiber. Thus, we propose a new approach that allows us not only to quantify the content of insoluble DF (IDF) and soluble DF (SDF) but also to estimate their chemical constituents, including their phenolic compounds and their respective antioxidant activity. The method can determine the chemical structure of DF resulting from four principal steps with chemical, enzymatic, and analytical procedures. The protocol can be completed in ~10 days, including ~5 days required for the fiber profile determination. Key words Antioxidant dietary fiber, Structural carbohydrates, Bound phenolic compounds

1

Introduction: Antioxidant Dietary Fiber Dietary fiber (DF) is a complex food component comprising various structural cell wall non-starch polysaccharides [1, 2]. Despite DFs being carbohydrate polymers with at least ten monomeric units, their composition varies substantially from one food source to another. However, their nondigestible characteristic, i.e., the incapacity of endogenous enzymes from the human small intestine to hydrolyze them, is a common property of all complex carbohydrates collectively grouped in the DF [1–3]. DFs are among the major components of plant foods, and their importance has emerged in the last years primarily by their beneficial effects on human health and diverse technological functions [4–6]. Conventionally, DF is divided into two categories, insoluble DF (IDF) and soluble DF (SDF), according to whether it can be dissolved in water [5, 6]. IDF and SDF composition exhibited significant compositional differences (neutral sugars and uronic acids, among others),

Cristobal N. Aguilar Gonzalez et al. (eds.), Food Waste Conversion, Methods and Protocols in Food Science, https://doi.org/10.1007/978-1-0716-3303-8_6, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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conferring entirely different physiological functions. For example, IDFs are mainly associated with regulating intestinal motility and reducing colon cancer risk, and SDFs, such as pectins, gums, β-glucans, fructooligosaccharides, and resistant starch, have common hypocholesterolemic effects [7]. Indeed, according to several authors, the classification of DF based on its main physiological effects might be more truthful [1, 8]. Phenolic compounds are a large variety of compounds in plants that share one or more phenol groups, including phenolic acids, flavonols, flavones, and others. These compounds are recognized for their potent antioxidant and free radical-scavenging properties, antimicrobial activity, anti-inflammatory effects, and other beneficial biological effects. Phenolic compounds usually are studied separately from DF. However, the phenolic compounds could also be considered DF components. Indeed, some researchers have pointed out that some of the health functionalities of DF usually associated only with its carbohydrate components are more probably due to the association of bound phenolics linked to DF [6, 8]. Bound phenolics associated with DF may remain unaltered through the gastrointestinal tract and reach the colon nearly intact. In the colon, they could undergo biotransformation by the gut microbiota and DF carbohydrate components acting as antioxidants and antimicrobial and prebiotic agents [9]. As a consequence of the importance of phenolics’ presence in DF, SauraCalixto (1998) suggested a new concept of “antioxidant dietary fiber” (ADF), which consists of a high DF source (higher than 50% on the dry basis) linked with natural antioxidants, mainly phenolic compounds [6, 10]. Determining the DF chemical structure (i.e., monosaccharides, lignin, uronic acids, phenolics) is fundamental to understanding fiber’s several health benefits. So, a methodology that allows not only measuring the content of IDF and SDF but estimating their chemical components, including their phenolic compounds and respective antioxidant activity, is proposed. 1.1 Overview of the Procedure

The DF chemical structure procedure is summarized in Fig. 1. It can be divided into four phases: pretreatments, enzymatic hydrolyses, SDF and IDF obtainment, and fiber profile determination with subsequent analysis. The pretreatments depend on the sample characteristics. The enzymatic hydrolyses involve the exposure of the sample to sequential enzymatic digestion by heat-stable α-amylase, protease, and amyloglucosidase. The SDF and IDF obtainment is estimated using the enzyme-gravimetric method, according to AOAC method 991.43 (1990), with slight modifications regarding SDF obtention followed by two different acid hydrolysis of IDF according to the NREL method [11] and acid hydrolysis in 6% sulfuric acid at 121 °C of SDF [12]. The last step is fiber profile determination and involves subsequent analyses to

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Fig. 1 Flow diagram of the DF chemical structure method. Timing and flow diagram of the DF chemical structure method for dried samples. The expected time frame in days (d) (left) and stages and corresponding step numbers in the Methods (right) are given. IDF, insoluble dietary fiber; SDF, soluble dietary fiber

quantify the fiber structure. Both hydrolysates are used to quantify uronic acids (UA) and neutral sugars (NS). Klason lignin (KL) and the resistant protein (RP) are also determined in IDF. The phenolic compounds were released from IDF and SDF using a hydrolysis process with NaOH [13] followed by its spectrophotometric analyses to measure total phenolic compounds (TPC) by the FolinCiocalteu and antioxidant activity. The identification and quantification of phenolic compounds could also be attained chromatographically.

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Materials

2.1 Pretreatment of Sample 2.2 Enzymatic Hydrolysis

1. n-Hexane. 2. Ethanol 80%. 1. MES-TRIS buffer, 0.05 M each, pH 8.2. 2. Total dietary fiber kit (TDF100A-1KT—Megazyme). 3. 0.561 N HCl solution. 4. 5% NaOH solution to raise pH and 5% HCl to drop pH.

2.3 Separation, Obtainment, and Washing of Soluble (SDF) from Insoluble Dietary Fiber (IDF)

1. Water at 70 °C.

2.4 Fiber Profile Determination

1. 72% H2SO4.

2. 96% ethanol. 3. Acetone. 4. Dialysis membrane with a molecular weight cutoff 12,000 Da.

2. 6% H2SO4. 3. Sugar standards: D-(+)glucose, D-(+)xylose, D-(+)galactose, L-(+)arabinose, and D-(+)mannose. 4. CaCO3. 5. 0.22 μM syringe filter. 6. Aminex Carbo-P, Bio-Rad. 7. Aminex HPX-87P heavy metal, 300 × 7.8 mM, Bio-Rad. 8. Ultrapure water. 9. 4 M NaOH. 10. 6M HCl. 11. Ethyl acetate. 12. Anhydrous Na2SO4. 13. Boric acid-sodium chloride solution: Dissolve 0.3 g of boric acid and 0.2 g of sodium chloride in 10 mL of distilled water. Dissolve 14. Dimethylphenol solution: 10 mg 3,5-dimethylphenol in 10 mL of acetic acid glacial.

3

of

Methods The quantification of antioxidant dietary fiber is determined on triplicate samples of dried materials. With each assay, run two blanks along with samples to measure any contribution from reagents to the residue.

Protocol for Antioxidant Dietary Fiber Determination. . .

3.1 Pretreatments of Sample

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A. Defatting (Only to Samples with Fat Content > 10%) 1. Weigh 1–2 g (2 g to low-fiber content samples) of the dried sample into 50 mL centrifugation tube. 2. Add 25 mL of n-hexane and place centrifuge tubes in an ultrasonic water bath for 10 min. 3. Centrifuge at 5000 RPM for 5 min and collect the supernatant. 4. Repeat the extraction process. B. Remotion of Low Molecular Weight Carbohydrates (Monosaccharides and Oligosaccharides) 1. Extract the dried sample or n-hexane extraction residue with 20 mL of 80% ethanol. 2. Place centrifuge tubes in an ultrasonic water bath for 10 min. 3. Centrifuge at 5000 RPM for 5 min and collect the supernatant. 4. Repeat the extraction process two more times. 5. Leave centrifuge tubes in the fume hood overnight to dry the residue.

3.2 Enzymatic Hydrolyses (AOAC Method 991.43 (1990))

1. Transfer each sample from the centrifuge tube to 600 mL beaker or Schott flask. 2. Dispense 40 mL MES-TRIS buffer, 0.05 M each, pH 8.2 at 24 °C. 3. Add 50 μL heat-stable α-amylase. 4. Incubate beakers (cover each beaker with aluminum foil squares) or Schott flask in a water bath at 98–100 °C with continuous agitation for 30 min. 5. Cool beakers to 60 °C. 6. Scrape the beaker wall with a spatula, if necessary. 7. Rinse with 10 mL water. 8. Adjust the temperature of the water bath to 60 ± °C. 9. Add 100 μL protease (no pH adjustment). 10. Place beakers on the water bath with continuous agitation for 30 min. 11. Remove sample beakers from the water bath. 12. Add 5 mL of 0.561 N HCl solution into each sample, stir, and check pH (pH should be 4.1–4.8). 13. Adjust pH, if necessary. Use 5% NaOH solution to raise pH and 5% HCl solution to drop pH.

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14. Add 200 μL amyloglucosidase. 15. Incubate in a shaking water bath at 60 °C for 30 min. 3.3 SDF and IDF Obtaining

A. Separation of Soluble (SDF) from Insoluble Dietary Fibre (IDF) 1. Filter through crucible n°2 (tared and weighed). 2. Wash with 2 water portions (10 mL) at 70 °C (use water to rinse the beaker before washing residue). 3. Save filtrate and water washings for determination of SDF. B. IDF Washing 1. Wash residue with 20 mL of 96% ethanol followed by 20 mL of acetone. 2. Oven-dry crucibles at 45 °C for 24 h for IDF analysis. 3. Cool the crucibles in a desiccator for approximately 1 h. 4. Weigh crucible containing dietary fiber residue. 5. Store the residue for IDF profile analysis. C. SDF Obtainment 1. Dialyze filtrate and water washings with dialysis membrane with a molecular weight cutoff 12,000 in 1.5 L of distilled water for 48 h. 2. Replace water after 12 h. 3. Freeze-dry samples after dialysis. 4. Weigh the freeze-dried sample. 5. Store the freeze-dried sample for SDF profile analysis.

3.4 Fiber Profile Determination

A. IDF Fiber Treatment 1. Weigh 300 mg of IDF sample to pressure tube (run the analysis in quadruplicate). 2. Add 3 mL of 72 % H2SO4. 3. Put IDF samples in the water bath (30oC) for 60 minutes (stirring every 5–10 min). 4. Transfer to 250 mL Erlenmeyer. 5. Add 84.00 ± 0.04 mL of deionized water to dilute the acid to a 4% concentration. 6. Autoclave at 121 oC for 1 h. 7. Filtrate with tared and weighed crucibles n°2. 8. Store a 5–10 mL sample to analyze the neutral sugars and uronic acids. B. SDF Fiber Treatment 1. Put at least 40 mg of the sample into a 100 mL pressure tube and cap it tightly. 2. Place the tubes in a safe autoclave rack.

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3. Hydrolyze samples using 12 mL of 6% H2SO4 in the autoclave at 121 °C for 1 h. 4. Store 5–10 mL to neutral sugars and uronic analysis. C. Preparation of a Set of Sugar Recovery Standards (SRS) 1. Weigh D-(+)glucose, D-(+)xylose, D-(+)galactose, L-(+) arabinose, and D-(+)mannose in concentrations closely resembling the concentrations of sugars in the test sample. 2. Add 10.0 mL deionized. 3. Add 348 μL of 72% sulfuric acid. 4. Transfer the SRS to a pressure tube and cap tightly. 5. Place the tubes in a safe autoclave rack. 6. Autoclave at 121 °C for 1 h. D. Klason Lignin Determination (Only in IDF) 1. Oven-dry the crucibles overnight at 105 °C (at least 16 h). 2. Record the weight of the crucibles. 3. Reserve two crucibles (RP) determination.

for

the

resistant

protein

4. Ash at 525 °C for 5 h and reweigh crucibles. 5. RP was estimated by micro-Kjeldahl method using a nitrogen-to-protein conversion factor of 6.25. 6. Klason lignin (KL) was the weight of residue after drying, subtracting the ash and RP. E. Neutral Sugar Analysis 1. Prepare a series of calibration standards, including D-cellobiose, D-(+)glucose, D-(+)xylose, D-(+)galactose, L-(+)arabinose, and D-(+)mannose with four-point calibration in concentrations ranging between 0.1 and 4.0 mg/ mL. 2. Neutralize using CaCO3 SDF hydrolysis liquid, IDF filtrates, and SRS until pH 5–6. 3. Allow to settle and decant off the supernatant. After settling, a pH of 7 will be reached. 4. Filtrate standards and neutralized solutions with a 0.22 μM syringe filter. 5. Transfer to an autosampler vial, seal, and label the vial. 6. Analyze the calibration standards, CVS, and samples by HPLC using the following conditions: Micro-guard column: Aminex Carbo-P, Bio-Rad. Carbohydrate analysis column: Aminex HPX-87P heavy metal, 300 × 7.8 mM, Bio-Rad.

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Injection volume: 10–50 μL, dependent on concentration and detector limits. Column temperature: 80–85 °C. Flow rate: 0.6 mL/min. Detector: Refractive index (RI). Run time: 35 min. 7. For the sugar recovery standards (SRS), calculate the amount of each component of sugar recovered after dilute acid hydrolysis, accounting for any dilution made prior to HPLC analysis. Average any replicate (%Rsugar) values obtained for each sugar. %Rsugar =

Conc: detected by HPLC, mg=mL × 100 Conc: of sugar before hydrolysis, mg=mL

8. Calculate the sugar concentration values obtained by HPLC for each hydrolyzed sample, reporting the dilution made prior to HPLC analysis and applying the %Rsugar values. Cx =

Conc: detected by HPLC, mg=mL × dilution factor × 100 %Rsugar =100

F. Uronic Acid Analysis Uronic acids (UA) were determined colorimetrically by adapting the 3-hydroxydiphenyl method of Blumenkrantz and Asboe-Hansen (1973) with d-galacturonic acid as standard. 1. Mix 250 μL of SDF hydrolysis liquid or IDF filtrates with 250 μL of boric acid-sodium chloride solution and 4 mL of 96% H2SO4. 2. Vortex the mixture and incubate in a water bath at 70 °C for 40 min. 3. Cool tubes at room temperature. 4. Add 200 μL of dimethylphenol solution and mix using a vortex. 5. Measure the absorbance at 400 nM and 450 nM. 6. The absorbance difference between 450 and 400 was quantified as galacturonic acid equivalent (stock solution, 200 μg/mL; standard curve concentrations,150, 100, 50, 25, 15, and 5 μg/mL). G. Bound Phenolic Compounds 1. Hydrolyze IDF and/or SDF sample (250–500 mg) using 4 M NaOH (8 mL) at room temperature on an orbital shaker at 250 rpm (4 h).

Protocol for Antioxidant Dietary Fiber Determination. . .

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2. Acidify to pH 1.5–2.0 using 6 M HCl. 3. Centrifugate (8000 RPM, 30 min). 4. Extract supernatant 5 times with ethyl acetate (10 mL). 5. Dry ethyl acetate using anhydrous Na2SO4 (funnel with cotton and a small amount of Na2SO4 to eliminate water residues). 6. Evaporate extracts until dry using a rotary vacuum evaporator at 30 °C. 7. Dissolve the resulting residue in methanol or other solvents to a final volume of 5 mL, and store at -20 °C until total phenolic analysis. 8. Aliquots of the extracts were used for spectrophotometric measurement by the Folin-Ciocalteu and ABTS or ORAC methods to determine total phenolic compounds (TPC) and antioxidant activity, respectively [4]. 9. The phenolic compounds could also be identified by HPLC [15].

4

Notes 1. The 80% ethanol supernatant could be used to determine soluble sugars and organic acids after ethanol evaporation in a rotary vacuum evaporator by HPLC-IR and HPLC-UV, respectively, or other sugar and organic acid quantification methodology. 2. The drying step between the remotion of simple carbohydrates and the enzyme treatment is essential to eliminate any ethanol residual. 3. The dialysis was applied to avoid the error caused by precipitating DF with ethanol [16, 17]. 4. A fresh SRS is not required for every analysis. A large batch of sugar recovery standards may be produced, filtered through 0.2 μM filters, stored in a freezer, and removed when needed. 5. A fresh set of standards is not required for every analysis. A large batch of standards may be produced, filtered through 0.2 μM filters, stored in a freezer, and removed when needed. 6. In the neutralization step of neutral sugar analysis, samples should never be allowed to exceed a pH of 9 to avoid a possible loss of sugars. 7. Check test sample chromatograms for the presence of cellobiose, oligomeric sugars, and sugar degradation:

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(a) Levels of cellobiose greater than 3 mg/mL indicate incomplete hydrolysis. (b) The presence of peaks eluting before cellobiose (retention time of 4–5 min using recommended conditions) may indicate high levels of sugar degradation products indicative of over-hydrolysis. All samples from batches showing evidence of incomplete or over-hydrolysis should have fresh samples hydrolyzed and analyzed.

Acknowledgments Thanks are due to Fundao para a Cieˆncia e a Tecnologia (FCT)/ MCT for the financial support for the Associate Laboratories CBQF (UIDB/50016/2020) through national founds and, where applicable, co-financed by the FEDER, within the PT2020 Partnership Agreement. The author Diva Santos would also like to thank FCT for her PhD grant (SFRH/BD/143493/2019). References 1. Johnson IT (2020) Fiber. In: Present knowledge in nutrition. Elsevier, pp 515–529 2. Xu J, Li Y, Zhao Y et al (2021) Influence of antioxidant dietary fiber on dough properties and bread qualities: a review. J Funct Foods 80: 104434. https://doi.org/10.1016/j.jff.2021. 104434 3. Jiang G, Bai X, Wu Z et al (2021) Modification of ginseng insoluble dietary fiber through alkaline hydrogen peroxide treatment and its impact on structure, physicochemical and functional properties. LWT 150:111956. https:// doi.org/10.1016/j.lwt.2021.111956 4. Ribeiro TB, Campos D, Oliveira A et al (2021) Study of olive pomace antioxidant dietary fibre powder throughout gastrointestinal tract as multisource of phenolics, fatty acids and dietary fibre. Food Res Int 142:110032. https:// doi.org/10.1016/j.foodres.2020.110032 5. Li N, Feng Z, Niu Y, Yu L (2017) Structural, rheological and functional properties of modified soluble dietary fiber from tomato peels. Food Hydrocoll:1–9. https://doi.org/10. 1016/j.foodhyd.2017.10.034 6. Liu S, Jia M, Chen J et al (2019) Removal of bound polyphenols and its effect on antioxidant and prebiotics properties of carrot dietary fiber. Food Hydrocoll 93:284–292. https:// doi.org/10.1016/j.foodhyd.2019.02.047

7. Yoshida BY, Prudencio SH (2020) Alkaline hydrogen peroxide improves physical, chemical, and techno-functional properties of okara. Food Chem 323:126776. https://doi.org/10. 1016/j.foodchem.2020.126776 8. Stephen AM, Champ MMJ, Cloran SJ et al (2017) Dietary fibre in Europe: current state of knowledge on definitions, sources, recommendations, intakes and relationships to health. Nutr Res Rev 30:149–190. https:// doi.org/10.1017/S095442241700004X 9. Ribeiro TB, Costa CM, Bonifa´cio Lopes T et al (2021) Prebiotic effects of olive pomace powders in the gut: In vitro evaluation of the inhibition of adhesion of pathogens, prebiotic and antioxidant effects. Food Hydrocoll 112: 1 0 6 3 1 2 . h tt p s : // d o i . o r g /1 0 . 1 0 1 6 / j . foodhyd.2020.106312 10. Saura-Calixto F (1998) Antioxidant dietary fiber product: a new concept and a potential food ingredient. J Agric Food Chem 46:4303– 4306. https://doi.org/10.1021/jf9803841 11. Sluiter A, Hames B, Ruiz R et al (2012) NREL/TP-510-42618 analytical procedure – determination of structural carbohydrates and lignin in Biomass. Lab Anal Proced 17:NREL/ TP-510-42618 12. Bravo L, Saura-Calixto F (1998) Characterization of dietary fiber and the in vitro indigestible

Protocol for Antioxidant Dietary Fiber Determination. . . fraction of grape pomace. Am J Enol Vitic 49: 135–141 13. Xie P-J, Huang L-X, Zhang C, Zhang Y-L (2015) Phenolic compositions, and antioxidant performance of olive leaf and fruit (Olea europaea L.) extracts and their structure–activity relationships. J Funct Foods 16:460–471. https://doi.org/10.1016/j.jff.2015.05.005 14. Blumenkrantz N, Asboe-Hansen G (1973) New method for quantitative determination of uronic acids. Anal Biochem 54:484–489. https://doi.org/10.1016/0003-2697(73) 90377-1 15. Ribeiro TB, Oliveira A, Campos D et al (2020) Simulated digestion of an olive pomace water-

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soluble ingredient: relationship between the bioaccessibility of compounds and their potential health benefits. Food Funct 11:2238– 2254. https://doi.org/10.1039/c9fo03000j ˜ as E, Bravo L, Saura-Calixto F (1994) 16. Man Sources of error in dietary fibre analysis. Food Chem 50:331–342. https://doi.org/10. 1016/0308-8146(94)90201-1 17. Deng Q, Penner MH, Zhao Y (2011) Chemical composition of dietary fiber and polyphenols of five different varieties of wine grape pomace skins. Food Res Int 44:2712–2720. https://doi.org/10.1016/j.foodres.2011. 05.026

Chapter 7 Use of Ultrasound Technology for Food Waste Breakdown Martina de la Rosa-Herna´ndez, M. Carmen Gutie´rrez-Sa´nchez, Diana B. Mun˜iz-Ma´rquez, Abigail Reyes-Munguı´a, and Jorge E. Wong-Paz Abstract Ultrasound technology use is a promising strategy for the exploitation of food waste by inducing structural changes on food matrices, favoring their decomposition. In this chapter we will be describing selected experimental methodologies that allow us to prove the influence of ultrasound on the waste decomposition, inducing its reduction and revaluation as a substrate to other applications. The conditions and recommendations of proposed method are also described. Our work aims to encourage the reader about the advantages of using ultrasound to minimize environmental and economic impact generated by food waste. Key words Ultrasound technology, Food matrix, Decomposition, Revaluation, Food waste

1

Introduction By-products or wastes originating from food activities such as agriculture, fishing, and commerce are not fully valued, as they easily accumulate and pollute the environment. Recent reports indicate that more than 931 million tons of food waste are generated worldwide from household, commercial, and other food services [1]. In contrast, industry, for example, has recorded a lower rate of waste generated of approximately 17 million produced during industrial processing. Faced with this problem, an alternative option arises for the use of most of these residues, which are a natural source of molecules of interest to the pharmaceutical, food, and cosmetology industries. These strategies include the use of these plant materials to extract important molecules or compounds contained in them that are applicable in various industrial sectors and research. Some traditional methods such as maceration, Soxhlet extraction, and distillation have made it possible to use residues to obtain and preserve most of the biological properties of food components [2].

Cristobal N. Aguilar Gonzalez et al. (eds.), Food Waste Conversion, Methods and Protocols in Food Science, https://doi.org/10.1007/978-1-0716-3303-8_7, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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Fig. 1 Release of bioactive compounds by acoustic cavitation in ultrasound

However, these methods seemed to be very promising, until they were not able to allow the highest yield of molecules of interest, in conjunction to the challenges involved in their successful development, extraction time, and indiscriminate use of organic solvents [3]. Therefore, more sophisticated and less conventional technologies using ultrasound waves, microwaves, solvent pressurization, and others [4] have been implemented to improve the extraction of bioactive molecules from discarded leaves, seeds, stems, and husks of fruits, vegetables, and plants. Ultrasound technology has been applied in different ways in the industry, such as filtration, freezing/crystallization, thawing, preservation, drying, foaming, cooking, emulsification, cutting, sterilization, rehydration, and component extraction [38]. The latter has been termed ultrasound-assisted extraction (UAE), which is an innovative and effective technology to recover high value-added molecules using food waste by achieving its reduction and minimizing the indiscriminate consumption of organic chemicals [9]. The method is based on the release of compounds through acoustic cavitation, that is, ultrasonic waves capable of breaking the cell wall of any solid surface (Fig. 1). It is interesting to study the effect of bubbles produced in cavitation and sound waves, since they manage to induce in isolation or together phenomena such as fragmentation and localized erosion, form pores, induce shear force, and cause greater

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absorption and swelling index in the plant cell of the matrix used [5]. In addition, the ultrasound technique requires simple and semiautomated equipment for its development where those phenomena mentioned above occur allowing to improve extraction yields against traditional methods [6]. Likewise, the success of the method involves the relationship of aspects that must be controlled such as the power and frequency of the ultrasonic waves, the solvent to be used, and the time, temperature, and duration of the extraction of the components. Ultrasound technology has allowed the isolation of secondary metabolites from by-products such as coffee (Coffea) husk [7], pomegranate (Punica granatum), roots such as ginger (Zingiber officinale), leaves of medicinal plants such as moringa (Moringa oleifera), chaya (Cnidoscolus aconitifolius), and others such as Jatropha dioica, Flourensia cernua, Turnera diffusa, and Eucalyptus camaldulensis [8]. Some important bioactive components obtained with UAE have been polyphenols, flavonoids [7], carotenoids [11], and polysaccharides [10] including fatty acids [12]. Likewise, UAE presents advantages during the realization of this technique, such as cost and time reduction of the process, as well as favoring a higher yield quantity and quality of the extracts. Based on the above, the following work aims to present to the reader the characteristics and important aspects of the selected protocols that use ultrasoundassisted extraction in the decomposition of food waste, induction of changes, up to reducing the environmental and economic impact.

2

Material

2.1 Equipment and Reagents

Ultrasound-assisted extraction (UAE) requires a digital ultrasonic bath (Branson® CPXH ultrasonic cleaner digital timer with heat) with a useful volume of 10 L and a convection drying oven (ECOSEL®) model 9025 h. The solvents used were distilled water (H2O) and acetone (C3H6O) at 70% v/v.

2.2 Selection of Plant Material

For ultrasound-assisted extraction, different residues derived from the food industry can be used, especially vegetables, fruit peels such as pomegranate, a variety of citrus fruits, red fruits, blueberries [14], and litchi [7], being widely used. Edible grains can also be treated with ultrasound with the intention of obtaining essential oils from some edible seeds such as cocoa, sunflower seed, sesame, pumpkin, chili, radish, and other vegetables [25]. Some less researched strategies are the use of ornamental and medicinal plants treated with ultrasound, since they are usually prepared by maceration, according to the conventional methods used by traditional medicine to extract compounds with important biological properties.

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2.3 Preparation of Plant Material

3

Once the food waste was selected, it was monitored so that all samples are prepared using the same experimental conditions. The residues were dehydrated in an air convection drying oven at temperatures of 45–60 °C for 2 days, based on the moisture percentage value of the samples. Subsequently, the samples were pulverized and sieved to a uniform particle size.

Methods

3.1 UltrasoundAssisted Extraction (UAE) for by-Products and Other Wastes

Metabolites called procyanidins were obtained from different commercially available roasted coffee beans according to Wong-Paz et al. [8] with some modifications. For ultrasound-assisted extraction (UAE), 20 g of the plant material, water, and 70% (v/v) acetone were used as solvents. The kinetics were from 0 to 20 min in an ultrasonic bath (model 2510, Brason) with temperatures programmed in the equipment of 20 °C and 45 °C. The first 5 min were compared with a percolation extraction under the same conditions. In the results it was observed that the UAE increased almost 50% of the yields of the coffee samples compared to the traditional method; however, there was no significant difference between the brands evaluated with a value of p < 0.05. On the other hand, Fig. 2 evidenced how it is possible to improve almost 100% of the yields of compounds obtained using 70% acetone as solvent in comparison to aqueous UAE. The implementation of the solvent has facilitated the extraction and allowed finding significant differences (p < 0.05) between the best compound contents expressed in milligram equivalents of procyanidin C1/g (mg EPC1/g) dry base of regional (5.78 ± 0.2), national (10.73

Fig. 2 Extraction of procyanidins from roasted coffee beans for 5 minutes. Validation UAE (lower case) and percolation (upper case)

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Fig. 3 Effect of temperature on ultrasound-assisted extraction kinetics (UAE-acetone) of coffee bean procyanidins at 20 °C (dotted line) and 45 °C (solid line)

± 0.3), and international (19.62 ±0.8). Also, the UAE-acetone proposed by Wong-Paz et al. [7] was evaluated under the effect of temperature for the extraction of procyanidins using roasted coffee bean to further improve yields. The results are shown in Fig. 3, where it is confirmed that temperature is a crucial factor in the release of the compounds studied. This is since temperature influences the direction of the ultrasound wave, since at high temperatures the ultrasound will have greater vertical direction toward the cell wall of the plant material that has been exposed to cavitation considerably increasing the yields [21]. In addition to the protocol described above, there are other proposals on the application of UAE using food products and wastes. For example, in Table 1, we describe some established methods as strategies in the utilization of various types of food wastes for obtaining bioactive components and the considerable reduction of these wastes. 3.2 Evaluation of the Metabolites Obtained

Extracts obtained by UAE can contain a higher number of bioactive components as the cell wall of the sample is breached by wave cavitation action, whereas extraction is achieved with gas- or liquid-phase solvents [43]. Both molecules anchored to the cell wall, and those located intercellularly have been identified using quantitative assays, the most common of which seeks the identification of molecules with high added value such as polyphenols (phenolic acids, flavonoids, and tannins), terpenes, and alkaloids [29]. The main assays selected for the determination of these metabolites from plants, fruits, and vegetables are the following: the Folin-Ciocalteu method for hydrolyzed polyphenols, the HCl-butanol technique for condensed tannins [30], and total flavonoids (n-quercetin method) [31].

Shells of: Handle

Sweet fruits

Banana

Melon Watermelon

Litchi Sonication time = 5, 20 min Sonication amplitude = 40–70% Blueberry pulp Blueberries Sonication temperature = 61.03 °C Sonication time = 23.67 min Strawberry pulp Strawberries Solvent: Acetone = 40%, 60%, 80% v/v Sonication time = 10 and 15 min Sonication temperature = 20, 35, 50 °C Pomegranate peel Grenada Solvent = distilled water (1:4 w/v) Sonication amplitude = 20, 60, and 100% Sonication time = 5, 10, 15 min Coffee pulp Coffee pulp Solvent = acetone (70% v/v) Sonication time = 20 min Room temperature Handle Sonication time = 15, 30, 45 min Frequency = 50 kHz, power 160 W Solvent = distilled water Sonication temperature = 25 °C Melon Solvent = ethanol (42%) Sonication temperature = 47.82 °C Sonication time = 31.63 min Banana Solvent = methanol, ethanol, and acetone (25%, 50%, 75%, 100%) Sonication temperature = 35 °C, 45 °C, 55 °C Sonication time = 60 min

Litchi husk and seed

Red fruits

UAE conditions

Type of waste

Food raw material

Table 1 Use of food waste by the pharmaceutical and food industry using ultrasound technology

[7]

Phenolic acids

[18]

[17]

[16]

[15]

Procyanidins, and

Preservatives, flavorings, colorings, antimicrobials, and antioxidants

[14]

[13]

[22]

Authors

Anthocyanidins,

Antioxidant and chemoprotective bioactive compounds such as

Extraction components

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Shellfish and crustaceans

Citrus

Sour fruits

Shrimp and Fish

Grapefruit Tanjarina

Lemon

Shrimp shell Solvent = petroleum ether, n-hexane, ethanol, acetone Sonication temperature = 25 to 45 °C Sonication time = 5 to 15 min Sonication amplitude = 20–100%

Amines, nutrients, pigments, carotenoids such as astaxanthin

Colorants, flavorings, vitamins, essential oils, fragrances, diuretics, antivirals, antimicrobials, antioxidants, and phenolic acids

Orange Sonication power = 100, 250, 400 W Solvent = 0%, 25%, 50% ethanol Sonication time = 5, 17.5, 30 min Mandarin Sonication frequency = 20 kHz Solvent = 50, 60% ethanol, acetone Sonication temperature = 50 ± 0.5 °C Sonication time = 5–15 min Citrus pomace Particle size (1.40–2.80 mm) Sonication time = 10–60 min Sonication temperature = 23–50 °C Ultrasonic power (150–250 W)

Shell of: Orange

Antimicrobial compounds, antioxidants, prebiotics, and enzymes

Shell and seed of: Tamarindo: Tamarindo Sonication amplitude = 25% and 50% Sonication time = 15, 30 min Room temperature Pitahaya pulp Pitahaya Pineapple peel Sonication temperature = 30–70 °C Solvent = 30, 60%. Sonication time = 5–25 min

(continued)

[26] [27]

[24]

[23]

[22]

[20]

[19]

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Peanuts and shells Mani Pumpkin seed Sonication time = 31.2 min Sunflower seed Sonication temperature = 36.6 °C Solvent = ethanol 93.2%

Edible grains

Antioxidant, antiaging compounds For food

Bioactive compounds and nutrients

Vegetables residues Sonication frequency = 24 kHz Sonication amplitude = 69.7% Sonication temperature = 53.43 °C Sonication time = 12 min

Spinach Broccoli Cauliflower Lettuce Asparagus Potato Tomato

Vegetables

Extraction components

UAE conditions

Type of waste

Food raw material

Table 1 (continued)

[25]

[5] [28]

Authors

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3.3 Yields and Decomposition of Food Wastes

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New technologies and the increasing demand for food products enriched with healthy ingredients have enabled industrial adaptation and research into cost-effective and less polluting extraction methods for sustainable processing of food waste in favor of the circular economy. Therefore, ultrasound technology is still being studied with the aim to know the advantages not only in yield rates but also in ideal and reproducible conditions for the reduction treatment of many wastes [27]. Recent works have published the reduction of food waste and residues to levels from 50% to 0% to be converted into value-added products, such as functional ingredients, bioactive compounds, biofuels, or bioplastics (Fig. 4) for the generation of new economic income for the industry [32]. Experimental designs have provided insight into the ideal conditions for the use of ultrasound in waste reduction with improved yields. For example, Maran and Priya [33] successfully employed and evaluated the efficiency of UAE in the extraction of natural pigments using red beet stalk waste by Box-Behnken response surface design. The authors propose an extraction at 53 °C, ultrasonic power of 89 W, sonication time of 35 min, and a solid/liquid ratio of 1:19 g/mL if yields greater than 1.29 mg/g betacyanin and 5.32 mg/g betaxanthin are desired.

Fig. 4 The effectiveness of ultrasound in the revalorization of food waste and the development of new products

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For obtaining antioxidants, Esteban-Lustres et al. [34] established a UAE optimization protocol for the recovery of antioxidant compounds from dehydrated fruit residues using Box-Behnken response surface design. The best concentrations and antiradical effect occurred in the extracts obtained under an operation of a sonication frequency of 24 kHz, a solid/liquid ratio of 1:15, an amplitude of 69.7% at a sonication temperature of 43 °C, and 12 min duration. Optimization of pectin polysaccharide extraction from oat husk has been successfully performed using ultrasound technology with ethanol for 60 min [35]. On the other hand, obtaining essential oils with ultrasound has also been optimized from sunflower husk and seed [36], with papaya seed [37] and other spices [36]. Moradi et al. [35] determined that to obtain better yields of sunflower seed oil rich in α-linolenic acid, UAE with hexane is used, with a solid/ liquid ratio of 1:12, in a sonication time of 120 min, an ultrasound frequency of 24 kHz, and temperature of 50 °C. 3.4 Structural Changes Induced by UAE

The good yields recorded in the literature by the effect of ultrasound using a variety of samples originate when the cavitation bubbles formed by the ultrasonic waves collapse causing them to collide and collide in an accelerated manner fragmenting the cellular structure of the plant material [38]. Therefore, the cell wall is compromised by its rapid fragmentation, resulting in solubilization of internal and external molecules in the solvent used. In addition, another phenomenon observed is the decrease in the particle size of the solid material itself reducing considerably the food waste [39]. However, not only the degradation of the plant material is possible, but also internal and external structural changes are created in the cell of the food matrices that can benefit the food in some of its properties or affect them. Some changes studied in food matrices and residues by ultrasound effect are the vibrations that favor cell permeability and solvent transfer internally and externally reducing the ultrasonic treatment time and facilitating the consecutive processes of filtration of liquid samples or juices [40]. Also, the uniform heat transfer and diffusion of the ultrasonic bath allow to modify the freezing points improving the preservation of the microstructure of the plant and even animal cell (if they were matrices or meat waste), thus opening better and faster cryopreservation points of the solids or extracts subjected to ultrasound [45]. On the contrary, there is also evidence that once the ultrasound-treated food matrices or wastes are thawed, they will thaw rapidly, preserving most of the physical and chemical characteristics, if the ultrasonic treatment times to which the microstructure is subjected are respected [42]. Cell softening is another effect on the structure of the material under ultrasonic effect [41]. All these modifications have been useful in the food industry and in research focused on the extraction of bioactive compounds.

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However, some modifications go beyond the cell wall of the plant because it has been observed that ultrasonic waves structurally degrade the molecules that have been extracted from the cell interior once they are solvent-bound, mainly if they are exposed to long periods of sonication and high temperatures [46]. In addition, it is important to mention that the type of molecule to be extracted, as well as the structure, cell wall resistance, and the effects of pressure, temperature, and time, determines the effect on the integrity of the metabolite to ultrasound. There are several phenomena that produce changes in the molecular structure of a compound, mainly the reactions generated during the cavitation of the waves that form bubbles, which when bursting manages to generate the formation of free radicals because of cellular rupture because of the constant temperature and pressure [41]. Therefore, it is advisable to use a low intensity of ultrasound to avoid the formation of these radicals, which can lead to lower yields on the recovery of antioxidant molecules that are fulfilling their biological activity during the extraction procedure. When working with bioactive compounds, they are often susceptible to high temperatures; isomerization effects of the molecules have been found when subjected to temperature gradients from 20 °C [47]. For their part, phenolic compounds are stable at temperatures below 40 °C during ultrasound treatments. However, thermal exposures exceeding values of more than 50 °C led to the degradation of most molecules [48]. Examples of compounds that have been studied for their degradation during ultrasound procedures are lutein; vitamins, such as cyanocobalamin; α-tocopherol; and organic and phenolic acids, such as chlorogenic acid, gallic acid, and others based on the type of sample to be used [49]; only condensed tannins, such as anthocyanidins, remain stable above 50 °C as long as they are not subjected to high pressure and temperature gradients since the interaction between the solid and liquid interfaces becomes violent [44, 50]. So that the reactions that occur by UAE can modify the bioactive components present in most food waste, it is necessary to experiment under mathematical models the ideal extraction conditions based on the type of plant material, chemical structure, functional group, time, and other physicochemical characteristics of the compound or compounds present in these. 3.5 Reduction and Revalorization of Food Wastes

The reduction of food waste is nowadays necessary in all underdeveloped and developed countries in order to avoid the pollution of the few aquatic and terrestrial ecosystems preserved so far. As for the urban context, the large quantities of food waste from the population and industries have become a difficult problem to solve. Therefore, in the last decade, the importance of creating

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sustainable alternatives that help to minimize this waste goes hand in hand to remedy the growing global warming and climate change. Some strategies that have been proposed and implemented are the use of green technologies that not only allow the rapid reduction of these wastes in quantity but also provide an economic value and an application to benefit various needs of society, for example, the use of food waste to produce biofuel, the obtaining of metabolites with bioactive properties, and their inclusion in the development of many products that benefit health, among others. One of the most researched topics in the revaluation of food waste is its use for the development of functional foods that include compounds with antioxidant, antibacterial, antifungal, anti-obesity, cardioprotective, chemoprotective, and even carcinogenic properties [51]. However, none of these applications would be possible without newly developed technologies that seek to optimize, reduce, and revalorize waste. This chapter has discussed the application of ultrasound technology for different purposes; however, one has been of vital importance to solve the overgrowth of food waste. The application of ultrasound for the reduction and revaluation of food waste is based on the ability of ultrasonic waves to degrade plant material from a volume of starting material and reducing the particle size [52]. The reduction of by-products using ultrasound to enhance enzymatic hydrolysis processes with different food waste as a function of time and temperature was successfully proven, in addition to generating economic value from the components obtained from each plant material. For example, Dangles and Fenger [48] treated shell waste for 24 hours with ultrasound and a proteolytic enzyme, this pretreatment allowed the considerable reduction of the waste from the perforation of the waste and the continuous diffusion of the enzyme in the wall of the food material. Other researchers, such as Zhang et al. [52], optimized based on a Taguchi experimental design the use of ultrasonic atomization for solid waste reduction during real-time industrial production. This system could be applied to solid food waste produced during industrialization. The use of ultrasound in the development of energy-efficient methods to convert food and agricultural wastes has been evidenced in current refereed reviews, such as the one by Wu et al. [53], where the efficiency of the system to obtain high value-added metabolites in less time and produce clean energy through biorefineries is described. Finally, the reduction of plant particles in the treated materials or during pretreatment with ultrasound will be evidenced from the implementation of techniques such as laser diffraction reduction or scanning electron microscopy (Fig. 5).

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Fig. 5 Techniques to demonstrate the effectiveness of ultrasound in the reduction of solid food waste

4 Notes: Prospects for Ultrasound Technology Ultrasound technology is undoubtedly an efficient, economical, and easy-to-use tool for the treatment or pretreatment of many food and agricultural wastes that arise in daily life. The various applications of ultrasound technology have had a great boom in the food and pharmaceutical industry, the most studied being food preservation, improvement of organoleptic properties, microbiological preservation methods, and extraction of metabolites with added value. The latter has been investigated with the aim of optimizing the obtainment and utilization of agricultural residues and some food wastes such as fruit, vegetable, and seafood peels. However, the effect of ultrasound on plant and animal cells still needs to be studied, especially if these components are to be used in the food sector or in the health area. In addition, the structural changes of the molecules extracted using UAE must be optimized in order to establish the best conditions for extraction and conservation of the biological properties of the extracted metabolites. The use of ultrasound as a pretreatment in solid food waste degradation processes should still be optimized because the conditions will be defined to the plant or animal material in question and therefore the economic cost during the time required for the reduction of the waste. References 1. UNEP-DTU Partnership and United Nations Environment Programme (2021) Reducing consumer food wastage through digital and digital technologies. Copenhagen and Nairobi 2. Abubakar AR, Haque M (2020) Preparation of medicinal plants: basic extraction and fractionation procedures for experimental purposes. J Pharm Bioallied Sci 12(1):1–10. https://doi. org/10.4103/jpbs.JPBS_175_19 3. Shahidi F, Yeo J (2018) Bioactivities of phenolics by focusing on suppression of chronic diseases: a review. Int J Mol Sci 19(6):1573

4. Jemain SFP, Jamal P, Raus AR, Amid A, Jaswir I (2017) Effects of process conditions on the ultrasonic extraction of phenolics scavenger from Curcuma caesia rhizome. Int Food Res J 24:422–427 5. Kumar K, Srivastav S, Singh-Sharanagat V (2021) Ultrasound assisted extraction (UAE) of bioactive compounds from fruit and vegetable processing by-products: a review. Ultrason Sonochem 70 6. Dalmau E, Rossello´ C, Eim V, Ratti C, Simal S (2020) Ultrasound-assisted aqueous extraction

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Use of Ultrasound Technology for Food Waste Breakdown 23. Gam DH, Hong JW, Kim JH, Kim JW (2021) Skin-whitening and anti-wrinkle effects of bioactive compounds isolated from peanut shell using ultrasound-assisted extraction. Molecules 26(5):1231. https://doi.org/10.3390/ molecules26051231 24. Sharayei P, Azarpazhooh E, Zomorodi S, Einafshar S, Ramaswamy HS (2021) Optimization of ultrasonic-assisted extraction of astaxanthin from green tiger (Penaeus semisulcatus) shrimpshell. Ultrason Sonochem 76. https:// doi.org/10.1016/j.ultsonch.2021 25. Yang K-R, Tsai M-F, Shieh C-J, Arakawa O, Dong C-D, Huang C-Y, Kuo C-H (2021) Ultrasonic-assisted extraction and structural characterization of chondroitin sulfate derived from jumbo squid cartilage. Foods 10:2363. https://doi.org/10.3390/foods10102363 26. Teoh ES (2016) Secondary metabolites of plants. In: Medicinal orchids of Asia. Springer, Cham, pp 59–73. https://doi.org/10.1007/ 978-3-319-24274-3_5 27. Venegas-Casanova EA (2012) Quantification of total flavonoids and tannins present in the aqueous extract of leaves of Thea sinensis L. and its antioxidant capacity. Scientia 4(2) 28. Sharma P, Gaur VK, Sirohi R, Varjani S, Kim SH, Wong JW (2021) Sustainable processing of food waste for production of bio-based products for circular bioeconomy. Bioresour Technol 325. https://doi.org/10.1016/j.biortech. 2021.124684 29. Porter L, Hrstich L, Chan B (1986) The conversion of procyanidins and prodelphinidins to cyaniding and delphinidin. Phytochemistry 25: 223–230 30. Maran JP, Priya B (2016) Multivariate statistical analysis and optimization of ultrasoundassisted extraction of natural pigments from waste red beet stalks. J Food Sci Technol 53(1):792–799. https://doi.org/10.1007/ s13197-015-1988-8 31. Yilmaz T, Tavman S (2015) Ultrasound assisted extraction of polysaccharides from hazelnut skin. Food Sci Technol Int 22. h t t p s : // d o i . o r g / 1 0 . 1 1 7 7 / 1082013215572415 32. Moradi N, Rahimi M, Moeini A, Parsamoghadam MA (2017) Impact of ultrasound on oil yield and content of functional food ingredients at the oil extraction from sunflower. Sep Sci Technol 53. https://doi.org/10.1080/ 01496395.2017.1384016 33. Moradi M, Rahimi M (2019) Effect of ultrasound- and pulsed electric field-assisted enzymatic treatment on the recovery and quality of

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Chapter 8 Integrated Biorefinery Strategy for Orange Juice By-products Valorization: A Sustainable Protocol to Obtain Bioactive Compounds Ana A. Vilas-Boas, Ricardo Go´mez-Garcı´a, De´bora A. Campos, Marta Correia, and Manuela Pintado Abstract Orange juice represents one of the main orange fruit consumption worldwide, and during this process, about 50% of the fresh fruit weight remains as by-products. These by-products comprise mainly peels and pulps, known as pomace, rich in bioactive compounds (BCs) such as essential oils, phenolic compounds (mainly hesperidin), and pectin. These BCs provide various health benefits, like antioxidant, antiinflammatory, anti-microbial, anti-cancerous, cardioprotective, and immuno-modulatory properties, as ingredients for food, nutraceutical, and pharmaceutical industries. Therefore, the use of orange by-products as feedstock in an innovative and sustainable route provides a unique opportunity for highquality value-added ingredients production and is a promising trend to minimize waste deposition. Furthermore, multiproduct recovery can increase the economic viability of recycling by-products and potentially alleviate climate challenges. Therefore, this chapter describes an integrated biorefinery protocol for recovery in a circular economy framework of three different BCs from orange juice by-products: essential oils, hesperidin rich-polyphenol extract, and pectin through the application of sustainable methods with a high yield of extractions. Key words Orange by-products, Circular economy, Hesperidin, Pectin, Essential oils

1

Introduction According to the last FAO statistical bulletin from citrus [1], orange is the major citrus fruit crop produced annually, accounting for a total production of 75.6 M tons in 2021. From this value, almost 30% are used for industrial processing, especially orange juice production. In addition, in Western Europe, orange juice accounts for 60% of all-consumed fruit juices and juice-based drinks [2]. This widespread success is mainly due to orange juice’s organoleptic characteristics, but also to a general consumer’s trend toward healthier eating in which fruit and vegetables play a major

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role, as they are naturally rich sources of minerals, vitamins, dietary fibers, and other health-promoting bioactive compounds (BCs). During the industrial juice squeezing, approximately 50% of the fresh fruit weight is wasted; therefore, around 37.8 M tons of orange by-products were generated only in 2021. The orange juice by-products are composed mainly of peels, pulps, and seeds, and the peels comprise about 36% of the total fruit weight and the pulps 18% [3]. The orange by-products are characterized by high moisture and organic matter content and low pH; therefore, their management is complex because, in the European Union, this kind of waste is legally not disposable in landfills [4]. During the last decades, the major established waste management practices include incineration and anaerobic digestion, while a small part is used as animal feed [5, 6]. The current disposal and poor management of by-products have negatively impacted the environment and social and economic sectors [7]. Furthermore, they are no longer feasible according to the UN’s Sustainable Development Goals (SDGs) to achieve a better and more sustainable future for all. Thereupon, in the last years, the use of orange juice by-products to obtain essential oils (EOs) and, most recently, pectin, is a new sustainable way for companies to solve the waste problem and take more economic profits from the by-products. At the commercial level, pectin is mainly extracted from citrus by-products (mainly from the white part of orange peels (albedo)), and it may be noted that conventional hot acid extraction with inorganic acids from orange peels produced high yields of pectin (20–30% wt) [8]. Nevertheless, these acids have the disadvantage of toxicity and harmful effects on the environment; therefore, recent studies using organic acids (e.g., citric acid) showed a more sustainable approach to maintaining the yield and quality than the conventional method [9]. The use of pectin has been approved by regulatory authorities as a food additive and there is no need to determine the Acceptablele Daily Intake (ADI). The pectin market is projected to grow at 7.6% CAGR between 2019 and 2026 because of its benefits at low quantities rather than the cost, pectin is popular in various scientific fields [10]. Due to its bioactivities, including slow gastric emptying, improvement of physical bowel function, reduced glucose and cholesterol absorption, and increase of fecal mass, it is a good candidate for several purposes such as functional foods and illness treatment, but also for drug delivery components and tissue engineering [11, 12]. Importantly, this biopolymer is resistant to the acidic/alkaline gastrointestinal media as well as the digestive enzymes that favors its application in colon delivery via the oral route under a specific condition [13]. The orange EOs are the most ancestral and famous BCs obtained from the orange peels with several bioactivities exhaustively reviewed, such as anti-inflammatory, anticancer, and antimicrobial activities [14]. EOs represent about 1–3% dry weight of

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orange peels, and the main constituent of these EOs is limonene (90–97%). Limonene has been identified as Generally Recognized As Safe (GRAS) by the Food and Drug Administration (FDA) and is associated with a forwarding trend to be employed in different markets, such as food, cosmetic, and pharmaceutical industries, due to raised concerns related to the less positive health effects associated to other typology of compounds (e.g., synthetic) [15]. Recently, its applicability has been explored for food preservation, antimicrobial packaging, flavoring, and biopesticide. Furthermore, during EO extraction, a co-product is also obtained— hydrolate—that has a very similar, although less intense, odor compared to its corresponding EO [16]. Contrasting EOs, they are water-soluble extracts and can be added to formulas with a high-water content. The characteristics of hydrolates, especially their biological properties, make them widely used in various industries, such as cosmetics and food. Beyond the EOs and pectin, the scientific community also report orange by-products as an excellent source of carotenoids and phenolic compounds with high potential to develop non-synthetic food additives to apply in the food, pharmaceutical, cosmetic, and nutraceutical industry [17]. Phenolic compounds in orange by-products are mainly flavonoids: hesperidin, narirutin [18, 19], and phenolic acids [20]. However, hesperidin is undoubtedly the most important, with a concentration of up to 2% wt [21]. Hesperidin has a broad spectrum of biological activities known, which were also widely reviewed [22]. These biological effects might be important in improving/preventing different pathologies, such as type 2 diabetes, cancer, and neurodegenerative and cardiovascular diseases already reported by in vitro and in vivo studies. Besides that, several human clinical trials showed the hesperidin effects upon antihypertensive, anti-inflammatory, antioxidant, anti-proliferative biomarkers in addition, this flavonoid showed positive effects on decrease cardiovascular and lipid peroxidation biomarkers [23–26]. Furthermore, recent evidence has highlighted hesperidin and other citrus flavonoids as possible microbiota modulator agents, actively inhibiting pathogenic bacteria (e.g., Helicobacter ganmani, Helicobacter hepaticus, and Helicobacter pylori) and selectively stimulating the growth of beneficial bacteria, such as Lactobacillus species [27, 28]. Traditionally, hesperidin has been obtained from different citrus peels throughout alkaline extraction, percolation, or continuous reflux [29]. As expected, this method is time-consuming and requires a high amount of harmful solvents. To overcome the limitations of conventional methods, new sustainable methods focused on obtaining hesperidin with minimal resources is a challenge has gained interest. Besides, currently, the producers of natural extracts must consider not only the production with high yields and quality, the safety for human health, but

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Fig. 1 Schematic protocol of integrated biorefinery strategy for orange juice by-products valorization

also the protection of the environment and preservation of biodiversity. Following this idea, “green extraction” is based on the design of extraction processes with reduced energy consumption, using alternative solvents and renewable natural products, ensuring a safe and high-quality product [30]. Therefore, in the last years, more sustainable methods such as microwave-assisted extraction [31], pressurized liquid extraction [32], and ultrasound-assisted extraction [33] have been used for the isolation of hesperidin from the plant material. As already pointed out and discussed, the correct management of orange juice by-products is a promising trend to minimize waste deposition and treat it appropriately to take advantage of the BCs, resulting in health and environmental benefits. Linked waste recycling with the application of the circular economy models to orange juice by-products is a great opportunity for facing the waste and by-products issues by bringing sustainable processing systems which allow the value chains to be more responsible and sustainable [7]. In a biorefinery concept adopting the circular economy, a framework is possible to introduce the orange by-products generated as feedstock in an innovative and technological route providing a unique opportunity for high-quality value-added ingredients production. Figure 1 shows an integrative approach targeting the total orange by-product valorization to obtain the new potential marketable value-added ingredients, such as EOS, hesperidin-rich polyphenol extract, pectin, and lignocellulosic material. The investment in a multiproduct recovery increases the economic viability of by-product recycling and has the potential to alleviate energy and climate challenge. Furthermore, new addedvalue ingredients are generated to compete in the food, cosmetic,

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and pharmaceutical markets. However, a technological difficulty is caused by the lack of knowledge in the area, associated with a scarcity of literature on practical studies in orange by-products biorefinery. Thus, this chapter aims to provide a novel, industrial, and relevant research line aiming to describe an integral protocol for recovery in a circular economy framework of three different bioactive compounds (BCs) from orange juice by-products: essential oils, phenolic compounds extract, and pectin.

2

Experimental Protocol for Biorefinery Process to Orange Juice By-products

2.1

Material

Distillated water, citric acid, ethanol 96% (v/v), and ethanol 70% (v/v).

2.2

Raw Material

The integrated approach in this protocol could be done with orange peels or with orange pomace. The methodology could be applied in the fresh raw matter to avoid the energy consumption of the drying process since during the entire process the matrix will always be in contact with water. Due to the biological nature of the orange juice by-products, the process must be carried out with a representative amount of sample (>10 kg) or in three different batches with more than 5 kg each. Furthermore, before start the integrative process it is important to evaluate the dry matter content as well as the nutritional composition of orange juice by-products according to the AOAC methods. The waste should be used directly in the first extraction (steam distillation) without any pretreatment to avoid the EOs loss; however, after this extraction, the sample must be grounded (Fig. 2) to allow a correct homogenization in the following extractions. According to European Pharmacopoeia, orange EOs should be obtained only by steam distillation or cold pressing. Despite

Fig. 2 The fresh orange waste after the EOs extraction (left side image) was pre-treated (ground in thermomixer) (right side image) before continuing the integrated biorefinery process

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Fig. 3 Schematic representation of steam distillation apparatus for EOs and hydrolate extraction used for orange by-products 2.3 Essential Oils Extraction by Steam Distillation

steam distillation being the most traditional method used for orange essential oil extraction, in the last years, it has been replaced by cold pressing technique due to the high energy consumption of the first process [34]. However, several justifications can be found in the literature for choosing steam distillation instead of coldpressed. While the scent of cold-pressed citrus EOs usually stays true to the scent of the peel/fruit, the oils produced via this method also tend to contain more phototoxic constituents. Phototoxicity causes the skin to be extra sensitive to sunlight, causing inflammation, blistering, reddening, and skin pigmentation. Because of this, it is not recommended to use cold-pressed oils topically on your skin. Instead, cold-pressed EOs should be used exclusively for inhalation and wash-off products. Another added advantage is that it has a longer shelf life too; as cold-pressed essential oils tend to oxidize faster, the limonene content is higher than cold-pressed methods, and the purity is higher because the cold-pressing method isolates non-aroma-active fats [35, 36]. To perform the EOs extraction by steam distillation, it is necessary to couple the distillation reactor to a boiler and a condenser (Fig. 3). While the boiler water heats up to its boiling temperature (100 °C), the raw material is placed in the distillation reactor. The reactor must not contain more than 3/4 of its capacity. The condenser must be full of cold water or with anti-freeze liquid to quickly condense the vapor oils and waters produced inside of the distillation reactor. The extraction should be carried out for approximately 45 min. However, if it is visible in the separating funnel that no more EO is being extracted, the process must be finished to avoid the thermal degradation of orange juice by-products biomass. At the end of this process, two products will be obtained: (i) hydrolate (aromatic waters) and (ii) orange EOs. The EO’s yield should be around 1–2% while large volumes of hydrolate will be collected.

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For these two added-value products obtained, the volatile characterization could be performed by gas chromatography-mass spectrometry according to the method described by Alves-Silva et al. [37], for example. However, according the European Pharmacopoeia, the sweet orange EOs should be almost exclusively D-limonene (> 97%). 2.4 Hesperidin-Rich Polyphenol Extract by Microwave Hydrodiffusion and Gravity (MHG)

Microwave Hydrodiffusion and Gravity (MHG) is an original combination of microwave heating and gravity working at atmospheric pressure with the potential for many extractive applications [38]. This technique represents an efficient, economical, and clean alternative since it requires low energy and little or no solvents. There are different models of MHG at laboratory and semiindustrial scales, all of which belong to Milestone [38]. The equipment is structured as an oven with a cavity at the bottom that allows the extract to flow by gravity through a condenser connected to a thermostatic bath (10 °C) that allows the collected extract to be cooled [39]. ETHOS X equipment is the most up-to-date model that can be used to extract pigments and flavonoids; therefore, this model should be selected to perform this protocol.

Fig. 4 Schematic representation of microwave hydrodiffusion and gravity technology to extract hesperidin-rich polyphenol extract from orange waste

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In this relatively simple method, the orange juice by-products (after the steam distillation process) are grounded and mixed with water (ratio 1:5 (m/v) and directly placed in a microwave reactor (Fig. 4). The use of water as a solvent is considered a green extraction, as it is a safe, non-toxic, non-corrosive, and non-flammable solvent. Afterwards, the equipment was programmed to operate for 20 min with a power ratio of 1.75 W/g of sample. According to Zill-e-Huma [40], powers higher than 2.5 W/g could destroy phenolic compounds and powers lower than 1.5 w/g are inefficient for the extraction of flavonoids (such as hesperidin and narirutin). However, the operator should validate that the extraction process must not exceed 85 °C to avoid the degradation of BCs and the burn of sample. The extraction process begins with an induction time of 3 up to 5 min (without any extract recovery); it ends with the appearance of the first colored drop of the extract. After, a constant increasing in the extraction flow rate occurs, ending with the appearance of the first colorless drop of the extract (20 min). The internal heating generated due to the microwaves interaction with polar molecules (water) allows the biological matrix rupture triggers the extravasation of cellular content (phenolic compounds). Heating under microwaves thus frees molecules of interest together with in situ water. This physical phenomenon, known as hydrodiffusion, allows the extract to diffuse outside the raw material and drop by earth gravity out of the microwave reactor (a heat exchanger outside the microwave oven cools the extract continuously). At the end of time extraction, the liquid extract (hesperidin rich-polyphenol extract) is collected and freeze-dried with an adjuvant (2% (m/v) maltodextrin should be added before the freezing). The remaining solid biomass is collected and saved for the following extraction process. 2.5 Pectin Extraction by Sustainable Hot Acid Extraction

After the extraction with MHG, the remaining solid residue was used to obtain pectin. The extraction was performed using commercial citric acid according to the flow process chart presented in Fig. 5. The major parameters for the protocol were: mass-solvent ratio of 1:50, pH 2.0, and extraction at 95 °C during 90 min with agitation (300 rpm). Briefly, 2/3 of the final water volume should be heated until 80 °C and adjust the pH with citric acid at 6 M until 3.0. After that, the sample was added, the pH was corrected to 2.0, and the remaining water volume was added. After the extraction, the sample was filtered using two layers of muslin cloth, and the filtrate was centrifuged at 15,000g for 15 min to remove insoluble particles from the pectin slurry. At this point, the lignocellulose fraction was collected and dried in vacuum oven at 60 °C for 6 h. Then, pectin precipitation was conducted by mixing the supernatant liquid with ethanol 96% (v/v) in a ratio of 1:2 (v/v) and incubated at 4 °C overnight (16–18 h). After that, the pectin was collected by filtration with four layers of muslin cloth and washed

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Fig. 5 Flow diagram of the process to extract pectin from orange juice by-products through hot acid treatment with citric acid

two times with ethanol 96% (v/v) and ethanol 70% (v/v). Finally, the precipitated pectins were freeze-dried and ground. The lignocellulosic fraction should be dried in a convective oven at 50 °C for 12 h.

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Conclusions Unquestionably, as in the past, even now, and future, sustainable solutions for the management of food by-products for their reincorporation in the industrial chains are the principal challenges in the industrial field. The unsustainable management of orange juice by-products is a risk to the environment and a loss of money to the companies; therefore, the integrated biorefinery protocol proposed in this chapter provides a unique circular concept approach to produce a wide range of new value-added bioactive ingredients to be put back into the supply chain, allowing economic growth from environmental losses. Furthermore, the bioactive ingredients obtained have a growing market demand and have several applications in food, cosmetic, pharmaceutical, and nutraceutical industries. Furthermore, all the methods proposed in this work are environmentally friendly, rapid to scale up, and respect the human food security.

Acknowledgments The authors would like to thank the scientific support by MEDISMART project-Mediterranean Citrus: innovative soft processing solutions for SMART (sustainable, Mediterranean, agronomically evolved, nutritionally enriched, traditional) products through national funds from FCT-Fundac¸˜ao para a Cieˆncia e Tecnologia (reference: PRIMA/0014/2019) and by CBQF-Centre of Biotechnology and Fine Chemistry from Escola Superior de Biotecnologia at Universidade Cato´lica Portuguesa under the FCT project UIDB/Multi/50016/2020. In addition, the author Ana. A. VilasBoas would like to acknowledge FCT for the individual PhD grant (2020.05655.BD) and De´bora Campos for the individual postdoctoral contract (2021.01947.CEECIND). References 1. FAO (2021) Citrus fruit statistical Compendium in Rome. FAO, Rome 2. Galaverna G, Dall’Asta C (2014) Production processes of orange juice and effects on antioxidant components. In: Processing and impact on antioxidants in beverages. Academic Press, Oxford, pp 203–214. https://doi.org/10. 1016/B978-0-12-404738-9.00021-0 3. Siddiqui SA, Pahmeyer MJ, Assadpour E, Jafari SM (2022) Extraction and purification of D-limonene from orange peel wastes: recent advances. Ind Crop Prod 177:114484.

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Chapter 9 Energy Integration of the Hydrothermal Pretreatment of Food Waste in Terms of a Sustainable Biorefinery Iosvany Lo´pez-Sandin, Rosa M. Rodrı´guez-Jasso, and He´ctor A. Ruiz Abstract The application of the biorefinery concept has contributed to the reduction of the volumes of food waste (FW), considered an economic and environmental problem. The reuse of these residues has made it possible to give them added value and take advantage of them in the production of biofuels and other value-added products. However, during the FW conversion process, there are limitations that can hinder its correct development and that can be solved through energy integration and different bioprocesses. In this sense, hydrothermal pretreatment stands out in FW processing. Therefore, the aim of the chapter is to propose a methodology for the energy integration of hydrothermal pretreatment in food waste biorefinery terms. Key words Bioenergy, Bioprocesses, Efficiency, Environment, Value-added products

1

Introduction Energy is a critical factor that controls the socioeconomic development of a country and is obtained mainly from fossil fuels, a nonrenewable energy source that has been depleted with its continuous use and the passage of time. Therefore, reducing dependence on these fuels has intensified the search for renewable energy sources such as biofuels produced from the biorefinery of food, agro-industrial, and other waste [1, 2]. Food waste (FW) has little use, and considerable volumes are generated annually that exceed the capacity and availability of landfills, representing an economic, environmental, and health threat [3, 4]. In addition, inadequate traditional methods are used for the management of FW, which do not comply with future energy and environmental standards [5]. However, from a biorefinery approach and due to the FWs’ physicochemical and biological nature, they can be used in the production of biofuels, chemicals, and biobased materials [2, 6]. The composition, the interaction between its components, and the desired final products are

Cristobal N. Aguilar Gonzalez et al. (eds.), Food Waste Conversion, Methods and Protocols in Food Science, https://doi.org/10.1007/978-1-0716-3303-8_9, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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fundamental in the valorization of the FW and in the efficiency of the selected biorefinery process [5]. For raw materials that present recalcitrant structures such as lignocellulosic materials, a prior pretreatment is necessary to break this structure and mainly facilitate the cellulose enzymatic degradation [7]. The profitability of the biorefinery process depends, among other factors, on the total integration of the production chain and energy integration, minimizing energy costs, operating costs, and environmental impact, in addition to maximizing production, quality, and social benefits [8, 9]. For the total integration of the chain, tools that optimize processes are necessary [8], among which we can mention are the following: (I) life cycle analysis (LCA), an effective tool to analyze material inventories, energy, and environmental impacts throughout the life cycle of a product or process [10], (II) systematic methodologies that use a superstructure-based process synthesis approach to determine potential growth pathways [11], and (III) the data envelopment analysis (DEA) method, widely used to study energy efficiency as a method of evaluating the total efficiency of factors [12]. Energy integration based on the implementation of tools or methodologies that allow the control and monitoring of energy use is an alternative to reduce costs and increase energy efficiency and the productivity of a process. Therefore, the chapter aim is to propose an energy integration method for hydrothermal pretreatment in terms of food waste biorefinery. Figure 1 shows the energy integration scheme of hydrothermal pretreatment in terms of food waste biorefinery.

2 2.1

Food Waste and Bioenergy Production Food Waste

FWs include both the edible and inedible parts of food that are generated throughout the entire food supply chain. It includes everything from spoiled food; fruit and vegetable waste; agricultural crop residues; waste left on plates from hotels, homes, and restaurants; and any other food that is wasted at any stage of the supply chains. Among the many factors that have led to the increase in FW generated during industrial, agricultural, or domestic processes, urban growth and lifestyle changes can be mentioned. This has led to a third of the total food produced being wasted each year [13, 14], the equivalent of approximately 1.3 billion tons of food worldwide [4, 14]. Therefore, completely avoiding food waste is practically impossible. However, it is possible to reduce the volume or quantity of food waste [2]. The main FWs composition is a mixture of carbohydrates (cellulose, starch, hemicellulose), lipids, proteins, and trace amounts of inorganic compounds [3, 14, 15], although its composition may vary depending on the type of food waste or production source, as

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Fig. 1 Energy integration scheme of hydrothermal pretreatment in terms of food waste (FW) biorefinery. Energy input (red) and output (green) in the process

well as its components and generation stage [3, 14]. FWs are generated throughout the food life cycle, from production to consumption, and can be divided into five main stages: (I) production stage, (II) handling and storage stage, (III) processing and packaging stage, (IV) distribution and commercialization stage, and (V) consumption stage. At all stages, a significant amount of FW is generated that decomposes and becomes an environmental problem due to the increase in greenhouse gas (GHG) emissions, bad odors, and the natural resource contamination (water, land, air), generally affecting human health. The organic composition and moisture of the FW facilitate the appearance of a variety of

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pathogens that reproduce in unpretreated food waste, which represents economic losses and the cause of various diseases due to the reproduction of pathogens [16–18]. In addition, the volumes of losses and FW throughout the supply chain are alarming figures that have drawn the attention of all sectors involved. This situation not only reduces the amount of food available but also the availability of many natural resources needed to produce food. This will become a critical and very important factor when the world population increases by more than 30% in the year 2050 [16], generating an urgent need to apply preventive measures and standard management practices to reduce the FW load [3]. In countries like the United States (USA), food and agriculture consume up to 16% of the energy, almost half of the land, and 67% of the use of fresh water in the country. This endangers the sustainability of the ecosystem with the depletion of natural resources [16]. 2.2 Bioenergy Production

The production of the bioenergy depends on the three main organic components of FW (lipids, proteins, carbohydrates), which can be biologically converted into some types of bioenergy through various processes: (I) fermentation of carbohydrates to obtain bioethanol or biobutanol and biohydrogen (dark fermentation); (II) transesterification of lipids (oils and fats) to produce biodiesel; (III) digestion and anaerobic conversion of carbohydrates, lipids, and proteins to produce biomethane or biogas; IV) pyrolysis and gasification; (V) hydrothermal carbonization; and (VI) incineration [15, 19]. In general, each component of FW favors the production of a certain product, and in some processes, they can have an inhibitory effect, for example, lipids are inhibitors to produce bioethanol, biomethane, and biohydrogen [15]. In addition, a high content of proteins and lipids can generate higher levels of ammonia and sulfur that result in an inhibitory effect on the metabolism of anaerobic consortia and thus limit one of the most used techniques due to its economic and environmental impact, the production of biogas [20]. Therefore, the composition of the FW is decisive in the biodegradation and transformation of this raw material into bioenergy and other products with high added value, which leads to the search for more efficient strategies or processes for its processing. Pereira et al. [21], based on the chemical composition of FW from a restaurant, estimated its potential for energy production and concluded that some carbohydrate-rich wastes have potential application in biogas production and others rich in lignin for energy cogeneration. Likewise, some bioprocesses such as acidogenesis, methanogenesis, solventogenesis, bioelectrogenesis, photosynthesis, fermentation, and oil production can be used to valorize FW from the production of biofuels, platform chemicals (pigments, enzymes, organic acids, essential oils), bioelectricity, biomaterials, biofertilizers, animal feed, etc. Likewise, the integration of these

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bioprocesses can be an alternative to increase their efficiency and sustainability. Furthermore, the application of the integrated biorefinery concept is a strategy that can lead to the development of the circular bioeconomy [22–24].

3

Biorefinery Concept: Food Waste Biorefinery Biorefinery

The concept of biorefinery means or can be described as a processing unit or a facility that uses biomass optimally to produce a wide range of products and has among its principles to be self-sufficient and environmentally friendly [25]. It can also be described as the sustainable bioprocessing of biomass to obtain a wide range of marketable products (food, feed, materials, chemicals, etc.) and energy (fuels, power, heat) [14]. The biorefinery integrates different conversion processes such as biochemistry, thermochemistry, combustion, and microbial growth platform to efficiently produce sustainable biobased products (bioenergy, biochemicals, biofuels, etc.) [25].

3.2 Food Waste Biorefinery

FW biorefinery is a process by which a broader range of food waste is converted into biofuels, platform chemicals, and biobased materials [5]. FW biorefineries have as their main objective the sustainable food production and the generation of low-carbon energy [26]. The biorefinery allows to value FW and minimize environmental challenges through its efficient use. The multiple bioproducts obtained through the integration of relevant techniques contribute to minimizing dependence on fossil fuels and moving toward the circular bioeconomy. Among the many products that can be produced simultaneously and sequentially, biofuels (biogas, biomethane, bioethanol, and biodiesel) stand out [2, 27, 28]. FW biorefinery is broadly classified into three main groups: (I) biological pathway, food waste is converted into value-added products using enzymes or microorganisms; (II) thermochemical process, FW is treated at high temperature using chemical products as solvent (liquefaction, pyrolysis, and gasification); and (III) chemical process, chemical products are used as solvent and catalyst in the recovery of FW. The integrated combination of two or more of these processes has shown higher conversion efficiency [2]. Besides, higher profitability can be obtained in a biorefinery installation with different raw materials [29].

3.1

4

Hydrothermal Pretreatment and Energy Integration

4.1 Hydrothermal Pretreatment

Currently, the biorefinery uses pretreatment techniques that ensure optimal use of biomass. Hydrothermal pretreatment is a clean technology that is advancing with promising results, in the fractionation of lignocellulosic biomass, with potential application on an

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Fig. 2 Scale reactor for hydrothermal pretreatment for biomass valorization

industrial scale in second-generation biorefinery and circular bioeconomy. This technology does not require chemical inputs other than liquid water or steam and heat, which can be conducted at elevated temperatures (150–240 °C) and pressures (approx. 1–3.5 MPa), for short periods of time (minutes) or extended periods of time (hours) [7, 30, 31]. Hydrothermal pretreatment, also known as liquid hot water, hydrothermolysis, subcritical water treatment, and autohydrolysis, converts organic feedstock into biofuel and high value-added products through the action of water at elevated temperatures in a sealed vessel or reactor (Fig. 2) [7, 32]. The process begins with the macromolecules of the biomass breaking down into their essential components (by hydrolysis). Then these are rebuilt to form new products, the same as saying that a sealed container with water at an

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elevated temperature induces the hydrolysis of complex biomass compounds into subunits such as glucose. Therefore, the elevated temperature reached by water can dissolve hydrophobic organic compounds such as free fatty acids, which are not soluble in water at room temperature [32]. Apart from temperature, the operating conditions of hydrothermal pretreatments can be varied through different parameters such as residence time, particle size, proportion of water, and solid biomass, among others [7]. In this sense, the severity factor [log (Ro)] is an index that provides a relationship between temperature and reaction time [31]. This can be used to visualize the effect of the experimental conditions on the result obtained [33], and it is expressed by the following equation: logðRoÞ = log t∙ exp ðT - T ref Þ=ω

ð1Þ

where t is the reaction time (min or h), Tref is the reference temperature (often 100 °C), T denotes the experimental temperature (°C), and ω represents the activation energy that uses a value of 14.75, which is related to the energy required for the hemicellulose solubilization [33, 34]. Depending on the temperature and pressure, hydrothermal pretreatment is classified into two types of processes: subcritical (below the critical point of water, i.e., 373 °C and 22.1 MPa) and supercritical that exceeds this point. Hydrothermal pretreatments is characterized by being cost-effective (e.g., avoiding costly or heavyduty materials of construction, catalysts, reagents, or neutralization steps), low energy (e.g., using feedstocks that do not require size reduction or other process), and easy to include in a process integration and intensification scheme [7]. The heterogeneous composition, high moisture content, and low calorific value of FWs currently make it a challenge for conversion into bioenergy and an impediment to the development and scaling of efficient and robust industrial processes [35]. However, pretreatment techniques have made it possible to significantly increase the calorific power of FW. For example, by increasing the temperature and torrefaction time of FW, the values of calorific power can increase, reaching values close to those of coal [36]. Also, the previous pretreatment with enzymes can increase the values of the calorific power of the hydrocarbon [37]. Likewise, FWs have been mixed with garden waste, and pretreatment (220 °C for 1 h) led to an increase in calorific value [38]. This same mixture has been used during solid-state anaerobic digestion to produce bioenergy. However, due to the recalcitrant nature of the yard waste, these must be thermally pretreated and subsequently mixed with FW to balance the nutrients for anaerobic co-digestion, which results in higher methane production and net energy gain [39]. The conditions that the water reaches during the hydrothermal pretreatment (temperature, pressure, time) allow the elimination of

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unwanted organisms or microbes present in the FW [32], being a potential alternative for the sterilization of FW with high moisture content [40]. Additionally, it solubilizes organic solids, resulting in higher recovery of fermentable sugars for bioenergy production [14]. However, a limitation of hydrothermal pretreatment is that it requires a large capital investment in equipment and infrastructure, which makes the process only techno-economically viable on a large scale, currently limiting its commercial expansion [32]. 4.2 Energy Integration

Biorefinery processes for bioenergy production require a large amount of energy, so the selection of raw material processing options and management decisions can be decisive in the results in energy and environmental terms [41]. Therefore, various technologies have been studied to increase their efficiency and reduce environmental impact, such as integrating different bioprocesses to generate a wider range of by-products and convert FW into higherquality products with effective recovery of bioenergy and biomaterials [42]. In this sense, the biochemical biorefinery has been used for the production of bioenergy by requires less energy than processes that include thermochemical transformations [4], as well as the use of hydrothermal pretreatment techniques in FW processing to leach solid animal fat that is mixed with other solid organic compounds and makes its direct extraction difficult [43], as occurs in the stages of high consumption of energy (lipid extraction and purification) during biodiesel production to leach solid lipids (fats) and remove the water before transesterification [15]. The hydrothermal pretreatment has improved the solubilization of the FW, managing to increase the energy conversion efficiency (78.6%) with respect to the untreated FW by 31.7% [44]. On the other hand, the anaerobic digestion from an energy point of view has shown to be a viable option for the treatment of FW, since it manages to recover a large part of the energy (81%) stored in the raw material [45]. However, considering the advantages and disadvantages of each bioprocess individually, the integration of several bioprocesses in a biorefinery platform may be the best option to maximize the potential of FW to produce bioenergy and high value-added products [46–48]. The individual application of a bioprocess limits the use of organic components, with part of these remaining by selective action as a certain amount of undesirable product and unstable residue [15]. Oliveira et al. [9] evaluated different biorefinery scenarios, and energy integration demonstrated a reduction in energy consumption of more than 50% compared to scenarios without integration. In this sense, Lin et al. [49] mentioned that a significant increase in efficiency (56.9%) can be achieved with the recovery of the heat generated during pretreatment. Likewise, the reduction of energy consumption and the emission of residual gases may be associated with the use of reactors that provide a controllable and stable

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biological environment [50]. In general, the energy integration of various bioprocesses can contribute to an efficient biorefinery that maximizes bioenergy production and minimizes the volume of FW [15].

5 5.1

Energy Balance Energy Efficiency

The energy balance comprises and applies different methodologies, indices, and energy terms, among which energy efficiency (EE) stands out [51]. EE can be defined as the energy required (input) to produce a quantity of product (output), always emphasizing producing more with less [52]. Also, it can be defined as the rational and efficient use of energy to obtain goods or services with the use of fewer energy resources, without affecting comfort levels, quality of life, or productivity. Thus, EE aims to reduce the amount of energy used in production systems and services, in addition to contributing to mitigating carbon emissions [53]. Therefore, policies that promote energy efficiency are an obvious response to climate change [54]. On the other hand, total factor energy efficiency (TFEE) is a widely accepted energy efficiency method. This, unlike traditional methods, considers that the energy input must combine with other factors such as labor and capital to obtain more tangible results or generate economic results [55]. TFEE is the ratio between the target power input and the actual power input, as shown below: 0≥

Target Energy Input ≤1 Actual Energy Input

ð2Þ

Therefore, TFEE can be considered an effective solution to the shortcomings of the traditional single-factor EE assessment [12]. Energy studies have it classified into three large groups: (I) energy analysis, (II) energy evaluation, and (III) methods of energy saving measures. Group I aims to increase the transparency of energy consumption with the systematic investigation and identification of the different energy consumers of a production system or process. This allows identifying processes, areas, or equipment with high energy consumption, representing a significant step to improve the EE of the production process. While group II aims to establish methodologies, tools, and evaluation indices that allow deepening into energy consumption within a production process and contribute to EE, in contrast, group III aims to identify corrective actions by selecting of the best technology to improve EE and environmental sustainability and proposing methods and tools that characterize EE measurements [56].

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5.2 Input and Output Energy

A crucial step for the success of the energy evaluation of the FW biorefinery process is to define the energy inputs and outputs. In this sense, there are diverse studies on EE, although a single standard has been defined to select the input and output variables [12]. However, the energy inputs and outputs will depend on the type of process and its scope. For example, [51], during the energy evaluation of the production of an agricultural crop, they considered the labor and the inputs necessary to conduct the process (electricity, diesel, machinery, chemical products, etc.) as energy input and the energy obtained from the biomass as energy output. Panepinto et al. [57] evaluated the EE of a wastewater plant and considered electricity as the input energy and the services generated as the output energy (equivalent population, volume of treated water, decrease in the chemical demand for oxygen and total nitrogen).

5.3 Calculation Procedure

In the energy balance, different methodologies have been used that combine energy inputs and outputs. However, the calculation procedure will adjust to the type of process and the energy source used to carry it out [51, 58, 59]. For the quantification of energy, different units of measurement are used, such as joules, kWh, or million tons of oil equivalent (Mtoe) [53], and the information for the balance must be compiled from the energy demand by the equipment, facilities, supplies, and labor necessary to execute each stage of the process, considering times, operating conditions, quantities, and the respective energy equivalent, in addition to the calorific value of the products generated [51, 59–62]. The energy equivalents and the calorific value can be determined or assumed from what has been reported in the literature (see Table 1) if they adjust to the process in execution. The energy balance of the hydrothermal pretreatment and other stages of the FW bioprocess can follow the calculation procedure described below.

5.3.1

Energy input (Ein) in MJ can be determined by the following equation:

Input Energy

E in =

E ER þ E HL þ E CR þ E EO

ð3Þ

where EER is the energy consumed (MJ) in electricity (MJ), EHL is the energy consumed (MJ) by human labor, ECR is the energy consumed (MJ) in chemical reactants, and EEO is the energy consumed (MJ) by other inputs such as water, enzymes, yeasts, etc. The forms of energy that are not required during the process take a value of 0. Electricity consumption considers electrical devices directly used in the FW biorefinery process, as well as associated equipment and facilities. Consumption data can be record by an electric meter

kg kg kg kg kg kW·h

Food Waste

Sulfuric acid

Sodium hydroxide

Other chemicals

Water

Electricity

kg kg kg

Phosphate (P2O5)

Potassium (K2O)

Calcium (CaO)

kg kg kg

Herbicides and growth regulators

Fungicides

Insecticides

2. Pesticides

kg

Nitrogen (N)

1. Mineral fertilizers

A. Production of a crop for food

L

Diesel

h kg

Manual Mechanized

Unit

Machinery

Human labor

Energy input

Concept

Table 1 Energy equivalents (energy input and output)

-1

278.00

92.50

238.70

2.12

11.15

15.80

77.50

11.93

0.005

8.74

10.41

0.702

5.35

(continued)

[66]

[66]

[66]

[66]

[66]

[66]

[51]

[51]

[65]

[64]

[64]

[63]

[23]

[51]

[51]

138.00a 47.80

[51] [51]

Source

1.96 1.05

(MJ·unit-1)

Energy Integration of the Hydrothermal Pretreatment of Food Waste in Terms. . . 135

kg kg

Formulation

3. Seed grain

ha ha ha

Heavy cultivator

Mulching machine

Rotary-harrow seeder

ha

Pesticide application run

Baling and carting (15 km)

8. Straw collection (collecting ratio 90%)

Combine and transport (15 km)

7. Grain harvest

Transport and on-field distribution

T of straw dry weight

t of grain dry weight

m-3

ha

Nitrogen fertilizer application

6. Stillage application

ha

Basic fertilizer application

5. Fertilizer and pesticide application

ha

Plow and cultipacker

4. Tillage and sowing

Unit

Concept

Table 1 (continued)

250.00

167.00

20.92

65.00

50.00

33.00

371.00

230.00

270.00

591.00

2.50

62.10

(MJ·unit-1)

[66]

[66]

[66]

[66]

[66]

[66]

[66]

[66]

[66]

[66]

[66]

[66]

Source

136 Iosvany Lo´pez-Sandin et al.

L of bioethanol (including a stillage recycling rate of 50%) L of bioethanol L of bioethanol

2. Mash process

3. Fermentation and yeast propagation

4. Distillation and dehydration

kg kg kg kg kg kg kg kg

1. Bioethanol

2. Bioethanol 10%

3. Bioethanol 20%

4. Bioethanol 30%

5. Bioethanol 5% and biodiesel 15%

6. Cellulose

7. Hemicellulose

8. Lignin

25.00

16.81

17.28

41.00

33.00

36.00

40.00

[69]

[68]

[68]

[67]

[67]

[67]

[67]

[65, 67]

[66]

Varying

(dependent on grain N, P2O5, K2O, CaO content, according to mineral fertilizer equivalents)

3. Stillage

26.80

[66]

17.30

kg of dry weight

2. Straw

[66]

[66]

[66]

[66]

[66]

[66]

L

21.20

4.74

0.06

0.40

75.00

3.80

1. Bioethanol

Includes the energy cost in steel production plus 50% for assembly

a

kg (if powder) or L (if liquid)

Technical enzyme preparations

Energy output

kg

Brewing malt (malting process incl. mean brewing barley production)

1. Saccharification substances

B. Bioethanol production

Energy Integration of the Hydrothermal Pretreatment of Food Waste in Terms. . . 137

138

Iosvany Lo´pez-Sandin et al.

with a frequency of 1 h. For the equipment(s) and/or installation (s) used for limited periods of time, they can be measured during their period of use. A general equation to calculate the energy consumed in electricity (EER) can be expressed as follows: E ER =

i=n i =1

CE  Q E

ð4Þ

where CE is the electricity consumption (kWh) by the equipment (s) and/or the electrical installation (s) and QE is the energy equivalent of electricity (MJ·kWh-1). The energy consumed in human labor (EHL) can be determined by the following equation: E HL =

i=n i=1

TW  QH

ð5Þ

where TW is the working time (h) of the operator(s) per unit of production and QH is the energy equivalent of human labor (MJ·h-1). The energy consumed in chemical reagents (ECR) can be determined by the following equation: E CR =

i=n

RC  Q R

i=1

ð6Þ

where RC is the chemical reagent(s) consumed (kg) and QC is the energy equivalent of the chemical reactant (MJ·kg-1). The energy consumed in other inputs (EEO) can be determined using the following equation: E EO =

i=n i=1

OS  Q S

ð7Þ

where OS is the input(s) consume (kg) and QS is the energy equivalent of the input (MJ·kg-1). 5.3.2

Output Energy

The energy output (Eout) can be determined using the following equation: E out =

i=n

P out  C V

ð8Þ

i=1

where Pout is the output product(s) (kg) during the FW biorefinery process as biofuels, biomaterials, chemicals, and other bioproducts and CV is the calorific value of the output product(s) (MJ·kg-1) and can be obtained using a bomb calorimeter or by searching the literature for a value that fits the running process conditions (Table 1).

Energy Integration of the Hydrothermal Pretreatment of Food Waste in Terms. . . 5.3.3 Energy Efficiency and Other Indices for the Process Energy Balance

139

Energy efficiency (EE) is the energy (MJ·kg-1) provided by the output product(s) of the biorefinery process divided by the input or required energy (MJ·kg-1) for your production. It is used when said process is for power generation and can be calculated using the following: EE =

E out E in

ð9Þ

Energy productivity (EP) is the relationship between the amount or weight of product(s) obtained, measured in units of mass (kg), and the energy input or required (MJ) to obtain them and can be calculated using the following equation: EP =

BW E in

ð10Þ

where BW is the processed biomass (kg). Specific energy (SE) is the ratio between the energy output or provided (MJ) by the product(s) obtained in the biorefinery process and the number of product(s) measured in units of mass (kg) and can be calculated using the following equation: SE =

E out BW

ð11Þ

Net energy gain (NEG) the difference between the energy output or produced (MJ·kg-1) by the product(s) obtained in the biorefinery process and the energy input or required (MJ·kg-1 to produce it(them) and can be calculated using the following equation: NEG = E out - E in

6

ð12Þ

Conclusions The need to reuse food waste has led to the search for new processing methods and technologies, as well as to increase its efficiency. In this sense, the biorefinery appears as a viable and efficient alternative to convert food waste into biofuels and other value-added bioproducts. However, during the performance of these bioprocesses, limitations arise that can hinder their correct development and that can be solved through energy and process integration. Thus, hydrothermal pretreatment stands out as an efficient alternative that can combine with other technologies, through indices such as energy efficiency, net energy gain, and others, that allow quality and efficiency to be increased and the emission of pollutants to be reduced at the atmosphere.

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Acknowledgments The authors would like to thank to the Autonomous University of Coahuila for the support provided to the Biorefinery Group and CONACYT by the scholarship 1047495 granted in the 2021 postdoctoral announcement stays in Mexico—Academic Modality. References 1. Kumar B, Bhardwaj N, Agrawal K, Chaturvedi V, Verma P (2020) Current perspective on pretreatment technologies using lignocellulosic biomass: An emerging biorefinery concept. Fuel Process Technol 199: 106244. https://doi.org/10.1016/j.fuproc. 2019.106244 2. Tsegaye B, Jaiswal S, Jaiswal AK (2021) Food waste biorefinery: pathway towards circular bioeconomy. Foods 10(6):1174. https://doi. org/10.3390/foods10061174 3. Paritosh K, Kushwaha SK, Yadav M, Pareek N, Chawade A, Vivekanand V (2017) Food waste to energy: an overview of sustainable approaches for food waste management and nutrient recycling. BioMed Res Int. https:// doi.org/10.1155/2017/2370927 4. Ebikade E, Athaley A, Fisher B, Yang K, Wu C, Ierapetritou MG, Vlachos DG (2020) The future is garbage: repurposing of food waste to an integrated biorefinery. ACS Sustainable Chem Eng 8(22):8124–8136. https://doi. org/10.1021/acssuschemeng.9b07479 5. Carmona-Cabello M, Garcia IL, Leiva-CandiaD, Dorado MP (2018) Valorization of food waste based on its composition through the concept of biorefinery. Curr Opin Green Sustainable Chem 14:67–79. https://doi.org/10. 1016/j.cogsc.2018.06.011 6. Karthikeyan OP, Mehariya S, Wong JWC (2017) Bio-refining of food waste for fuel and value products. Energy Procedia 136:14–21. https://doi.org/10.1016/j.egypro.2017. 10.253 7. Ruiz HA, Conrad M, Sun SN, Sanchez A, Rocha GJ, Romanı´ A, Castro E, TorresRosa A, Rodrı´guez-Jasso M, Andrade LP, Smirnova I, Run-Cang S, Meyer AS (2020) Engineering aspects of hydrothermal pretreatment: From batch to continuous operation, scale-up and pilot reactor under biorefinery concept. Bioresour Technol 299:122685. https://doi.org/10.1016/j.biortech.2019. 122685 8. Budzianowski WM, Postawa K (2016) Total chain integration of sustainable biorefinery

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Part III Food Waste as a Carbon Source for Fungi Based Processes: Bioactives Obtention and Releasement

Chapter 10 Solid-State Fermentation as Strategy for Food Waste Transformation Israel Bautista-Herna´ndez, Mo´nica L. Cha´vez-Gonza´lez, Arturo Siller Sa´nchez, Karen N. Ramı´rez Guzma´n, Cristian Torres Leo´n, Pedro Aguilar Za´rate, Cristobal N. Aguilar Gonzalez, and Deepak Kumar Verma Abstract An excessive amount of food waste is generated daily globally, which has a rich composition of diverse compounds, making its use desirable. There are several strategies to take advantage of food waste; among the most promising is solid-state fermentation. Solid-state fermentation is a tool that allows the development of microorganisms on solid or semisolid substrates in the absence of free water. This particularity allows agro-food waste to be used as fermentation substrates to generate various products such as the release of bioactive compounds, enzyme production, waste detoxification, and others. This chapter describes solid-state fermentation as a valuable tool for utilizing food waste and the characteristics that agro-industrial waste must possess to be considered suitable fermentation substrates. The protocol for this strategy is established, specifying the minor details surrounding the bioprocess. In addition, particular examples of success on various food wastes are described. Key words Solid-state fermentation, Waste management, Bioprocess, Bioactive compounds, SSF protocol

1

Introduction In the last century, the human population has been growing in exponential rate, and the prediction englobes a global human population from 7.7 to 9.7 billion in 2050 [1]. As a result, the food and agro-industrial industry have increased the production of products for human consumption. Complementarily, a slow progress in the development of sustainable and effective lines of waste management has generated the accumulation of agro-industrial residues [2, 3]. These residues englobe an environmental topic that may cause environmental damage; for example, it could release natural compounds that in higher concentrations could be toxic or

Cristobal N. Aguilar Gonzalez et al. (eds.), Food Waste Conversion, Methods and Protocols in Food Science, https://doi.org/10.1007/978-1-0716-3303-8_10, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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antinutritional (plants and animals) or changing nutritional properties in the ecosystem (e.g., eutrophication) [4]. Under the environmental risk, new strategies for residue management have been developed to revalorize the residual material to a raw material in other industrial processes. In this context, innovative technologies encompass an interesting alternative with the application of “green solvents,” short operating times, lower extraction temperature, and higher quality of the compounds [5–7], but they comprise expensive technology that requires specific knowledge for the operational procedure. Thus, another interesting alternative includes the application of microorganisms for the recovery or production of chemical components that could be applied in the food industry (bioactive secondary metabolites) [2, 3, 8]. In this context, the solid-state fermentation (SSF) encompasses a relevant tool for the extraction of polyphenolic compounds by providing the enzymatic system to degrade the cell wall (hydrolysis) and release of compounds in the extraction media [9–13]. The SSF is defined as a technology that uses microbial growth to produce biomolecules on a solid substrate used as physical support and as a source of nutrients in the absence or near absence of free-flowing liquid [8], with relevant advantages such as low energy consumption and a simpler extraction method for bioactive components. The SSF offers the possibility of applying bacteria or fungal strains, but the majority of studies has focused on fungal strains for efficacy, water requirements, enzymatic machinery, and the capacity to colonize the interparticle spaces of solid matrices [8]. Some examples of an application of SSF by fungal strain to recover bioactive compounds englobe the study of Torres-Leo´n et al. [12] where using an Aspergillus niger GH1 fungal strain enhanced the phenolic content and antioxidant activity in Mexican mango seed; the application showed an increment from 984 mg GAE/100 g to 3288 mg GAE/100 g (mg GAE = milligram equivalent of gallic acid) at 20 h of fermentation. Otherwise, another fungal strain of interest is Trichoderma harzianum, where the application has shown a 6.38-fold increase in total polyphenolic compounds in curry leaves with respect to the initial time [14] and 5.6-fold in turmeric fermented material with respect to the initial time [15]. Also, there are studies from other fungal strains to be applied for the bioactive compound recovery as Rhizopus spp. (R. oligosporus and R. oryzae). In this context, the main purpose of this protocol is to provide and describe solid-state fermentation as a valuable tool for utilizing food waste and the characteristics that agro-industrial waste must possess to be considered suitable fermentation substrates.

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Materials • Conditioning of the substrate – Support (agro-industrial wastes). – Drying chamber. – Cutter. – Sieve. • Inoculum preparation – Strain (fungus, yeast, bacteria). – Medium. – Flasks. – Tween 80. – Magnetic bar. – Micropipette. – Neubauer counting. – Microscope. • Humidity setting – Water activity meter. – Drying oven. – Desiccator. – Centrifuge. • Fermentation setup – Reactor. – Erlenmeyer flasks.

3

Methods

3.1 Conditioning of the Substrate

In recent years, different materials have been used as a substrate for fermentation processes; however, agro-industrial waste has shown increasing interest because, in addition to being used as fermentation supports, they can also be used as a substrate, since it has been shown that these matrices are made up of diverse and abundant compounds and nutrients of interest that microorganisms can use for their food [16, 17]. The agro-industrial waste that has been used is the peel and seeds of fruits and vegetables, as well as residues of cereals, grains, and legumes, including waste from the meat and dairy industry [18]. Therefore, it is important and necessary to carry out some primary processing, such as washing, disinfecting, component separation, size reduction, etc., in order to move on to the next point in the process.

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Drying is important for microbiological and enzymatic stabilization by reducing water activity of the substrate. Dehydrating agroindustrial residues helps increase its useful life since it does not have free water. Dehydrating helps prevent microbial growth or enzymatic processes, since having so low humidity decreases the chemical reactions within the product and controls the product’s respiration rate. [19] However, it is important to maintain the drying process at a temperature not higher than 60 °C since some of the biocomposites could be compromised. It seeks to reach a humidity of between 5% and 10% where the residues are stable [20, 21]. During the drying process for the processing of the raw material, it is essential to consider the characterization of the material, such as the amount of water it has, texture, porosity, volume, and chemical and physical composition it contains. Depending on the waste characteristics, the water that composes it can move inside it by different mechanisms, so the diffusion coefficient is important as it represents the movement of water from inside to outside. Mechanisms, such as liquid, surface, capillary, thermal, and steam diffusion, among others, are the main ones found in agro-industrial waste [22]. The diffusion capacity of the water in the fermentation substrate will be vital to ensure the correct development of the microorganisms during the fermentation process. There are different drying methods: sun drying, hot air drying, contact drying, infrared drying, freeze drying, fluid bed drying, and dielectric drying. However, to choose the best one, it is necessary to analyze some characteristics such as time available, requirements, costs, advantages, and disadvantages that the different types of drying entail. For example, drying in the sun could be an excellent option because of how economical it could be. It should be considered that since the material is exposed, depending on its composition and amount of sugars, it could be affected by microbial or pest contamination [23]. In the case of infrared drying, the depth of radiation penetration depends on the properties of the material to be dried and the wavelength of the radiation. The most widely used technique in agribusiness is infrared drying. Hot air eliminates the water contained in the product through evaporation, which prevents the growth of some bacteria that cannot live in dry media [24].

3.1.2 Grinding and Sieving

The following process for conditioning the raw material is the grinding process in which the particle size is reduced with the help of specialized equipment through mechanical force application. There are four most used reduction methods: compression, where solids are reduced into larger particles using up and down pressure; cutting, where specific cuts are made with the help of blades; rubbing or shearing, in which fine particles are obtained; and by impact where the particles are reduced with continuous

3.1.1

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blows. Currently, there is a wide variety of grinding equipment on the market with different mechanisms and components that are becoming more specialized and with greater technology [25]. In addition to reducing the particle size and being able to make the fermentation processes simpler, the grinding process is important so that the microorganism used for the bioprocess can have better adhesion to the substrate, which also helps to make the compounds or nutrients more available and their penetration into the matrix easier. In addition, it is also important because this will be the means by which moisture and communication will be available in the system [26]. Therefore, it is crucial after the grinding process to make a characterization of the particle sizes that were achieved. It is recommended to make the characterization using different sizes of sieves that will help to classify the particles. In this way, it is advisable to study the impact of particle size on the growth of the microorganism, adaptation of the microorganism, percentage of product extraction, and yield, among others. It has been shown that the size of the particle used in solid-state fermentation has influenced the ability of the microorganism to adapt, as well as its ability to produce different secondary metabolites, which represent compounds of industrial interest [20, 21]. 3.1.3

Storage

3.2 Inoculum Preparation

The last step in the preparation of raw material for its use as fermentation support is its adequate storage. It is important to keep it at an adequate temperature and in the dark. This is due to the high content of bioactive compounds that they possess, among which are sugars, minerals, vitamins, phenolic compounds, proteins, lipids, etc. Being sensitive, they could be affected and denatured. It is also important to keep them in a moisture-free environment since the matrices could rehydrate and become contaminated with opportunistic microorganisms. Another relevant step in the solid-state fermentation process englobes the inoculum preparation. It is a very important stage prior to starting the solid-state fermentation. Two strategies are considered for inoculum preparation; one is the conidia suspension, and the other is biphasic fermentation [27]. The first method is closely related to the preparation of inoculum for fungal strains and the second for the preparation of bacteria, yeast, and fungi inoculum. The following methodology presents both inoculum preparation and should work for most of the microbes used in solid-state fermentation.

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Fig. 1 Inoculation for the obtention of conidia in agar PDA or DSA

Conidia Suspension 1. The initial media is defined by the researcher. The main used culture media to activate fungus is potato dextrose agar and dextrose Sabouraud agar. The preparation must be carried out according to manufacturer instructions. For inoculum preparation, preferably prepare the media in an Erlenmeyer flask. The volume of the flask may vary according to the requirements of the researcher. 2. Inoculate the agar with spore suspension previously preserved. Most of the spores are cryopreserved. Add the desired volume of preserved spores into the flask containing the media (Fig. 1). The agar must be previously sterilized, tempered, and semiliquid in order to homogenize the spores into the media. This procedure must be carried out in sterile conditions. The flasks containing the inoculated media must be incubated at the microbe’s required temperature during the period that allows producing conidia. 3. It is required to prepare 10–15 mL of sterile Tween 80 solution (0.05–0.1% v/v in distilled water). The solution could be prepared and sterilized using a 15 mL centrifuge tube with a

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Fig. 2 Counting conidia into Neubauer chamber. (a) The Neubauer chamber, (b) view of the grid at 10× and signalization of squares to be counted (stars or numbers), and (c) view of the grid at 40× where the conidia must be seen clearly. All the conidia found within the medium square are counted even if they are partially out. For example, the gray conidia will be counted but not the red-colored (a) Count the conidia found in the grid. It must be counted from 5 to 13 squares. If too many conidia are visualized, dilutions must be carried out (b) Estimate the conidia content with the following equation: Conidia/mL = (Average conidia/ square)
(25 total squares)
(1 x 104)
(dilution factor)

cap. It is recommended to introduce a sterile magnetic bar into the tube in order to avoid contamination of the culture because of manipulation. 4. Once the conidia are produced, they must be harvested. Flood the Erlenmeyer flask with the Tween 80 solution, and carefully scrape the conidia using the magnetic bar and a magnetic stirrer at 100 rpm during 10–15 min. 5. Place the conidia solution into a sterile centrifuge tube with a cap with the aid of a 5 mL pipette. Avoid scratching the mycelium. The solution could be stored at 4 °C for a maximum of 4 days. 6. Then count the conidia in a Neubauer counting chamber as follows: (a) Place the chamber into the microscope, and focus the chamber at 10× in order to visualize the whole Neubauer chamber grid (it must be 25 main squares divided into 20 small squares). (b) Focus the microscope to 40×; the conidia must be seen clearly (Fig. 2). 7. Take the number of conidia required for the inoculation of the solid support that will be fermented. Most of the solid supports are inoculated with 1x107 spores per gram of support.

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Biphasic Fermentation This procedure is commonly used for large-scale solid-state fermentation bioreactors and must work for the culture of fungi, bacteria, and yeast. 1. Revitalizing the strains. Nutrient medium or MRS medium are commonly used for bacteria reactivaton (De Man, Rogosa, and Sharpe) medium. For yeast, YPD (yeast extract-peptone-dextrose) medium is used and for fungi the abovementioned potato dextrose or dextrose Sabouraud medium. Many researchers elaborate their synthetic media according to the nutrient requirements of the microbes. 2. Prepare from 50 to 100 mL of culture media and sterilize. Then inoculate the microorganism under sterile conditions. Culture the microbes according to the required time, temperature, and agitation speed. Generally, the culture finishes at the mid-exponential growth phase of the microorganism. 3. Once the time has finished, inoculate the solid support by adding 1–5% v/w of the prepared inoculum. It is very important to consider the amount of water added to the solid support to adjust the fermentation’s initial humidity content.

4

Humidity Setting This section presents the details for calculating the moisture adjustment parameters in the plant material for the solid-state process. The calculation of factors such as moisture, water activity (aw), water absorption index (WAI), and critical humidity point (CHP) are essential to guarantee the success of a solid fermentation using agro-industrial byproducts. To make use of agro-industrial wastes as fermentation substrates, mosture and aw are considered critical control points in microbial growth. Microbial growth is closely dependent on food’s water (free and bound) contents [28]. The aw values can be determined using a water activity meter. Equipment like AquaLab (Decagon Devices, 3TE, USA) or LabSwift-aw (Novasina, Lachen, Switzerland) are very reliable and give quick results. Typically, moisture and dry matter can be determined by the AOAC methods via a thermogravimetric approach. These methods are based on determining the mass of water present in a known mass sample [29]. Briefly dry the empty dish and lid in the oven at 105 °C for 3 h and transfer to the desiccator. Weigh about 5 g of sample into the crucible. Afterward, after 3 h at 105 °C, the crucible should be taken to the desiccator

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and weighed cold. The percentage of humidity is calculated with the following equation: Moisture ð%Þ =

ðWeight before drying - Weight after dryingÞ × 100 ðWeight before dryingÞ

ð1Þ

The moisture percentage can also be calculated with a moisture balance; this equipment is small and gives fast and precise results using the gravimetric moisture measurement principle. The humidity or weight loss results can be monitored on the equipment screen (final value or the value each time until constant weight). A weight greater than 0.5 g is necessary, and the test is destructive. The WAI is the quantity of water that the support can absorb [30]. According to Raimbault [31], to be used as substrate in SSF, the agro-industrial byproduct must attain several requirements, such as the ability to absorb water amounting to one or several times its dry weight, with a relatively high water activity on the solid/gas interface. Therefore, agro-industrial byproducts with high WAI are preferred for SSF since their moisture content can be modified during solid-state fermentation [32, 33]. WAI can be calculated by weight difference. Briefly, the samples (2.5 g) are dissolved in 30 mL of distilled water in glass containers with lids and cooked at 70 °C for 30 min in a water bath. The cooked samples are cooled to room temperature, transferred to pre-weighed centrifuge tubes, and centrifuged at 3000 RPM for 20 min. The supernatant is decanted (this will be used to determine CHP), and the sediment is weighed. WAI is calculated using the following equation: WAI

g Weight of sediment = g Weight of dry solids

ð2Þ

CHP represents the water linked to the support, which the microorganisms cannot use for their metabolic reactions [30]. According to He et al. [34], the agro-industrial subproduct must possess appropriate available moisture to support microbial growth and metabolism. The CHP can be calculated with any of the methods mentioned above to determine moisture (AOAC or thermobalance) by identifying the transition point between the periods of constant drying and decreasing drying speed. The supernatant obtained in the previous test (WAI) is used for the determination of CHP. The factors that control dehydration are temperature, humidity, air velocity, and dehydration time [35]. The fastest and most accurate method to calculate CHP is drying kinetics using a thermobalance at 60 °C with 1 g of the supernatant [30]. The calculation of the humidity value is made for each time with Eq. 1 (consider the final weight). CHP can be

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Fig. 3 Curve in the drying process

determined with a prototype drying process. Figure 3 shows a typical curve in the drying process. In this drying curve, the constant rate period, terminating at the CHP, followed by the falling rate period, can be identified. CHP is the moisture content at which the drying rate begins to decline [36]. Another way to determine CHP is by plotting the drying rate (evaporation per unit area) against moisture dry mass basis (for more details, the fundamentals of drying engineering can be consulted); the CHP is the point where the rate of drying changes. In summary, CHP is a humidity value that divides two drying periods: the period where all the free water is eliminated (speed is constant) and the period where only the bound water remains (fall in drying speed); this value is important in fermentation technology since the bound water cannot be used by the microorganism. The PCH is the limit humidity value at which fermentation can occur.

5

Fermentation Setup Finally, the fermentation assembly procedure is an important step to carry out a correct solid-state fermentation process. In addition to the previous preparations (substrate conditioning, humidity adjustment, and inoculum preparation), certain conditions must be met to ensure the proper development of this methodology. The most important point is to take care of sterility during the process [37], both in the place where it will be carried out and in the utensils and materials used. The solid-state fermentation process is very versatile, concerning the use of different types of reactors and amounts of substrates [38]. According to the final objective of the process depends the selection of type of reactor; the objectives can be as varied as where processes can be set up to analyze the behavior of a microorganism-substrate or the optimization of a process, carrying out fermentation kinetics at different times, modifying different factors (humidity, pH, substrate, supplementary

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Fig. 4 Scheme for the assembly of a laboratory-scale solid-state fermentation

media), analyzing the consumption-accumulation of products of interest, also the objective could be to obtain a higher volume of a product of interest (from a known process) where scaling up can be implemented in the process. The type of reactor can also be chosen depending on how sampling will be done (if included) or take the process to final time. Another important point to consider is that the reactor selection is conditioned by the final volume of the final product to be recovered. In this case, the protocol of a laboratory-scale fermentation setup will be demonstrated (Fig. 4), with the objective of studying the behavior of the accumulation-consumption of antioxidant compounds using an agro-industrial waste as substrate, using a filamentous fungus, performing a 120-h kinetic with sampling every 12 h, which will be carried out at a temperature of 30 °C, with an initial humidity of 70% and inoculum of 1 × 107 spores/ reactor, where each reactor will have 5 g of substrate. Some materials for assembly procedure englobe Erlenmeyer flasks, beaker, autoclave, spatula, micropipette (1000–100 μL, 200–20 μL), pipette (10–1 mL), pointers, laminar flow hood, and ethanol. Also, the volume capacity of material could change according to the SSF size and requirements. Procedure 1. The aim is to work under sterile conditions, so the process will be carried out in a laminar flow hood previously disinfected of any type of microorganism foreign to the process, with the use

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of alcohol, as well as the use of UV lamp, assuring that the workplace is innocuous. 2. The materials that will be used must be sterilized with the use of autoclave (temperature conditions 121 °C for a time of 15 min). For handling and preservation of the innocuousness of the materials prior to the process, it is recommended to line the objects with paper before sterilization, and for the reactors, cotton plugs are used, which allow the transfer of oxygen and keep the interior innocuous. 3. The amount of substrate is added to each reactor and reserved for later use. 4. Once the amount of water to be added for the 5 g of substrate is known, adjusting to 70%, also taking into account the amount of spore stock solution previously recovered, using the beaker and pipettes, the water to be used per reactor is added, together with the amount of spore stock solution to have the inoculum, a mixture that will be deposited in each of the reactors. 5. With the use of the spatula, the solid-liquid mixture should be homogenized in the reactor. 6. The cotton plug is placed again, labeled correctly, and placed in the incubator with the temperature at which the process will be carried out, which was previously disinfected in a similar way to the flow hood. References 1. UNITED NATIONS. DESA (2019) World population prospects 2019 – highlights. Department of Economic and Social Affairs World Population Prospects 2019, pp 1–39 2. Ravindran R, Jaiswal AK (2015) Exploitation of food industry waste for high-value products. Trends Biotechnol xx:1–12. https://doi.org/ 10.1016/j.tibtech.2015.10.008 3. Maria S, Shirahigue LD, Ceccato-antonini SR (2020) Agro-industrial wastes as sources of bioactive compounds for food and fermentation industries 4. Lopes FC, Ligabue-braun R, Bernardo S (2021) Agro-industrial residues: eco-friendly and inexpensive substrates for microbial pigments production. 5. https://doi.org/10. 3389/fsufs.2021.589414 5. Mohammadpour H, Sadrameli SM, Eslami F, Asoodeh A (2019) Industrial crops & products optimization of ultrasound-assisted extraction of Moringa peregrina oil with response surface methodology and comparison with Soxhlet method. Ind Crops Prod 131:106–116.

https://doi.org/10.1016/j.indcrop.2019. 01.030 6. Reddy AVB, Moniruzzaman M, Madhavi V, Jaafar J (2020) Recent improvements in the extraction, cleanup and quantification of bioactive flavonoids, 1st edn. Elsevier 7. Verma DK, Srivastav PP (2020) Extraction, identification and quantification methods of rice aroma compounds with emphasis on 2-Acetyl-1-Pyrroline (2-AP) and its relationship with rice quality: a comprehensive review. Food Rev Int 00:1–52. https://doi.org/10. 1080/87559129.2020.1720231 8. Savino S, Bulgari D, Monti E, Gobbi E (2021) Agro-industrial wastes: a substrate for multienzymes production by Cryphonectria parasitica, pp 1–15 9. Martins S, Mussatto SI, Martı´nez-avila G et al (2011) Bioactive phenolic compounds: production and extraction by solid-state fermentation. A review. Biotechnol Adv 29:365–373. https://doi.org/10.1016/j.biotechadv.2011. 01.008

Solid-State Fermentation as Strategy for Food Waste Transformation 10. Venil CK, Zulaikha N, Yusof B Solid state fermentation utilizing agro- industrial waste for microbial pigment production, pp 375–381 11. Sadh PK, Duhan S, Duhan JS (2018) Agro – industrial wastes and their utilization using solid state fermentation: a review. Bioresour Bioprocess:1–15. https://doi.org/10.1186/ s40643-017-0187-z 12. Torres-leo´n C, Ramirez-guzman N, Ascaciovaldes J, Serna- L (2019) AC SC. LWT Food Sci Technol. https://doi.org/10.1016/j.lwt. 2019.06.003 13. Chen G, Chen B, Song D (2021) Co-microbiological regulation of phenolic release through solid-state fermentation of corn kernels (Zea mays L.) to improve their antioxidant activity. 142 14. Salah HA, Bassuiny RI, El-Khonezy MI et al (2019) Impact of solid state fermentation by Trichoderma spp. on phenolic content, antioxidant and antibacterial activities of curry leaf powder. J Food Meas Charact 13:1333–1340. https://doi.org/10.1007/s11694-01900048-0 15. Mohamed SA, Saleh RM, Kabli SA, Al-garni SM (2016) Influence of solid state fermentation by Trichoderma spp. on solubility, phenolic content, antioxidant, and antimicrobial activities of commercial turmeric. Biosci Biotechnol Biochem 8451:1–9. https://doi.org/ 10.1080/09168451.2015.1136879 16. Torres-Leo´n C, Ramı´rez-Guzman N, Lon˜ o-Hernandez L et al (2018) Food waste don and byproducts: an opportunity to minimize malnutrition and hunger in developing countries. Front Sustain Food Syst 2:52. https://doi.org/10.3389/fsufs.2018.00052 17. Angulo-Lo´pez JE, Flores-Gallegos AC, TorresLeo´n C et al (2021) Guava (Psidium guajava l.) fruit and valorization of industrialization by-products. Processes 9:1–17. https://doi. org/10.3390/pr9061075 18. Flores-Maltos DA, Mussatto SI, Esquivel JCC et al (2014) Typical mexican agroindustrial residues as supports for solid-state fermentation. Am J Agric Biol Sci 9:289–293. https:// doi.org/10.3844/ajabssp.2014.289.293 19. Reynaldo De la Cruz-Quiroz SR, CNA (2018) Production of a biological control agent: effect of a drying process of solid-state fermentation on viability of trichoderma spores. Int J Green Technol 4:01–06. https://doi.org/10.30634/ 2414-2077.2018.04.1 20. Torres-Leo´n C, Ramirez-Guzman N, AscacioValdes J et al (2019) Solid-state fermentation with Aspergillus niger to enhance the phenolic contents and antioxidative activity of Mexican

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mango seed: a promising source of natural antioxidants. LWT 112:108236. https://doi.org/ 10.1016/j.lwt.2019.06.003 21. Robledo A, Aguilera-Carbo´ A, Rodriguez R et al (2008) Ellagic acid production by Aspergillus niger in solid state fermentation of pomegranate residues. J Ind Microbiol Biotechnol 35:507–513. https://doi.org/10.1007/ s10295-008-0309-x 22. Reinaldo Sanchez Arriagada (2007) Determinacion coeficiente convectivo de materia en el secado de so´lidos. Maderas Cienc y Tecnol 9: 245–268 23. Correˆa PC, de Oliveira GHH, Santos E de S (2012) Thermodynamic properties of agricultural products processes. Phys Prop Foods Nov Meas Tech Appl 72:131–142. https://doi. org/10.1201/b11542 24. Correˆa PC, de Oliveira GHHH, Baptestini FM, Mayra Darliane Martins Silva Diniz AADP (2012) Tomato infrared drying: modeling and some coefficients of the dehydration process. Chil J Agric Res 262–267 25. Chauruka SR, Hassanpour A, Brydson R et al (2015) Effect of mill type on the size reduction and phase transformation of gamma alumina. Chem Eng Sci 134:774–783. https://doi.org/ 10.1016/j.ces.2015.06.004 26. Lopez YB, Roa DF, Bravo JE (2022) Efecto del tratamiento te´rmico en la estabilidad de geles obtenidos a partir de harinas de quinua. Inf Tecnolo´gica 33:203–214. https://doi.org/ 10.4067/s0718-07642022000100203 27. Jaronski ST, Mascarin GM (2017) Mass production of fungal entomopathogens. In: Lacey LA (ed) Microbial Control of insect and mite pests: from theory to practice. Academic Press, Yakima, pp 141–155 28. da Silva MU, Sato J, Ribeiro PM, Janeiro V, Ribeiro LB, Vasconcellos RS (2022) Modeling moisture adsorption isotherms for extruded dry pet foods. Anim Feed Sci Technol 290: 1 1 5 3 1 8 . h t t p s :// d o i . o r g / 1 0 . 1 0 1 6 / j . anifeedsci.2022.115318 29. AOAC (2000) Loss on drying (moisture) 930.15-1930. Arlington 30. Torres-Leo´n C, Ramirez-Guzman N, AscacioValdes J, Serna-Cock L, dos Santos Correia MT, Contreras-Esquivel JC, Aguilar CN (2019) Solid-state fermentation with Aspergillus niger to enhance the phenolic contents and antioxidative activity of Mexican mango seed: a promising source of natural antioxidants. LWT Food Sci Technol 112:108236. https://doi. org/10.1016/j.lwt.2019.06.003

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31. Raimbault M (1998) General and microbiological aspects of solid substrate fermentation. Electron J Biotechnol 1:1–15 32. Robledo A, Aguilera A, Rodriguez R, Martinez JL, Garza Y, Aguilar C (2008) Ellagic acid production by Aspergillus niger in solid state fermentation of pomegranate residues. J Ind Microbiol Biotechnol 35:507–513. https:// doi.org/10.1007/s10295-008-0309-x 33. Orzua MC, Mussatto SI, Contreras JC, Rodriguez R, de la Garza H, Teixeira JA, Aguilar CN (2009) Exploitation of agro industrial wastes as immobilization carrier for solid-state fermentation. Ind Crops Prod 30:24–27. https://doi.org/10.1016/j.indcrop.2009. 02.001 34. Buenrostro-figueroa JJ, Vela´zquez M, Floresortega O, Ascacio-valde´s JA, Huerta-Ochoa S, Aguilar CN, Prado-Barragan L (2017) Solid state fermentation of fig (Ficus carica L) by-products using fungi to obtain phenolic compounds with antioxidant activity and qualitative evaluation of phenolics obtained. Process Biochem 62:16–23. https://doi.org/10. 1016/j.procbio.2017.07.016

35. He Q, Peng H, Sheng M, Hu S, Qiu J, Gu J (2019) Humidity control strategies for solidstate fermentation: Capillary water supply by water-retention materials and negativepressure auto-controlled irrigation. Front Bioeng Biotechnol 7:1–13. https://doi.org/ 10.3389/fbioe.2019.00263 36. Torres-Leo´n C, Aguilar C (2022) Food preservation. In: Sepulveda L, Aguilar NC, Porteen K, Haghi A (eds) Quantitative methods and analytical techniques in food microbiology, 1st edn. Apple Academic Press, Abingdon, pp 39–57 37. Fang J, Liu Y, Huan C, Xu L, Ji G, Yan Z (2020) Comparison of poly-γ-glutamic acid production between sterilized and non-sterilized solid-state fermentation using agricultural waste as substrates. J Clean Prod 255:120248 38. Arora S, Rani R, Ghosh S (2018) Bioreactors in solid state fermentation technology: design, applications and engineering aspects. J Biotechnol 269:16–34

Chapter 11 Protocol for the Solid-State Fermentation-Assisted Extraction (SSFAE) of Bioactives from Tomato Waste: The Case of Carotenoids Juanita Y. Mendez-Carmona, Karen N. Ramı´rez-Guzman, Juan A. Ascacio-Valdes, Leonardo Sepulveda, Jose´ Sandoval, and Cristobal N. Aguilar Gonzalez Abstract Tomato agro-industry generates significant amounts of waste along the production chain. This residue is a good source of nutrients and bioactive compounds. The tomato waste is a carotenoid-rich biomass, mainly lycopene which is recognized for its powerful biological activity with relevance in the pharma, food, and cosmetic industry; for this reason, several extraction technologies have been evaluated to separate these biomolecules; one of them is the solid-state fermentation-assisted extraction (SSFAE). In this chapter, we describe a protocol that has been developed to recover carotenoids, mainly lycopene, from tomato waste using SSFAE. Specifically, we used the tomato waste as solid support for the fermentation, employing a fungal strain as source of the enzymatic and metabolic machinery to separate the target molecule from the tomato biomass. Our goal is to provide the reader with guidelines on how to apply the SSFAE of bioactives from tomato waste. Key words Tomato waste, Carotenoids, Lycopene, Solid-state fermentation, Aspergillus niger

1

Introduction Carotenoids include a variety of tetraterpenes distributed in plants, animals, bacteria, fungi, and algae. About 850 different natural carotenoids have been reported. Most pigments present 8 isoprene units with 40 carbons skeleton that usually contain 9 double conjugate bonds [1]. They are isoprenoids characterized by their attractive shades that come in yellow, orange, red, and purple. Its synthesis is photosynthetic in plants and not photosynthetic in bacteria and fungi. That is why the intake of foods rich in carotenoids through the daily diet of animals and humans is indispensable to obtain the benefits demonstrated in this group of pigments [2],

Cristobal N. Aguilar Gonzalez et al. (eds.), Food Waste Conversion, Methods and Protocols in Food Science, https://doi.org/10.1007/978-1-0716-3303-8_11, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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benefits demonstrated in various studies from the usual consumption in the diet, which evoke in the reduction of the development of some types of cancer (prostate, cervical, colorectal, breast, and liver, among others) and in the prevention of cardiovascular diseases; skin, bone, and eye disorders; infectious diseases; and leukemia, to name a few [3–5]. Therefore, the commercial growth that carotenoid pigments have acquired in recent decades is not new. However, considering that molecules are highly sensitive to heat, oxygen, light, exposure to oxidative enzymes, acids and transition metals, there is a need to implement and develop extraction methodologies that not only quantify high amounts of carotenoids but allow the recovery of molecules using more environmentally friendly technologies [6], the need to implement and develop extraction methodologies that don’t only quantify high amounts of carotenoids but allow the recovery of molecules by means of more environmentally friendly technologies, which in turn allow the application of the recovered extracts in the food industry while preserving the quality thereof, in addition to meeting current demands governed by a circular economy. One of the ecologically viable alternatives includes the revaluation of agro-industrial wastes for the extraction and recovery of biologically active compounds. In this sense, tomato is one of the most productive vegetables worldwide [7]. Therefore, large quantities of the product are lost and/or wasted at different stages of the production chain. This not only represents food losses but also includes the loss of nonrenewable natural resources involved in harvesting the product. Tomato contains some nutrients including protein, fiber, saturated fatty acids, as well as vitamins and minerals. On the other hand, it is important to recognize that the presence of secondary metabolites such as carotenoids, mainly lycopene and β-carotene, ascorbic acid, tocopherol represents an advantage in favor of the prevention of chronic-degenerative diseases including cardiovascular and neurodegenerative diseases, as well as some types of cancer [8]. The tomato stands out for its lycopene content, which represents approximately 80% of the total carotenoids present in the fruit [9]. Given the importance of nutrition and human health that could be attributed to these compounds, various technologies have been used to extract and recover carotenoids from tomato residues. These include microwave-assisted extraction, ultrasoundassisted extraction, electrical pulses, enzyme-assisted extraction, and high-pressure-assisted hydrostatic extraction, to name a few [10]. Despite the known advantages of so-called emerging technologies, due to the high costs in the acquisition and maintenance of some equipment, as well as the complexity of certain extractive processes, other alternatives have been explored and successfully applied in the extraction of bioactive compounds from agroindustrial waste, such as solid-state fermentation (SSF). The SSF is defined as the fermentative process involving solids in absence, or

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near absence, of free water. Nonetheless, the substrate must have sufficient moisture to make the growth and metabolism of the microorganism suitable [11]. SSF has become an excellent technique for biotransforming agro-industrial waste and by-products. In this sentence, SSF as a process of extraction and recovery of bioactive compounds present in food waste has been extensively explored, thanks to the fact that it offers greater efficiency in the fermentative process, an increase in the concentration and quality of the final products, and greater stability of the active compounds [12, 13]. It is important to note that this strategy called SSFAE was originally proposed by our research group [14, 15]. Due to the significant amounts of residual tomato generated, as well as the nutritional properties and bioactive compounds of interest contained in the fruit, the present protocol proposes to use the rejected tomato for sale and/or consumption due to its excessive state of maturation as a substrate for SSFAE for the recovery of carotenoids.

2

Materials

2.1

Raw Material

1. The tomato waste (Solanum lycopersicum) was recollected, selected, and processed (see Note 1).

2.2

Microorganism

1. Aspergillus niger GH1 (A. niger GH1) strain from the DIA/UADEC (Departamento de Investigacio´n en Alimentos/Universidad Auto´noma de Coahuila) collection were used in this study.

2.3 Solid-State Fermentation

1. Raw material (previously processed). 2. Petri dishes (60 × 15 mm). 3. Distilled water. 4. A. niger GH1 reactivated on PDA agar for 5 days at 30 °C.

2.4 Recovery of Carotenoids

1. Acetone in a 1:2 (v/w) ratio for the recovery of the bioactive compounds in the SSF.

2.5 Analysis of Carotenoids

1. Quartz cell. 2. 0.45 mL membrane filter. 3. Acetone.

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Methods

3.1 Processing of the Raw Material

1. The tomato waste was collected and selected based on quality criteria recognized mainly by the perception of sight and touch related to color and texture. Tomatoes of intense red color were selected, irregularly, with loss of firmness to the touch, for their rejection to the sale and consumption of the fruit, in addition to selecting those tomatoes without apparent microbiological contamination. 2. The tomato wastes were washed with water and then immersed in a commercial bactericidal solution regarding 10 drops of the product per liter of water, left to stand for 15 min, and finally the tomatoes were cut into quarters for storage at -20 °C until further use. 3. The tomato waste was defrosted for 12 h protected from light, to dehydrate in a conventional oven at 60 °C for 12 h, and finally grind the samples in a mill to a 75 μM particle size. The ground tomato was stored in closed containers, protected from light at room temperature until later use.

3.2

Test for Support

1. The potential of SSF support of the sample was determined by the evaluation of the water absorption index (WAI), critical humidity point (CHP), water activity (aw), pH, and invasion capacity (Table 1). 1.1 To determine the WAI, 2.5 g of the sample were dissolved in 30 mL of distilled water and cooked at 70 °C for 30 min in a water bath. The cooked paste was cooled to room temperature and transferred to pre-weighed centrifuge tubes and centrifuged at 3000 rpm for 20 min. The supernatant liquid was decanted, and the resulting gel weighed. WAI was calculated using the following equation and expressed as grams of gel obtained per gram of solid: Table 1 Characterization of tomato waste as a support for SSF Analyses

Range

CHP (%)

75.04

WAI (g/g)

3.14

aw

0.35

pH

4.13

Growth rate (mm/h)

0.89

Humidity (%)

70

WAI water absorption index, CHP critical humidity point, aw water activity

Protocol for the Solid-State Fermentation-Assisted Extraction (SSFAE). . .

WAI

g g

=

165

weight of sediment weight of dry soilds

1.2 CHP was estimated using a thermobalance. The drying kinetics was performed with 0.60 ± 0.01 g of the sample impregnated with saturated water (WAI result) at 60 °C. 1.3 Aw was evaluated using AquaLab to 25 ± 2 °C. 1.4 pH was measured using a pH meter; 0.300 g of sample was homogenized with 30 mL of distilled water. Subsequently, pH was measured by direct immersion of the electrode. 1.5 To estimate the number of solids (substrate) and liquid for the fermentation process, the balance of matter was calculated based on the data previously obtained in the moisture determination on the processed sample. The adjustment was made to 70% humidity and 30% solids. 1.6 The invasion capacity (fungal growth) was evaluated in 10-cm-diameter petri dishes with the tomato that was previously processed. A. niger GH1 was reactivated on PDA agar for 5 days at 30 °C. Explants (8 mM) were inoculated in the center of the plates; radial growth was monitored kinetically (6 h). All tests were performed in triplicate, and the invasion capacity was expressed as the growth rate in mm/h. 3.3 Solid-State Fermentation

1. Reactors of petri dishes (60 × 15 mM) with 2.91 g of sample and 7.09 mL of distilled water inoculated with 1 × 106 spores per gram of support were used (see Note 2). 2. The samples were incubated at 30 °C. 3. The extraction was performed with acetone in a 1:2 (w/v) ratio. The extracts were stored in containers protected from light at -20 °C until analysis (see Note 3). 4. Total sugar consumption was determined by the phenolsulfuric method proposed by Dubois et al. [16], and the fungal biomass estimated under the methodology proposed by Co´rdova-Lo´pez et al. [17], to select the final fermentation time in relation to the total carotenoid recovery and biomass. (see Note 4) (Table 2).

3.4 Quantification of Carotenoids

1. Total carotenoid concentration was detected and quantified by high-performance liquid chromatography (HPLC) and spectrophotometrically at an optical density of 460 nM following the below equation:

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Table 2 Productivity for treatment by time and estimation of total carotenoid content

Biomass (mg/g dw)

Total carotenoid recovery Performance (mg L/mg L) (mg/g dw)

Standard Treatment Average deviation

Standard Average deviation

Carotenoid/ biomass

Productivity

0

0.054

0.0253233

0.0034

0.000847813

0.062962963

0.0000

6

0.053

0.00267604

0.0019

0

0.035849057

0.0003

12

0.051

0.00292382

0.0037

0.001243991

0.07254902

0.0003

24

0.087

0.00619454

0.0051

0.001195629

0.05862069

0.0002

36

0.062

0.00292382

0.0102

0.000521229

0.164516129

0.0003

48

0.061

0.00183359

0.01

0.004019708

0.163934426

0.0002

dw dry weight

Yx =

v  A  106 E  100  ms 1%

where Yx represents the carotenoid yield (μg/g dw), A the optical density (OD480nM), v the volume of solvent used (mL), ms the dry cell mass (g), and E1% the specific absorptivity of solvent (2150) [18]. 2. For the HPLC carotenoid determination, 1.5 mL of extract were filtered through a 0.45 μM membrane filter and subsequently injected into the HPLC equipment (Pursuit XRs C18 column, 150 × 4.6 mM, 5 μM). The HPLC analysis was determined by gradients: phase A, methanol; phase B, acetone 75%; and phase C, acetone 95%, flow 1.5 mL/min, and UV detector at 460 nM.

4

Notes 1. The tomato was collected from the local market in the city of Saltillo, Coahuila, Mexico. The tomatoes selected were those discarded for consumption and sale due to their excessive state of ripeness (+6) [19]. 2. The spore count in the Neubauer chamber was performed; then the inoculum size to be added was calculated, which would be subtracted from the distilled water mL calculated for each reactor. Finally, the homogeneous mixture of the liquid with the solids was performed.

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3. For the recovery of the fermented mass compounds, the amount of solvent stipulated for each reactor was added; a homogeneous mixture was made with the help of a sterile spatula, to finally extract the compounds of interest using a sterile muslin fabric to separate/filter the liquid from the solid compound. 4. The profiles of glucose consumption, biomass production, and total carotenoid concentration were modeled to validate and control the bioprocess. To determine the biomass, it is necessary to reserve a part of the fermented solid support. The fermented sample was frozen at -20 °C until further use.

Acknowledgments Me´ndez-Carmona thanks the National Council of Science and Technology, Mexico (CONACYT), for the financial support given to her master’s degree studies at DIA-FCQ/UADEC. References 1. Maoka T (2020) Carotenoids as natural functional pigments. J Nat Med [Internet] 74(1): 1–16. Available from: https://doi.org/10. 1007/s11418-019-01364-x 2. Zia-Ul-Haq M (2021) Historical and introductory aspects of carotenoids. In: Zia-UlHaq M, Dewanjee SRM (eds) Carotenoids: structure and function in the human body. Cham, Springer, pp 1–42 3. Mele´ndez-Martı´nez AJ, Mandic´ AI, Bantis F, Bo¨hm V, Borge GIA, Brncˇic´ M et al (2020) A comprehensive review on carotenoids in foods and feeds: status quo, applications, patents, and research needs. Crit Rev Food Sci Nutr [Internet] 1–51. Available from: https://doi.org/ 10.1080/10408398.2020.1867959 4. Zia-Ul-Haq M, Duo D, Riaz M (2021) Carotenoids: structure and function in the human body. Springer, 859 p 5. Nabi F, Arain MA, Rajput N, Alagawany M, Soomro J, Umer M et al (2020) Health benefits of carotenoids and potential application in poultry industry: a review. March: 1809–1818 6. Zhao X, Kehong L, Hong Z (2022) Carotenoids in cereals and related foodstuffs: a review of extraction and analysis methods. Food Rev Int

7. Li N, Wu X, Zhuang W, Xia L, Chen Y, Wu C et al (2021) Tomato and lycopene and multiple health outcomes: umbrella review. Food Chem 343(September) 8. Ali MY, Sina AAI, Khandker SS, Neesa L, Tanvir EM, Kabir A et al (2021) Nutritional composition and bioactive compounds in tomatoes and their impact on human health and disease: a review. Foods 10(1) ´ etkovic´ G, C ´ anadanovic´-Brunet J, 9. Stajcˇic´ S, C ˇ etojevic´-Simin D (2015) Djilas S, Mandic´ A, C Tomato waste: carotenoids content, antioxidant and cell growth activities. Food Chem 172:225–232 10. Ashraf W, Latif A, Lianfu Z, Jian Z, Chenqiang W, Rehman A et al (2020) Technological advancement in the processing of lycopene: a review. Food Rev Int:1–27 11. Pandey A (2003) Solid-state fermentation. Biochem Eng J 13:81–84 12. Chilakamarry CR, Mimi Sakinah AM, Zularisam AW, Sirohi R, Khilji IA, Ahmad N et al (2022) Advances in solid-state fermentation for bioconversion of agricultural wastes to value-added products: Opportunities and challenges. Bioresour Technol [Internet] 343

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(August):126065. Available from: https://doi. org/10.1016/j.biortech.2021.126065 13. Aguilar CN, Aguilera-Carbo A, Robledo A, Ventura J, Belmares R, Martinez D et al (2008) Production of antioxidant nutraceuticals by solid-state cultures of pomegranate (Punica granatum) peel and creosote bush (Larrea tridentata) leaves. Food Technol Biotechnol 46(2):218–222 14. Martins S, Mussatto SI, Martı´nez-Avila G, ˜ ez-Saenz J, Aguilar CN, Teixeira JA Montan (2011) Bioactive phenolic compounds: Production and extraction by solid-state fermentation. A review. Biotechnol Adv [Internet] 29(3):365–73. Available from: https://doi. org/10.1016/j.biotechadv.2011.01.008 15. Ventura J, Belmares R, Aguilera-Carbo A, Gutie´rrez-Sanchez G, Rodrı´guez-Herrera R, Aguilar CN (2008) Fungal biodegradation of tannins from creosote bush (larrea tridentata) and tar bush (fluorensia cernua) for gallic and

ellagic acid production. Food Technol Biotechnol 46(2):213–217 16. Dubois M, Gilles KA, Hamilton JK, Rebers PT, Smith F (1956) Colorimetric method for determination of sugars and related substances. Anal Chem 28(3):350–356 17. Co´rdova-Lo´pez J, Gutie´rrez-Rojas M, ˜ eda G, Favela-Torres Huerta S, Saucedo-Castan E (1996) Biomass estimation of aspergillus Niger growing on real and model supports in solid state fermentation. Biotechnol Tech 10(1):1–6 18. Rodriguez-Amaya DB (2001) A guide to analysis in carotenoid. ILSI Press, Washington 19. Camelo AF Manual para la preparacio´n y venta de frutas y hortalizas [Internet]. Servicios Agricolas de la FAO. 2003 [cited 2022 Feb 20], p 1. Available from: https://www.fao.org/3/ y4893s/y4893s00.htm#Contents

Chapter 12 Protocol for the Production of Trichoderma Spores for Use as a Biological Control Agent Through the Revalorization of Agro-industrial Waste Salvador A. Saldan˜a-Mendoza, Mo´nica L. Chavez-Gonza´lez, and Cristobal N. Aguilar Gonzalez Abstract Trichoderma is one of the most widely used biological control agents worldwide. In addition to protecting crops through various mechanisms such as competition for nutrients and space, mycoparasitism, antibiotic production, and induction of the systemic immune response in plants, it is widely recognized for its beneficial effects on crops through the development of plants by increasing the availability of nutrients and promoting growth and development of the root system, as well as the degradation of synthetic pesticides. The most used form for its application in the field is spores, which can be produced by solidstate fermentation systems. This characteristic allows the revalorization of agro-industrial residues as a substrate in the production processes of this biological control agent, contributing to the reduction of production costs as well as the mitigation of the sources of contamination represented by the disposal of these wastes in the fields. This protocol establishes the guidelines to follow for the evaluation of the physicochemical properties of an agro-industrial residue for its use as a support in solid-state fermentation processes, as well as for its revalorization through its use in the production of Trichoderma spores for its use as a biological control agent through solid-state fermentation processes. Key words Biocontrol, Agro-industrial residues, Trichoderma, Solid-state fermentation

1

Introduction Every year the world population increases considerably, making it necessary to work on aspects that ensure the maintenance of food demand. One of agricultural production’s main problems is the attack by phytopathogens, which generate enormous crop losses due to multiple diseases [1]. The most used strategy for the control of phytopathogens is the use of synthetic pesticides. However, the uncontrolled and long periods of application of these chemicals have generated resistance in phytopathogens to combat, in addition to severe problems of environmental contamination and damage to

Cristobal N. Aguilar Gonzalez et al. (eds.), Food Waste Conversion, Methods and Protocols in Food Science, https://doi.org/10.1007/978-1-0716-3303-8_12, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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health, being associated with mutagenic damage, carcinogenicity, and teratogenicity [2]. For these reasons, current international policies point to the use of safer natural alternatives for the control of phytopathogens and crop protection, such as the use of biological control agents [3], among which Trichoderma stands out. Trichoderma is a filamentous fungus widely recognized for its ability to control phytopathogens through different mechanisms, such as competition for nutrients and space, mycoparasitism, antibiosis, and induction of the systemic immune response in plants. In addition, it provides benefits for crops by removing synthetic pesticides in the field, increasing the availability of nutrients, and promoting the growth of plants and their root systems, which contributes to the increase in agricultural production [4–6]. Its application in the field is commonly executed through spores, which must be applied in large quantities to achieve a successful effect [7]. Another of the serious problems currently facing the agricultural industry is the generation of enormous amounts of lignocellulosic waste in the form of seeds, stems, husks, bagasse, and skins as residues from the processing of wine, beer, coffee, vegetables, and fruits, among many others [8]. These wastes are commonly discarded into the environment where they become a severe pollution problem, incinerated to prevent the proliferation of parasites and microorganisms, or, in the best of cases, destined for animal feed. However, thanks to their composition, rich in sugars and nutrients, they can be revalorized through biotechnological systems to generate new value-added products [8, 9], in this case, Trichoderma spores for use as biocontrol agents. Solid-state fermentation (SSF) is a biotechnological process characterized by free water’s absence or near absence. It does not require sophisticated equipment for its development and allows the use of agro-industrial residues as a substrate without requiring rigorous pretreatments [10]. In this way, the application of SSF through the revalorization of agro-industrial waste contributes by making the production of biocontrol agents cheaper, where the substrates represent up to 40% of the costs, reducing the impact that waste has on the environment as a source of contamination [11]. This system has been widely used for developing bioprocesses, especially those in which filamentous fungi are involved since it mimics the conditions in which they develop in nature compared to its liquid counterpart, submerged fermentation (SmF) [12]. Likewise, it has been reported that the spores produced in SSF have a longer shelf life than those obtained through SmF processes [7]. Numerous investigations have reported the production of Trichoderma spores through the revalorization of agro-industrial residues through SSF processes [11, 13–15]. In the same way, our

Protocol for the Production of Trichoderma Spores. . .

171

research group has reported the production of T. asperellum spores through the revaluation of corn cob residues with excellent results [16]. This protocol provides a detailed explanation of the procedures necessary to identify if a residue has the appropriate physicochemical characteristics to be used as a support in SSF processes, as well as the procedure to follow for its use by SSF for the production of Trichoderma spores for its use as a biocontrol agent.

2

Materials

2.1

Raw Material

1. Agro-industrial residue to be revalorized by SSF.

2.2

Microorganism

1. Trichoderma sp. strain to be used to produce spores for its application as a biological control agent.

2.3 Tests for the Use of Agro-Industrial Residues as Support for SSF

1. Sterile water. 2. Falcon Tubes. 3. Centrifuge. 4. Balance. 5. Thermobath. 6. Thermobalance. 7. Potentiometer or pH test strips. 8. Petri dishes (60 × 15 mM). 9. Vernier.

2.4 Preparation of SSF for Trichoderma sp. Spore Production

1. Polyethylene bags. 2. 250 mL Erlenmeyer flasks. 3. Autoclave. 4. 1% Tween 80 solution. 5. Magnetic stirrer. 6. Stirring rack. 7. 10 mL stoppered glass tubes. 8. Neubauer chamber. 9. 1 mL micropipette. 10. 200 μL micropipette. 11. mL tips. 12. 200 μL tips.

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Methodology

3.1 Raw Material Pretreatment

1. A residue will be selected for the development of the process (see Note 1). 2. The selected residue will be dried in an oven for 24 h at 60 °C. 3. Once dry, it will undergo a grinding process to reduce the particle size (see Note 2). 4. Finally, the material will be stored at room temperature (25 °C) until use (see Note 3).

3.2 Tests for the Use of Agro-industrial Residues as Support for SSF

3.2.1

WAI Determination

The substrate to be revalorized must have characteristics that allow it to be used as a support in the SSF process. For this, analyses will be carried out to determine the water absorption index (WAI), critical humidity point (CHP), and pH determination using the ˜ a-Mendoza et al. [10]. methods reported by Saldan 1. 1.25 g of the dry sample will be weighed and mixed with 30 mL of sterile distilled water using a Falcon tube. 2. The sample will be placed in a water bath at 60 °C for 30 min and mixed every 10 min. 3. Subsequently, it will be centrifuged at 4900 rpm, and the supernatant will be decanted. 4. The formed gel will be weighed, and the WAI will be determined using the following formula: WAI =

3.2.2

CHP Determination

gel weight weigt of dry sample

1. Drying kinetics will be carried out using a thermobalance. 0.5 g of the gel obtained in the WAI determination will be placed and subjected to a drying process at 120 °C, evaluating the percentage of moisture lost every minute. 2. A drying speed graph will be prepared using the information obtained from this test, determining the critical point of humidity, the point at which the initial drying speed is reduced.

3.2.3

pH Determination

1. One gram of the dry sample will be homogenized in 10 mL of distilled water at room temperature (25 °C). 2. Subsequently, the pH of the sample will be determined by using a potentiometer or, failing that, by using pH test strips.

Protocol for the Production of Trichoderma Spores. . .

3.3 Evaluation of the Adaptation Capacity of the Trichoderma sp. Strain to the Substrate

173

For evaluation of the adaptation capacity of the microorganism to the substrate; this will determine the radial growth velocity using ˜ a-Mendoza et al. [10] with the method described by Saldan modifications: 1. The Trichoderma sp. strain will be reactivated on PDA agar at 25 °C for 4 days. 2. On the other hand, a petri dish will be prepared with 5 g of the dry substrate to be evaluated and impregnated with 5 mL of sterile distilled water, which must be distributed evenly, without compacting, on the surface of the plate. 3. A 5 mm mycelial disk will be obtained from the peripheral area of the Trichoderma sp. colony using a punch and placed in the center of the plate with the substrate. 4. The assay will be left to incubate at 25 °C. The radial growth of the microorganism will be determined using a Vernier every 12 h until the whole invasion. The invasion capacity will be expressed as the growth rate (mm/h).

3.4 Preparation of SSF for Trichoderma sp. Spore Production

Following the methodology described by De la Cruz-Quiroz et al. [17] with modifications: 1. Polyethylene bag bioreactors with 50 g of the selected substrate, previously dehydrated, will be used. The fermentation conditions will be 50% humidity and an inoculum of 1 × 107 spores/mL. 2. The Trichoderma sp. strain will be reactivated in three 250 mL Erlenmeyer flasks with 50 mL of previously sterilized PDA (121 °C, 15 psi, 20 min) at 30 °C for 4 days (see Note 4). 3. 50 mL of a 1% (v/v) Tween 80 solution (see Note 5) will be added and left stirring with the help of a magnetic stirrer for 3 min to recover the spores. 4. The content of the first flask will be transferred to the second, and the same process will be repeated without the addition of Tween 80 solution. Subsequently, the process will be repeated with the third flask to obtain the stock solution of spores. 5. Take 1 mL of the stock solution of spores and dilute it in 9 mL of sterile distilled water to obtain a 1:10 dilution. The process will be repeated to obtain a 1:100 dilution. 20 μL of the 1:100 solution will be taken for the spore count in the Neubauer chamber. The spores in 13 of the 25 central squares of the chamber will be counted (see Note 6). The spore count will be carried out in triplicate. The spore concentration of the spore stock solution will be determined using the following equation: espores=mL = A  25  10,000  FD  13

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where A is the average number of spores obtained from the triplicate count and FD is the dilution factor used, which in this case corresponds to a value of 100. 6. Once the spore concentration of the stock solution has been determined, the calculation will be made to adjust the volume of the inoculum using the formula described below: V1=

C2  V 2 C1

where C1 corresponds to the spore concentration of the stock solution, V1 is the unknown volume to be used in mL to obtain the desired volume, C2 corresponds to the desired spore concentration, and V2 corresponds to the desired volume of solution. 7. The substrate will be impregnated with the inoculum solution and mixed to homogenize. 8. The fermentation will be incubated at 25 °C until the total invasion of the substrate (see Note 7). 9. At the end of the fermentation, a sample of 1 g of ferment will be taken and mixed with 9 mL of sterile distilled water. The spore count will be carried out using a Neubauer chamber using the previously described methodology. The spores produced by this method can be used directly in the field, preferably incorporated into the root area of the crops to be treated. Preventive treatment with Trichoderma sp. spores guarantees better results for the control of diseases caused by the attack of phytopathogens.

4

Notes 1. The selected waste must be clean and free of contamination. 2. Avoid using too small particle sizes at the level of powders. Too small particle size can agglomerate during SSF, generating gas exchange problems and limiting the availability of the interparticle space, thus limiting the microorganism’s access to nutrients and hindering its development. 3. Use sealed bags or plastic boxes. Insects can quickly invade the raw material. 4. For this point, the flasks should present a uniform dark-green color, characteristic of this microorganism, provided by sporulation. 5. Uncountably agglomerated spores will be considered as only one spore during the Neubauer chamber count.

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6. The incubation period may vary depending on the Trichoderma sp. strain used and its invasion capacity on the substrate to be revalorized.

Acknowledgments ˜ a-Mendoza thanks the National Council of Science and Saldan Technology, Mexico (CONACYT), for the financial support given to his PhD studies at DIA-FCQ/UADEC. References 1. Dukare AS, Paul S, Nambi VE, Gupta RK, Singh R, Sharma K, Vishwakarma RK (2019) Exploitation of microbial antagonists for the control of postharvest diseases of fruits: a review. Crit Rev Food Sci Nutr 59:1498–1513 2. Li Y, Xia X, Zhao Q, Dong P (2022) Physiological and molecular plant pathology the biocontrol of potato dry rot by microorganisms and bioactive substances: a review. Physiol Mol Plant Pathol 122:101919 3. European Commission (2023) Farm to fork strategy 4. Rodrigues AO, May De Mio LL, Soccol CR (2023) Trichoderma as a powerful fungal disease control agent for a more sustainable and healthy agriculture: recent studies and molecular insights. Planta 257:1–15 5. Woo SL, Hermosa R, Lorito M, Monte E (2022) Trichoderma: a multipurpose, plantbeneficial microorganism for eco-sustainable agriculture. Nat Rev Microbiol ˜ oz6. Escudero-Leyva E, Alfaro-Vargas P, Mun Arrieta R, Charpentier-Alfaro C, GranadosMontero MDM, Valverde-Madrigal KS, Pe´rez-Villanueva M, Me´ndez-Rivera M, Rodrı´guez-Rodrı´guez CE, Chaverri P, Mora Villalobos JA (2022) Tolerance and biological removal of fungicides by trichoderma species isolated from the endosphere of wild rubiaceae plants. Front Agron 3:1–14 7. Hong-Jun L, Wan-Dong D, Chao L, LingXue M, Hong-Xu L, Rong L, Qi-Rong S (2021) Spore production in the solid-state fermentation of stevia residue by Trichoderma guizhouense and its effects on corn growth. J Integr Agric 20:1147–1156 8. Go´mez-Garcı´a R, Campos DA, Aguilar CN, Madureira AR, Pintado M (2021) Valorisation of food agro-industrial by-products: from the past to the present and perspectives. J Environ Manag 299:113571

9. Yaashikaa PR, Senthil Kumar P, Varjani S (2022) Valorization of agro-industrial wastes for biorefinery process and circular bioeconomy: a critical review. Bioresour Technol 343: 126126 ˜ a-Mendoza SA, Ascacio-Valde´s JA, 10. Saldan Palacios-Ponce AS, Contreras-Esquivel JC, Rodrı´guez-Herrera R, Ruiz HA, Martı´nezHernandez JL, Sugathan S, Aguilar CN (2020) Use of wastes from the tea and coffee industries for the production of cellulases using fungi isolated from the Western Ghats of India. Syst Microbiol Biomanuf 11. Mulatu A, Alemu T, Megersa N, Vetukuri RR (2021) Optimization of culture conditions and production of bio-fungicides from trichoderma species under solid-state fermentation using mathematical modeling. Microorganisms 9 12. Amaya-Chantaca D, Flores-Gallegos AC, Ilina´ A, Aguilar CN, Sepu´lveda-Torre L, Ascacio-Vadle´s JA, Cha´vez-Gonza´lez ML (2022) Comparative extraction study of grape pomace bioactive compounds by submerged and solidstate fermentation. J Chem Technol Biotechnol 97:1494–1505 13. Sala A, Vittone S, Barrena R, Sa´nchez A, Artola A (2021) Scanning agro-industrial wastes as substrates for fungal biopesticide production: use of Beauveria bassiana and Trichoderma harzianum in solid-state fermentation. J Environ Manag 295:113113 14. Sala A, Barrena R, Sa´nchez A, Artola A (2021) Fungal biopesticide production: process scaleup and sequential batch mode operation with Trichoderma harzianum using agro-industrial solid wastes of different biodegradability. Chem Eng J 425:131620 15. Zhang C, Ali Khan RA, Wei H, Wang R, Hou J, Liu T (2022) Rapid and mass production of biopesticide Trichoderma Brev T069 from cassava peels using newly established solid-state

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fermentation bioreactor system. J Environ Manag 313:114981 16. De la Cruz-Quiroz R, Roussos S, HernandezCastillo D, Rodrı´guez-Herrera R, Lo´pezLo´pez LI, Castillo F, Aguilar CN (2017) Solid-state fermentation in a bag bioreactor: effect of corn cob mixed with phytopathogen biomass on spore and cellulase production by trichoderma asperellum. In: Fermentation processes. InTech

17. De la Cruz-Quiroz R, Ascacio-Valde´s JA, Rodrı´guez-Herrera R, Roussos S, Aguilar CN (2019) Phytopathogen biomass as inducer of antifungal compounds by trichoderma asperellum under solid-state fermentation. In: Secondary metabolites of plant growth promoting rhizomicroorganisms. Springer, Singapore, pp 113–124

Chapter 13 Submerged Fermentation as a Strategy for the Valorization of Fish By-products to Obtain High-Protein Meals Anna Marı´a Polanı´a, Alexis Garcı´a, Liliana London˜o-Hernandez, Germa´n Bolivar, and Cristina Ramı´rez Abstract During food processing, a large amount of by-products are generated which, due to their composition, can be used to produce value-added compounds. Particularly the fishing industry generates by-products such as bones, scales, skin, etc., which could be used to produce high commercial value products such as flours. However, one of the major drawbacks is their short shelf life due to their high lipid content. To valorize these by-products, one methodology that can be applied is submerged fermentation (SmF). SmF is a process that involves inoculation of the microbial culture into the liquid medium to produce metabolites or products of interest. The objective of this chapter is to explain in detail how to perform the SmF process on fish by-products, which starter cultures could be applied, how to extract the oil from the paste that is formed, the process to dry it, how to determine the fat content, lipolytic activity in the solid and liquid substrate, the determination of α-amino acid, and degree of protein hydrolysis; finally some aspects that should be considered during the process are mentioned. Key words Fermentation, Fisheries, By-products, Protein, Fish meal

1

Introduction Fisheries and aquaculture are one of the most important sectors worldwide supporting the production of healthy food. According to the FAO report [1], it is estimated that by 2018 world fish production reached about 179 million tons, of which 80 million tons correspond to capture fisheries and the remaining to aquaculture production. Likewise, it is estimated that 156 million tons of the total production is destined for human consumption. During the processing and marketing of fishery products, a large amount of by-products are produced, including head, skeleton, fins, viscera, scales, and shells, among others, which are often improperly handled, causing environmental problems and consequent economic losses. These by-products represent between 30% and 70% of the

Cristobal N. Aguilar Gonzalez et al. (eds.), Food Waste Conversion, Methods and Protocols in Food Science, https://doi.org/10.1007/978-1-0716-3303-8_13, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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weight of the fish, so their production is considerably high [2]. By-products or fishery wastes are mostly composed of protein (15.5–16.3%), fat (5.8–11.1%), fiber (0.2–0.3%) and carbohydrates (2.1–6.9%) [3, 4], so their utilization through different technologies is feasible. Biodiesel [5], polymers [6], feed [7], and fish oil [8], among others, have been obtained using fish wastes. One of the uses of fish waste that has gained greater interest in recent years is as raw material to produce fish meal, to reduce the intensive hunting of fish used for this purpose. However, fish meal from by-products obtained by traditional technologies such as drying and milling is of lower quality and has a short shelf life due to lipid oxidation processes [9]. Therefore, the use of alternative technologies such as fermentation is recommended to produce fish meal. Through fermentation it is possible to improve the quality of proteins, oils, and compounds of fish by-products, besides being considered a clean technology [10]. In general terms fermentation processes can be divided into two: solid-state fermentation and submerged fermentation. In solid-state fermentation (SSF), the process is carried out in a solid substrate without free water but with sufficient water to allow microbial growth. This process is used in a wide variety of agro-industrial wastes for the production of different compounds of interest in different industries such as food, pharmaceutical, and chemical, among others [11]. In the submerged fermentation (SmF) process, inoculation of the microbial culture in liquid medium is necessary to produce desired products. A large part of the products marketed are produced through this process such as antibiotics and enzymes [12]. Often these media are optimized with the use of some nutrients to improve the culture medium. Generally, these microorganisms are grown in closed reactors that have high oxygen concentrations and contain the fermentation medium [13]. It is possible to classify this fermentative process into anaerobic and aerobic. Through the aerobic process, it is possible to produce enzymes and antibiotics, where oxygen is incorporated in the liquid medium, while butanol production is carried out anaerobically, where the entry of oxygen must be avoided. One of the main advantages of this process is its ease of scale-up and automation; in addition, there are no problems related to heat transfer, which is one of the disadvantages of solid-state fermentation. Some of the disadvantages of this process are the complexity of the medium, high production costs, and low productivity [14]. In order to obtain fish meal through submerged fermentation processes, it is important to consider several aspects: the type of microorganism and the concentration of the starter culture, the temperature, oxygen, and relative humidity conditions. Therefore, this chapter describes the methodology used to obtain fish meal through fermentation processes.

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Materials The raw material necessary to obtain high-protein meal can be obtained from by-products obtained from the fish filleting process. Heads, tails, bones, muscle mass, and cartilage from different fish species can be used to this process. The by-products should be frozen (-18 °C) to avoid decomposition during the transport process. However, it is necessary that they be thawed under refrigeration for 24 h before starting the fermentation process. Starters

Some of the strains recommended for the process are Pediococcus acidilactici and Staphylococcus warneri because they present good lipolytic activity. It is also possible to isolate bacteria from the same by-products, but their capacity to synthesize lipase must be identified (see Note 5).

2.2 Fermentation Process

1. Prepare the plastic containers where the process will be carried out; in this case, 21 × 18 cm buckets were used.

2.1

2. Add 1 kg of non-sterilized by-products in each container. 3. Add 10% sucrose and 10% inoculum. 4. In this work, four treatments were carried out to evaluate the effect and evolution of the inoculum on fish by-products through fermentation: T1, control (endogenous bacteria); T2, inoculum with Pediococcus acidilactici bacteria (C20) (8.77 Log CFU/mL); T3, inoculum with lipolytic Staphylococcus warneri bacteria (BL24) (8.63 Log CFU/mL); and T4, mixed inoculum (BL24/C20) (see Note 1) . 5. To create anaerobic conditions, a plastic bag was placed over the by-products, and the empty space of the container was filled with water (Fig. 1). 6. The temperature process was 35 °C, for 4 days, and relative humidity was 83%.

Fig. 1 Arrangement of the components in the fermentation process

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2.3 Reagents of TNBS Method

2,4,6-Trinitrobenzenesulfonic acid (TNBS), sodium sulfite, and Lleucine are reagents necessary to realize the quantification of TNBS method (see Note 2).

2.4

To quantify this parameter is necessary: De Man, Rogosa, and Sharpe (MRS) agar, incubator, cream-agar (CA) culture medium (trypticase soy broth [TSB], 2% agar–agar, and 1% heavy cream [35% milk fat]), and cellulose nitrate filter (25 mM diameter/ 0.2 μM pore size).

3

Lipolytic Activity

Methods

3.1 Paste Oil Extraction

Once the fermentation time its completed, it is necessary to separate the paste from the medium. Three phases are possible to observe: lipidic, aqueous, and solid phase. Use a centrifuge to facilitate the separation process (3500 rpm for 30 min); after this step, oil phase would be present at the surface and could be extracted using a pipette. Aqueous phase is also separated and used to evaluate the degree of hydrolysis obtained. Solids are manually recovered and collected for the drying process.

3.2

Paste obtained after fermentation process have a high moisture content; to assure stability and regulatory requirements is necessary to be dried to 6–10% w.b.

Paste Drying

1. Use a mold to create a thin film (2 mm), and spread it in the drying surface (trays or paper towels could be used as drying surface). 2. A forced convection oven with an air velocity of 1.6 m/s at 60 ° C could be used for the drying process (normally 3 h are required to achieve the desired moisture range under these conditions) (see Note 4). 3. After the process is finished, recover the dried meal for further analysis. 3.3

Fat Content

Fat content is an important quality parameter to evaluate in fish meal; a higher fat content could lead to oxidation reactions and deteriorate of the product. For determination of the fat content: 1. Initially weigh 5 g of fish meal and dry to 95–100 °C for 3 h. 2. Place the fish meal in a filter paper, and wrap it tight (it is possible to use pita thread to firmly adjust the paper around the sample). 3. Put the sample in a Soxhlet extraction system and use ethyl ether solvent for extraction for 16 h.

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4. At the end of extraction period, recover the solvent until volume in the flask is 10–15 mL. Evaporate the solvent at 35 °C and weigh the resulting oil in the flask. 3.4 Lipolytic Activity in a Solid Substrate

1. The bacteria must be activated in De Man, Rogosa, and Sharp (MRS) agar. 2. Subsequently, incubate at 35 °C for 24 h. 3. Cream-agar (CA) culture medium can be used to determine lipolytic activity. 4. A colony at least 48 h old should be selected and seeded in two parallel lines on CA culture medium. 5. Incubate for 72 h at 35 °C. 6. If a halo is observed around the colony, it should be interpreted as a positive lipolysis [15]. 7. To preserve the bacteria, it is recommended to use 30% glycerol at -15 °C. 8. The lipid hydrolysis zone was assessed by impregnating a cellulose nitrate filter with 500 μL of the 24 h culture. 9. It is allowed to dry and overlaid on the center of the agar-cream surface. 10. Plates should be incubated at 35 °C for 72 h. 11. The halo diameter should be measured daily and subtracted from the filter diameter (see Note 6).

3.5 Lipolytic Activity in Liquid Substrate

1. Kinetics should be performed in MRS and TSB broth to evaluate the synthesis of extracellular lipases. 2. It is possible to use 1 L Schott flasks, using 540 mL as working volume, 10% being the inoculum. 3. Before inoculation, the strain must be adapted to the culture media for at least two generations in 10 mL and 54 mL volumes. 4. The media should be incubated at 35 °C for 30 h. To evaluate the lipolytic activity, it is recommended to analyze the amount of free fatty acids every 6 h, following the methodology described by Rai et al. [16] with some modifications: 1. Take 12 mL of the culture medium and centrifuge at 8500 rpm for 15 min. 2. The supernatant should be filtered using No. 2 paper, and then retain the lipid solution.

Whatman

3. Take 1 mL of this lipid solution, and add 1 mL of lipase substrate (fish oil) in 1.5 mL of 0.1 M Tris–HCl buffer at pH 8.0. 4. This mixture is incubated at 37 °C for 15 min.

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5. The hydrolysis should be stopped by adding 3 mL of 95% ethyl alcohol. 6. The mixture should be titrated by adding a 0.01 N NaOH solution and using phenolphthalein as indicator. 7. The blank is determined using the same reaction mixture but without adding the lipid solution. 8. It is recommended to perform the test at least in duplicate. The Equation 1 is used to determine the lipolytic activity: N 1V 1 = N 2V 2

ð1Þ

where N1 = normality of the fatty acids released in the solution (one unit of normality = 103 μM), V1 = volume of the reaction mixture (3.5 mL), N2 = normality of the NaOH solution (0.01 N), and V2 = volume of NaOH required for titration, which equals the difference between the volume required to titrate the sample and the volume required to titrate the blank [17, 18]. To estimate a lipase unit, the amount of lipase required to release 1 μM of fatty acid per minute under assay conditions (U = μM/15 min) can be determined. Enzyme activity should be defined per unit volume (U/mL). The kinetic parameters that can be evaluated are specific growth (μmax), doubling time (td), and % substrate consumption age (% CS). These can be determined by applying Eqs. (2, 3, and 4), respectively: μmax =

ln x - ln x 0 t ln 2 μmax

ð3Þ

ðS 0 - S Þ  100 S0

ð4Þ

td = CS =

ð2Þ

where x is the cell concentration (colony-forming units (CFU)/ mL), x0 is the initial cell concentration, t is the fermentation time to the exponential phase, So is the initial total sugar concentration (g/L), and S is the final total sugar concentration (g/L). 3.6 Determination of α-Amino Acid

1. For the solid fraction obtained from the fermentation process, the α-amino acid content can be determined with the 2,4,6trinitrobenzenesulfonic acid (TNBS) method of Benjakul and Morrisey [19] with some modifications by Ramirez et al. [20]. 2. Samples should be dissolved in 0.2 M phosphate buffer and pH 8 in a 2:3 ratio (silage/buffer). 3. The sample should be thoroughly homogenized and centrifuged at 4000 rpm at 4 °C for 15 min. 4. It is necessary to realize an L-leucine standard curve with concentrations from 0.09 and above of 1.8 mM.

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5. The concentration of α-amino acids can be determined with known concentrations of L-leucine and expressed as mM of α-amino acids. 6. The mixture absorbance was measured at 420 nM in a spectrophotometer (see Note 3). 3.7 Determination of Degree of Protein Hydrolysis

1. The supernatant obtained in item 3 of Subheading 3.5 is used for this determination. Take 64 μL of the supernatant, 1 mL of phosphate buffer (0.2 M, pH 8.2), and 0.5 mL of 0.01% TNBS, and place them in an amber tube with lid. 2. This mixture should be shaken for 10 s and placed in a water bath at 50 °C for 30 min in the dark. 3. The reaction is stopped by adding 1 mL of 0.1 M sodium sulfite. 4. The mixture is left at room temperature for 15 min, and the absorbance at 420 nm should be recorded with a spectrophotometer. The degree of hydrolysis (DH) is calculated using the following equation (Eq. 5): %DH =

ðNH2 Þtx - ðNH2 Þt0  100 ðNH2 ÞHT - ðNH2 Þt0

ð5Þ

where (NH2)tx = amount of α-amino terminal groups at time tx of fermentation, (NH2)t0= amount of α-amino terminal groups in the silage at time 0, and (NH2)HT= amount of α-amino terminal groups in unfermented waste after total acid hydrolysis. Total acid hydrolysis was determined by the method of Baek and Cadwallader [21]: 1. The unfermented sample is dissolved in 0.2 M phosphate buffer and pH 8 in a 2:3 ratio (silage/buffer). 2. This sample is centrifuged at 4000 rpm at 4 °C for 15 min. 3. 0.5 mL of the supernatant is taken and placed in a side release balloon to be mixed with 4.5 mL of 6 N HCl. 4. The atmosphere in the balloon is modified with nitrogen and maintained at 100 °C for 24 h. 5. Subsequently, the reaction mixture is neutralized by the addition of 4.5 mL of 6 N NaOH. 6. The hydrolyzed sample is filtered using Whatman No. 1 paper to remove ash. 7. Finally, the degree of hydrolysis of the filtrate is determined by the TNBS method.

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Notes 1. At treatment (T4), BL24 was added first, and C20 was added 2 days later, until the 4-day fermentation process was completed. 2. Residual sugars in the samples may react with the TNBS reagent, so it is recommended to dilute your sample and then apply the TNBS method [9]. 3. The absorbance obtained from these samples should be subtracted from the absorbance measured for the fish by-product meal sample to avoid overestimating the α-amino acid content [9]. 4. Alternative drying systems such as tunnel belt could be used if a continuous production system is required for dried meal. Refractive window is another interesting system that could assure short process time and retention of the quality of the product [22]. 5. It is recommended that the microorganisms selected for the fermentation process be molecularly identified [23]. 6. It is advisable to perform this assay in triplicate for lipolysispositive bacteria.

References 1. FAO (2020) El estado mundial de la pesca y la acuicultura. https://www.fao.org/3/ca9231 es/ca9231es.pdf. Accessed 20 Aug 2022 2. Wang CH, Doan CT, Nguyen VB, Nguyen AD, Wang SL (2019) Reclamation of fishery processing waste: a mini-review. Molecules 2 4 ( 1 2 ) . h t t p s : // d o i . o r g / 1 0 . 3 3 9 0 / molecules24122234 3. Isibika A, Vinnera˚s B, Kibazohi O, Zurbru¨gg C, Lalander C (2021) Co-composting of banana peel and orange peel waste with fish waste to improve conversion by black soldier fly (Hermetia illucens (L.), Diptera: Stratiomyidae) larvae. J Clean Prod 318. https://doi.org/10.1016/j.jclepro. 2021.128570 4. Coppola D, Lauritano C, Palma Esposito F, Riccio G, Rizzo C, de Pascale D (2021) Fish waste: from problem to valuable resource. Mar Drugs 19(2). https://doi.org/10.3390/ md19020116 5. J. Ching-Velasquez et al., “Production and characterization of biodiesel from oil of fish waste by enzymatic catalysis,” Renew Energy, vol. 153, pp. 1346–1354, Jun. 2020, doi:

https://doi.org/10.1016/j.renene.2020. 02.100 6. S. Mohapatra et al., “Bioconversion of fish solid waste into PHB using Bacillus subtilis based submerged fermentation process,” Environ Technol, vol. 38, no. 24, pp. 3201–3208, Dec. 2017, doi: https://doi.org/10.1080/ 09593330.2017.1291759 7. Tropea A et al (2021) Aquafeed production from fermented fish waste and lemon peel. Fermentation 7(4). https://doi.org/10.3390/ fermentation7040272 8. Inguglia L et al (2020) Salmo salar fish waste oil: fatty acids composition and antibacterial activity. PeerJ 8. https://doi.org/10. 7717/peerj.9299 9. Natalia QP, Cristina RT, Germa´n BE (2022) Lipolytic effect of staphylococcus warneri for obtaining high-quality fishmeal from fish waste fermentation. Waste Biomass Valorization 13: 2519–2530. https://doi.org/10.1007/ s12649-021-01668-8 10. Marti-Quijal FJ, Remize F, Meca G, Ferrer E, Ruiz MJ, Barba FJ (2020) Fermentation in fish and by-products processing: an overview of current research and future prospects. Curr

Submerged Fermentation as a Strategy for the Valorization of Fish. . . Opin Food Sci 31:9–16. https://doi.org/10. 1016/j.cofs.2019.08.001 11. Aita BC et al (2019) Production of cell-wall degrading enzymes by solid-state fermentation using agroindustrial residues as substrates. J Environ Chem Eng 7(3). https://doi.org/10. 1016/j.jece.2019.103193 12. Sharma R, Oberoi HS, Dhillon GS (2016) Fruit and vegetable processing waste. In: Agro-industrial wastes as feedstock for enzyme production. Elsevier, pp 23–59. https://doi. org/10.1016/B978-0-12-802392-1. 00002-2 13. Doriya K, Jose N, Gowda M, Kumar DS (2016) Solid-state fermentation vs submerged fermentation for the production of l-Asparaginase, pp 115–135. https://doi.org/ 10.1016/bs.afnr.2016.05.003 14. Babbar N, Oberoi HS (2014) Enzymes in value addition of agricultural and agro-industrial residues. In: Brar SK, Verma M (eds) Enzymes in value-addition of wastes, pp 29–50 15. Ramı´rez-Lo´pez C, Ve´lez-Ruiz JF (2016) Aislamiento, Caracterizacio´n y Seleccio´n de Bacterias La´cticas Auto´ctonas de Leche y Queso Fresco Artesanal de Cabra. Informacio´n Tecnolo´gica 27(6):115–128. https://doi.org/10. 4067/S0718-07642016000600012 16. Rai AK, Jini R, Swapna HC, Sachindra NM, Bhaskar N, Baskaran V (2011) Application of native lactic acid bacteria (LAB) for fermentative recovery of lipids and proteins from fish processing wastes: bioactivities of fermentation products. J Aquat Food Product Technol 20(1):32–44. https://doi.org/10.1080/ 10498850.2010.528174 17. Jini R et al (2011) Isolation and characterization of potential lactic acid bacteria (LAB) from freshwater fish processing wastes for

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application in fermentative utilisation of fish processing waste. Braz J Microbiol 42(4): 1516–1525. https://doi.org/10.1590/ S1517-838220110004000039 18. Tembhurkar VR, Kulkarni MB, Peshwe SA (2012) Optimization of lipase production by pseudomonas spp. in submerged batch process in shake flask culture. Sci Res Rep 2(1):46–50 19. Benjakul S, Morrissey MT (1997) Protein hydrolysates from Pacific whiting solid wastes. J Agric Food Chem 45(9):3423–3430. https://doi.org/10.1021/jf970294g 20. Ramı´rez JCR et al (2013) Preparation of biological fish silage and its effect on the performance and meat quality characteristics of quails (Coturnix coturnix japonica). Braz Arch Biol Technol 56(6):1002–1010. h t t p s : // d o i . o r g / 1 0 . 1 5 9 0 / S1516-89132013000600016 21. Baek HH, Cadwallader KR (1995) Enzymatic hydrolysis of crayfish processing by-products. J Food Sci 60(5):929–935. https://doi.org/10. 1111/j.1365-2621.1995.tb06264.x 22. Leiton-Ramı´rez, Y.M.; Ayala-Aponte, A.; Ochoa-Martı´nez, C.I. (2020) Physicochemical properties of guava snacks as affected by drying technology. Processes 8, 106. https://doi.org/ 10.3390/pr8010106 23. Sunanta Bunmadee, Jantima Teeka, Thanasak Lomthong, Dolnapa Kaewpa, Prapatsorn Areesirisuk, Atsadawut Areesirisuk (2022) Isolation and identification of a newly isolated lipase-producing bacteria (Acinetobacter baumannii RMUTT3S8-2) from oily wastewater treatment pond in a poultry processing factory and its optimum lipase production, Bioresource Technology Reports, Volume 20, 101267, ISSN 2589-014X, https://doi. org/10.1016/j.biteb.2022.101267

Chapter 14 Monitoring Methods for Anaerobic Digestion of Food Waste: Physicochemical and Molecular Analysis Mario Alberto Yaverino-Gutierrez, Juan Gerardo Flores-Iga, Martha Ine´s Velez-Mercado, Aldo Sosa-Herrera, Marı´a de las Mercedes Esparza-Perusquia, Miriam P. Lue´vanos Escaren˜o, Ayerim Y. Herna´ndez Almanza, Fernando Herna´ndez Tera´n, Javier Ulises Herna´ndez-Beltra´n, and Nagamani Balagurusamy Abstract Anaerobic digestion (AD) is a process in which organic matter such as food waste is degraded and converted into methane, a renewable energy source. AD is a complex biochemical process that occurs in four steps: hydrolysis, acidogenesis, acetogenesis, and methanogenesis. The monitoring of AD is essential to evaluate the performance of a biodigester in order to carry out corrections if needed. Several approaches, such as physicochemical analyses, enzymatic assays, and molecular analyses, enable the whole understanding of this process. Physicochemical analyses and enzyme assays help in understanding the biotransformations that occur during the process and the related microbial activity. In contrast, molecular analyses help to determine the microbiome diversity, dynamics, and various stages of the process and the expression of specific genes and their dynamics and functionality by employing proteomic analyses. Various methods of physicochemical, enzymatic, and molecular analyses are described in this chapter. Key words Anaerobic digestion, Monitoring, Physicochemical, Enzyme, Assay, Metagenomics, Proteomics, Gene expression

1

Introduction The anaerobic digestion (AD) of organic waste is one of the most attractive, cost-effective, and sustainable technologies in waste reduction and a generator of methane, a renewable energy—renewable insofar as using the effluent slurry from the process as an organic fertilizer [1, 2]. AD occurs in four stages: hydrolysis, acidogenesis, acetogenesis, and methanogenesis. Different factors can affect the anaerobic digestion process, such as the presence of inhibitors in waste and undesirable operating conditions, which

Cristobal N. Aguilar Gonzalez et al. (eds.), Food Waste Conversion, Methods and Protocols in Food Science, https://doi.org/10.1007/978-1-0716-3303-8_14, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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Fig. 1 Different techniques employed for monitoring of anaerobic digesters, viz., physicochemical, metagenomics, proteomics, and gene expression to analyse microbial diversity, functions and to identify biomarkers to determine its performance

can result in failures of the biodigester. Hence, continuous monitoring is necessary to identify the state and performance of the process [3, 4]. Psychochemical, enzymatic, and molecular analyses (Fig. 1) can provide a complete idea on this complex biochemical process, which is governed by microbial activities of different groups of microorganisms and the environment inside the biodigester as a consequence of the operating conditions necessary for AD. Chemical oxygen demand (COD), volatile fatty acid (VFA) profile, and methane measurement are some of the physicochemical methods that provide descriptions of the organic matter degradation, the formation of intermediary compounds, and methane yield. Determining hydrolytic enzymes and oxidoreductases measures the rate of conversion and the capability to degrade wastes through the detection of the formation of metabolic intermediates, respectively. Because these processes are carried out by a microbial metabolism, descriptions of the microbial consortium and its functionality are needed. Molecular analyses such as 16S amplicon metagenomics can describe microbial diversity and their dynamics at different stages of the AD process; gene expression analyses, such as on mcrA, a key gene for methane synthesis, can give us glimpses into the activity and functionality of methanogens, which can help us to identify the performance of the biodigester. Similarly, proteomics can help us to identify the protein profiles, through which metabolic pathways, activities, and dynamics can be studied and help to select biomarkers for quick insights into the AD process. In this chapter, we describe the materials and protocols for all the analyses mentioned above, for the monitoring of biodigesters fed with food and/other organic waste.

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Materials

2.1 Physicochemical Analysis 2.1.1 Total Solids (TSs) [5]

1. Evaporating dishes of porcelain, platinum, or high-silica glass (see Note 1). 2. Muffle furnace at 550 °C. 3. Steam bath. 4. Desiccator, with an indicator of moisture concentration. 5. Drying oven, capable of operating at 103–105 °C. 6. Analytical balance, with enough sensitivity for weighting until 0.1 mg. 7. Magnetic stirrer. 8. Pipets (wide bore). 9. Beaker. 10. Graduated cylinder.

2.1.2 Total Suspended Solids (TSSs) at 103–105 °C [5]

(see Note 2).

2.1.3 Fixed Solids and Volatile Solids (FSs and VSs), Ignited at 550 °C [5]

(see Note 2).

2.1.4 Chemical Oxygen Demand (COD) [5]

1. Standard potassium dichromate solution, 0.04167 M. 2. Sulfuric acid–silver sulfate reagent: AgSO4 in a relation 5.5 g/kg H2SO4. 3. Ferroin indicator solution: 1.148 g of 1, 10-phenanthroline monohydrate, 695 mg FeSO4·7H2O, 100 mL of distilled water. 4. Standard ferrous ammonium sulfate (FAS) titrant, approximately 0.25 M. For the preparation of FAS titrant, 0.25 M approximal (see Note 5). 5. Mercuric sulfate, HgSO4, in crystals or powder. 6. Potassium hydrogen phthalate (KHP) standard: 425 mg of KHP, 1 L of distilled water. 7. Reflux apparatus, can be made with an Erlenmeyer flask of 500 or 250 mL, with a ground-glass 24/40 neck and 300 nm jacket Liebig, or by using a condenser adequate to the neck of the ground-glass joint, and for this, a hot plate with enough power to produce a superficial heat of 1.4 W/cm2 is needed. 8. Blender and pipets.

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1. Sodium carbonate solution, approx. 0.05 N. 2. Standard sulfuric acid or hydrochloric acid, 0.1 N. For the preparation of the solution, see Note 4; all steps will be mentioned in that part. 3. Standard sulfuric acid or hydrochloric acid, 0.02 N. 4. Bromocresol green indicator, pH 4.5. 5. Mixed bromocresol green-methyl red indicator solution. This indicator could be used in two forms: an aqueous solution or an alcoholic solution (either of them can be used without affecting the results): 5.1 Aqueous: take 100 mg of bromocresol green sodium salt and 20 mg of methyl red sodium salt and dissolve them into 100 mL of distilled water. 5.2 Alcoholic: same quantities as in aqueous solution but use 100 mL of 95% ethyl alcohol for dissolution. 6. Metacresol purple indicator solution, pH 8.3. 7. Phenolphthalein solution indicator, alcoholic, pH 8.3. 8. Sodium thiosulfate 0.1 N. 9. Electrometric titrator: any commercial pH meter or electrically operated titrator with a glass electrode can be used, but it must be able to measure units of 0.05 pH, and do not forget to calibrate it by following the manufacturer’s instructions. 10. Titration vessel: depending on the size and form of electrodes, vessel should be chosen, and if conventional electrodes are used, the recommended volume is 200 mL. 11. Magnetic stirrer, pipets (volumetric), flask (1000–200– 100 mL), burets (50, 25, 10 mL), and polyolefin bottle of 1 L.

2.1.6 Total Volatile Fatty Acids (Total VFAs) [6]

1. Sulfuric acid, diluted. Mix sulfuric acid and water in equal proportions of volume. 2. Acid ethylene-glycol reagent. Prepare solution as indicated in Note 3. 3. Sodium hydroxide, 4.5 N. 4. Hydroxyammonium sulfate solution, 10%. 5. Hydroxylamine reagent. Take 20 mL of NaOH 4.5 N solution and mix it with 5 mL of hydroxyammonium sulfate before use. 6. Acid ferric chloride reagent. Dissolve 20 g of ferric chloride in 500 mL of distilled water, gently add 20 mL of concentrated sulfuric acid, and set the volume to 1 L. 7. Spectrophotometer UV-visible.

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1. Hydrochloric acid (HCl)—5 M. 2. Orthophosphoric acid. 3. Screw-cap tubes. 4. Ether. 5. Microcentrifuge. 6. Magnesium sulfate (MgSO4), anhydrous. 7. Volatile acid standard solution: 100 mL of distilled water, 0.057 mL of acetic, 0.075 mL of propionic, 0.092 mL of isobutyric, 0.091 mL of butyric, 0.127 mL of isovaleric, 0.125 mL of valeric, 0.126 mL of isocaproic, and 0.126 mL of caproic. Lab supplier. 8. Chromatograph (600 Clarus Perkin Elmer Chromatograph equipped with an Elite-Wax capillary column 30 m × 0.32 mm and a flame ionization detector, or FID). Nitrogen/argon as carrier gas, and hydrogen and oxygen for flame. Oven temperature, 135 °C; injector and detector temperature, 150 °C. 9. Microcentrifuge.

2.1.8 Methane Measurement [7]

1. Methane 99.9%.

2.2 Enzymatic Assays

1. 80% (w/v) ammonium sulfate.

2.2.1

1. 20 mM phosphate buffer, pH 6: 2.5 mg/mL of monosodium phosphate monohydrate, 0.52 mg/mL of disodium phosphate heptahydrate, distilled water.

Amylase [8]

2. Chromatograph (600 Clarus Perkin Elmer Chromatograph equipped with an Elite-Wax capillary column 30 m × 0.32 mm and an FID). Nitrogen/argon as carrier gas, and hydrogen and oxygen for flame. Oven temperature, 70 °C; injector and detector temperature, 90 °C.

2. 3,5-dinitrosalicylic acid (DNS) reagent.

2. 1% (w/v) starch solution in 20 mM phosphate buffer, pH 6. 2.2.2

Exoglucanase [9]

1. 0.1 M citrate buffer, pH 4: 11.5 mg/mL of citric acid, 10.3 mg/mL of trisodium citrate, distilled water. 2. 1% (w/v) crystalline cellulose in 0.1 M of citrate buffer, pH 4.

2.2.3 Endoglucanase [10]

1. 50 mM of phosphate buffer, pH 7.5: 1.3 mg/mL of monosodium phosphate monohydrate, 10.8 mg/mL of disodium phosphate heptahydrate, distilled water. 2. 0.1% (w/v) carboxymethyl cellulose in 50 mM of phosphate buffer, pH 7.5.

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1. 50 mM of phosphate buffer, pH 7.5: 1.3 mg/mL of monosodium phosphate monohydrate, 10.8 mg/mL of disodium phosphate heptahydrate, distilled water. 2. 0.2% (w/v) cellobiose in 50 mM of phosphate buffer, pH 7.5.

2.2.5

Lipase [11]

1. 50 mM of Tris buffer, pH 8.5: 6.06 mg/mL of Tris base, 6 mg/mL of sodium azide, distilled water. 2. 20 mM of p-nitrophenyl palmitate in 50 mM of Tris buffer, pH 8.5. 3. 20% (w/v) sodium dodecyl sulfate (SDS).

2.2.6

Chitinase [12]

1. 0.1 M of phosphate buffer, pH 7: 5.3 mg/mL of monosodium phosphate monohydrate, 16.3 mg/mL of disodium phosphate heptahydrate, distilled water. 2. 0.1% (w/v) colloidal chitin: 66.66 mg/mL of chitin powder in concentrated HCl (see Note 6). 3. Schales’s reagent.

2.2.7

Xylanase [13]

1. 0.1 M of sodium acetate buffer, pH 5: 5.7 mg/mL of sodium acetate, 1.7 mg/mL of acetic acid, distilled water. 2. 2% (w/v) birchwood xylan solution in 0.1 M of sodium acetate buffer, pH 5.

2.2.8

Protease [9]

1. 50 mM of glycine-NaOH buffer, pH 9: 3.8 mg/mL of glycine, 0.4 mg/mL of NaOH, distilled water. 2. 1% (w/v) casein in 50 mM of glycine-NaOH buffer, pH 9. 3. 10% (w/v) trichloroacetic acid (TCA). 4. Folin and Ciocalteu’s phenol reagent. 5. 0.5 M of Na2CO3.

2.2.9

Pullulanase [14, 15 ]

1. 0.1 M of sodium acetate buffer, pH 5: 5.7 mg/mL of sodium acetate, 1.7 mg/mL of acetic acid, distilled water. 2. 1% (w/v) pullulan in 0.1 M of sodium acetate buffer, pH 5.

2.2.10 Benzoyl-CoA Reductase [16]

1. Mops/KOH reaction buffer: 163 mM of Mops/KOH, pH 7.3, 10.87 mM of MgCl2. 2. 0.5 M of ATP. 3. 0.1 M of methyl viologen. 4. 20 mM of Benzoyl-CoA.

2.2.11 Phloroglucinol Reductase [17, 18]

1. Potassium dihydrogen phosphate (KH2PO4) - Ethylenediaminetetraacetic acid (EDTA) reaction buffer: 54.3 mM of KH2PO4, pH 7.2; 1.1 mM of EDTA; 1.1 mM of cysteine chloralhydrate.

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2. 50 mM of phloroglucinol. 3. 10 mM of NADPH. 2.2.12 Resorcinol Reductase [19]

1. KH2PO4 reaction buffer: 108 mM of KH2PO4, pH 7; 1.08 mM of diothioerytrhitol. 2. 40 mM of resorcinol. 3. 40 mM of methyl viologen.

2.2.13 Resorcinol Hydrolase [20]

1. Tris–HCl reaction buffer: 55 mM of Tris–HCl, pH 7.1; 2.7 mM of CaCl2. 2. 40 mM of resorcinol. 3. 40 mM of K3[Fe (CN)6].

2.3

Metagenomics

Materials and solutions must be sterilized and free of nucleases that can degrade DNA, and they can be purchased from a molecular biology reagent supplier. Most reagents are stored at room temperature (-20 °C) unless otherwise indicated. Use an ice bucket to keep the reagents cool.

2.3.1 Sample Preparation [21]

1. Phosphate buffer: 0.12 M, pH 8.

2.3.2

1. Extraction buffer, pH 8: 605.7 mg of Tris–HCl (50 μM), 18.60 mg of EDTA (50 μM), 8.77 g of NaCl, 100 mL of distilled water, adjust pH with HCl.

DNA Extraction [21]

2. 2 mL of microtubes.

2. Snailase solution: 20 mg/mL, Sigma-Aldrich. 3. 10% sodium dodecyl sulfate (SDS) (see Note 7). 4. Lysozyme: 50 mg/mL, Thermo Scientific. 5. Proteinase K: 10 mg/mL, Thermo Scientific. 6. Chloroform/isoamyl alcohol 24:1: Add 48 mL of chloroform in a flask for storage, and then add 2 mL of isoamyl alcohol— for a final volume of 50 mL (see Note 8). 7. Isopropanol, general lab supplier. 8. 70% ethanol, general lab supplier. 2.3.3

16S Amplicon [22]

1. V3-V4 16s rRNA Amplicon PCR Forward Primer (1 μM), suggested: 5’-AYTGGGYDTAAAGNG-3′ (Su et al., 2011, Garcı´a-Lozano et al., 2019). 2. V3-V4 16s rRNA Amplicon PCR Reverse Primer (1 μM), suggested: 5’-TACNVGGGTATCTAATCC-3′ (Su et al., 2011, Garcı´a-Lozano et al., 2019). 3. 2× KAPA HiFi HotStart ReadyMix, KAPA Biosystems.

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4. 10 mM of Tris–HCl, pH 8.5: Add about 20 mL of water to a 100 mL graduated cylinder. Weigh 121.14 mg of Tris base (10 mM) and transfer it to the cylinder. Add water to a volume of 80 mL. Mix and adjust pH with HCl to 8.5. Make up to 100 mL with water. Store at 4 °C. 5. 96-well 0.2 mL PCR plate, Bio-Rad. 6. AMPure XP beads, Beckman Coulter. 7. 80% ethanol. 2.3.4 [22]

Library Preparation

1. 2× KAPA HiFi HotStart ReadyMix, KAPA Biosystems. 2. Nextera XT Index 1 Primers (N7XX) from the Nextera XT Index kit, Illumina. 3. Nextera XT Index 2 Primers (S5XX) from the Nextera XT Index kit, Illumina. 4. PCR-grade water, general lab supplier. 5. TruSeq Index Plate Fixture, Illumina. 6. 96-well 0.2 mL PCR plate, Bio-Rad. 7. Microseal ‘A’ film, Bio-Rad. 8. 10 mM of Tris–HCl, pH 8.5: Add about 20 mL of water to a 100 mL graduated cylinder. Weigh 121.14 mg of Tris base (10 mM) and transfer to the cylinder. Add water to a volume of 80 mL. Mix and adjust pH with HCl to 8.5. Make up to 100 mL with water. Store at 4 °C. 9. AMPure XP beads, Beckman Coulter. 10. 80% ethanol.

2.3.5 Sequencing Preparation [22]

1. HT1 Hybridization Buffer, Illumina. 2. 0.2 N NaOH solution: Add about 50 mL of water to a 100 mL graduated cylinder. Weigh 0.8 g of NaOH (0.2 M) and transfer it to the cylinder. Mix until dissolved, and make up to 100 mL with water. Store at room temperature. 3. PhiX Control Kit v3, Illumina. 4. MiSeq reagent cartridge, Illumina.

Equipment

1. Centrifuge. 2. Vortex. 3. Water bath. 4. Ice cube. 5. Thermal cycler. 6. Magnetic stand.

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7. UV-vis spectrophotometer/nanodrop spectrophotometer. 8. MiSeq System, Illumina. 2.4

Gene Expression

2.4.1 Sample Preparation

2.4.2 RNA Extraction [23, 24 ]

All the materials, solutions, and surfaces used must be RNAse-free and sterile to prevent RNA degradation, and the reagents can be purchased from any molecular biology supplier. Solutions are normally stored at -20 °C unless otherwise specified. Use an ice bucket to keep the reagents cool. Lysis buffer Tris-EDTA-Sodium dodecyl sulfate (TES): 10 mM of Tris–HCl, pH 7.5; 10 mM of EDTA; and 0.5% sodium dodecyl sulfate (SDS). 1. Stock solution of Tris–HCl (1 M): Dissolve 121.1 g of Tris base in 800 mL of H2O, and after dissolving it, make it up to 1 L. Take an aliquot and adjust it to the required pH with concentrated HCl. 2. Stock solution EDTA (0.5 M): Dissolve 186.1 g of EDTA, and make up to 1 L. Take an aliquot and adjust it to the required pH. 3. Acid saturated phenol solution (pH 4.3): Liquify 100 g of phenol crystal at 60 °C. Add 100 mL of liquified phenol to a beaker, mix it with an equal volume of DEPC-treated water, and wait until a thin aqueous layer appears on the surface. Transfer the down layer to a beaker and wash it multiple times with citrate buffer (0.1 M, pH 4.3). Take the down layer and use it or store it at 20 °C (see Note 9 and 10). 4. Saturated phenol: Liquify 100 g of phenol crystal at 60 °C. Add 100 mL of liquified phenol to a beaker, mix it with an equal volume of DEPC-treated water, and wait until a thin aqueous layer appears on the surface. Transfer the down layer to a beaker and store it at -20 °C until ready for use. 5. Chloroform 99%. 6. Sodium acetate (3 M): Weigh 24.61 g of sodium acetate dihydrate and transfer it to an volumetric flask (100 ml) with 40–50 ml of distilled water. After the salt is dissolved, the volume of the flask is made up to 100 mL with water. 7. Ethanol 100%. 8. Ethanol 70%. 9. Tris-EDTA (TE) buffer (pH 8): Add about 20 mL of water in a 100 mL volumetric flask. Take 1 mL of stock solution Tris– HCl (1 M, pH 8) and 0.2 mL of stock solution EDTA (0.5 M, pH 8), then transfer them to the solution flask.

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2.4.3 qRT-PCR for Relative Expression of mcrA Gene [25, 26]

1. Primer qmcrA-F (forward) 10 μM, TTCGGTGGATCDCARAGRGC3’ (36).

suggested:

5’

2. Primer mcrA-rev (reverse) 10 μM, suggested: CGTTCATBGCGTAGTTVGGRTAGT 3′ (37).

5’

3. RT-PCR-quality H2O. 4. SYBR Green qPCR Master Mix, Thermo Fisher. 5. iScript cDNA synthesis kit, Bio-Rad Laboratories, Inc. 6. cDNA. 7. 96-well 0.2 mL PCR plate, Bio-Rad. 2.4.4

Equipment

1. Refrigerated centrifuge. 2. Vortex. 3. Water bath. 4. UV-vis spectrophotometer. 5. Thermal cycler. 6. qPCR thermal cycler.

2.5 2.5.1

Proteomics SDS-Page [27]

1. Lysis buffer: 3% (v/v) β-Mercaptoethanol, 0.83 mg/mL of bromophenol blue, 7.6 mg/mL of Trizma, 10 mg/mL of SDS, 3% (v/v) glycerol, distilled water. 2. 0.1 M of phosphate buffer, pH 7.5: 2.3 mg/mL of monosodium phosphate monohydrate, 22.3 mg/mL of disodium phosphate heptahydrate, distilled water. 3. 30% (w/v) acrylamide–bis acrylamide: 29% (v/v) acrylamide, 1% (v/v) bis acrylamide. 4. 1.5 M of Tris–HCl, pH 8.8: 40 mg/mL of SDS, 182 mg/mL of Trizma base, distilled water (see Note 11). 5. 10% (w/v) sodium dodecyl sulfate (SDS). 6. 10% (w/v) ammonium persulfate (APS) (see Note 12). 7. -N, N, N′, N′-tetramethylethylenediamine (TEMED). 8. 0.5 M of Tris–HCl, pH 6.8: 40 mg/mL of SDS, 60.4 mg/mL of Trizma, 50 mL of distilled water (see Note 11). 9. Running buffer 5×: 75 mg/mL of glycine, 15.1 mg/mL of Trizma base, 5 mg/mL of SDS. 10. Coomassie stain solution: 0.02% (w/v) Coomassie, 5% (w/v) aluminum sulfate, 10% (w/v) ethanol, 2% (w/v) orthophosphoric acid. 11. Distaining solution: 30% methanol, 5% acetic acid, 65% milli-Q water.

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2D-Page [28]

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1. Immobilized pH gradient (IPG) strip. 2. Rehydration buffer: 7 M of urea, 2 M of thiourea, 1% ASB-14, 40 mM of Tris, 0.001% bromophenol blue. 3. Mineral oil. 4. Equilibration buffer I: 6 M of urea, 0.375 M of Tris–HCl, pH 8.8, 2% SDS, 20% glycerol, 2% (w/v) Dithiothreitol (DTT). 5. Equilibration buffer II: 6 M of urea, 0.375 M of Tris–HCl, pH 8.8, 2% SDS, 20% glycerol, 2.5% (w/v) iodoacetamide. 6. Overlay agarose: 0.5% low melting point agarose in 1× Tris/ glycine/SDS and 0.003% bromophenol blue.

2.5.3 Mass Spectrometry [29, 30]

1. 100 mM of NH4HCO3 and acetonitrile: 7.9 mg/mL of NH4HCO3, 4.1 mg/mL of acetonitrile, distilled water. 2. Digestion buffer: 50 mM of NH4HCO3, 5 mM of CaCI2, and 12.5 ng/μL of trypsin. 3. 5% (w/v) formic acid and acetonitrile. 4. 5%–40% acetonitrile, 0.1% formic acid.

2.5.4

Equipment

1. Protein electrophoresis equipment (SDS-PAGE, 2D-PAGE, IEF). 2. Spectrophotometer. 3. LC-MSMS/MS.

3 3.1 3.1.1

Methods Physicochemical Total Solids [5]

1. The preparation of the evaporating dish depends on the analysis type. Total solids or volatile solids require a range of temperatures, but this range varies depending on which analysis will be conducted (see Note 13). 2. Sample a volume with a yield residue in the range of 2.5–200 mg, which must have been previously identified. 3. Collect a well-mixed sample and measure its volume in a previously weighed dish. If the sample is homogeneous, it must be taken at the midpoint, not at the bottom of the flask or container of the sample. At the same time, the sample must not be taken near the vortex, so the sample must be taken at the midpoint, between the wall and the vortex, to guarantee the dryness of the sample. 4. Take a well-mixed sample and measure its volume in a previously weighed dish. For a homogeneous sample, take it at the midpoint of the vessel, not near the vortex, to guarantee the dryness of the sample.

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5. Take the sample and put it into a drying oven to eliminate the moisture. While in transit, stir the sample with a magnetic stirrer. If necessary, successive portions of the sample must be added to the same dish just after evaporation, and be careful to avoid splattering the sample in the evaporation, adjusting the temperature to approximately 2 °C lower than its boiling point when evaporating. 6. Dry the evaporated sample in the oven at a temperature of 103–105 °C for 1 h. 7. Put the dish into a desiccator to balance the temperature, and finally weigh the dish. These steps of drying, desiccating, and cooling must be repeated until a constant weight has been maintained (see Note 14). Calculation: mg total solids=L =

ðA - B Þ  1000 mL ðsample volumeÞ:

where A = dried residue + dish (weight of both in mg) and B = weight of dish (mg). 3.1.2 Total Suspended Solids (TSS) at 103–105 °C [5]

1. If preprepared fiber filter disks are already available, please jump ahead to step 6, but if not, please continue in step 2. 2. Take a disk and put it with the creased side up on the filtration apparatus, then apply vacuum and wash them with 20 mL of reagent-grade water. The washing should be repeated three times. 3. Remove all traces of water by using suction, turn off the vacuum machine, and throw out the washing; then remove the filter from the filtration machine, and transfer it into an aluminum weighing dish. If a Gooch crucible is used, remove the combination between the filter and the crucible. Dry it in an oven at 103–105 °C for 1 h; if volatile solids are going to be measured, ignite it at 550 °C for at least 15 min in a muffle furnace. 4. Take the disk and put it into a desiccator until a balance between temperature and weight has been reached; these last steps of igniting, cooling, desiccating, and weighing must be repeated until a weight that does not change by more than 4% from its previous weight has been reached. Lastly, store the disk in a desiccator. 5. Select a sample, taking a sample volume with a yield in the range of 2.5–200 mg of dried residue. If after it has been filtered, it still does not have enough yield, the volume of the sample must be increased until it reaches 1 L. If a filtration time

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of more than 10 min is required, then either the volume should be decreased or the filter diameter should be increased. 6. Use an arm-filtering machine, then filter the sample and start with suction. Moisten the filter with distilled water to obtain a better seat; also stir the sample with a magnetic stirrer, and speed it up until it has uniform particles (almost homogenous). This strategy is more practical. Centrifuge is not a good option, because centrifugal force can separate by size and density, which means that it has a poor accuracy because the samples have different withdrawal points. 7. Place a known volume onto the seated glass-fiber filter; at the time of being stirred, if the sample is homogenous, ensure that the volume is extracted from the midpoint of the container, not the vortex. More specifically, it should be taken between the wall and the vortex, at the middle of the container. Then wash it with distilled water. Wash the filter three times, using a volume of 10 mL. Between washings, allow the filter to drain, and then continue with suction for at least 3 min just after filtration has ended (some samples with high content of dissolved solids may require additional washings). 8. Be careful when taking away the filter from the filtration machine, and move the filter onto an aluminum weighing disk as support. If a Gooch crucible is used, remove the crucible and filter combination from the crucible adapter. 9. Dry for at least 1 h between 103 °C and 105 °C in an oven, then put it into a desiccator and weigh it. Continue to repeat these last steps (of drying, desiccating, and weighing) until a constant weight has been reached and maintained (see Note 14). Calculation: mg total suspended solids=L =

ðA - B Þ  1000 mL ðsample volumeÞ

where A = dried residue + filter (weight of both in mg) and B = weight of filter (mg) (see Note 15). 3.1.3 Fixed Solids and Volatile Solids (FSs and VSs), Ignited at 550 °C [5]

1. Residue from total solids assays is recollected and then ignited at 550 °C in a muffle furnace until it reaches a constant weight (see Note 13). 2. Ignite a blank glass-fiber filter with samples at the same time. Before inserting the samples, heat the furnace up to temperature. A recommended sample of 200 mg of residue takes 15–20 min to complete ignition. Nevertheless, more than one sample, or samples with bigger weights than 200 mg, of

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residue may take longer for ignition because the furnace could be overtaxed. 3. Cool the dish or filter the disk at environmental temperature for a considerable time until most of the heat has gone, then put it into a desiccator. The desiccator must not be overburdened. Finally, weight of filter the disk (after it has cooled, to balance the temperature inside the desiccator). 4. Repeat the steps of igniting, cooling, and weighting until a constant weight has been reached or until the weight is not more than 4% of the previous weight. Also, if the determination is in duplicate samples, the average weight must agree within 5%. If the blank filter lost weight, this is a clear signal that this type of filter is inadequate for this assay. Calculation: mg volatile solids=L = mg fixed solids=L =

ðA - B Þ  1000 mL ðsample volumeÞ

ðB - C Þ  1000 mL ðsample volumeÞ

where A = residue + dish before ignition (weight, mg), B = residue + dish after ignition (weight, mg), and C = filter/or dish it was used (weight, mg) (see Note 16). 3.1.4 Chemical Oxygen Demand (COD) [5]

1. The procedure changes depending on organic matter and O2 concentration. To learn more, see Note 20. If the samples have a chemical oxygen demand (COD) greater than 50 mg of O2/ L, please proceed to step 2; otherwise, proceed to step 8. 2. Mix the sample if required, and pipet 50 mL into a 500 mL refluxing flask (for a sample with a COD of 900 mg of O2/L, take a smaller portion of the sample and dilute it to 50 mL; see Note 21). 3. Carefully and slowly add 1 g of HgSO4 and 5 mL of acid sulfuric reagent to dissolve the HgSO4. During the time of mixing, cool it to avoid losing volatile materials. Next, add 25 mL of 0.04167 M of K2Cr2O7 solution and mix. The flask must be attached to the condenser to gain better control, then place it into cooling water. 4. Add the rest of the volume (70 mL) with an acid sulfuric reagent through the condenser open part (the reflux mix must be well mixed before applying heat, to avoid heating the bottom of the flask, because otherwise, a blast of flask content may occur). 5. Cover the open-end part of the condenser with a small beaker to prevent external material from entering the condenser and thus affecting the refluxing mixture. Then reflux for 2 h.

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6. Wash down the condenser and cool it with distilled water, then disconnect the next condenser and dilute the mixture with distilled water until it reaches double the original volume. Cool the mixture to room temperature and titrate the excess of K2Cr2O7 with FAS titrant (approximately 10–15 mL is required: 2 or 3 drops). Even though the amount of ferroin indicator is not crucial, use the same volume in all titrations, taking the final point of titration. The first change of color from a blue-green to reddish-brown, even if the change occurs for only a minute or so, is enough to be considered a final point. 7. In duplicate determinations, their average variation must be lower than 5%. Samples that have components such as suspended solids or other components may interfere with oxidation, so additional determinations may be needed. The bluegreen coloration could appear again, so it would be convenient to reflux and titrate a blank that contains the reagents of the sample with the same volume of distilled water. 8. In an alternative procedure for low-COD samples, all the steps of the procedure, from step 1 to 7, are the same, with some exceptions—using the standard 0.004167 M K2Cr2O7 and FAS 0.025 M titrant (different concentrations with respect to the one used before). These procedures require taking precautions because even a small amount of organic matter in or on the glassware, such as from the atmosphere, could cause enormous errors. If more precision and more sensitivity are required, the best option is to take a larger volume of the sample for reflux, and follow the steps as mentioned earlier (see Note 22 for more information on this alternative procedure). Calculation: COD, mg O2 =L =

ðA - B Þ  M  8000 mL ðsample volumeÞ

where A = mL of FAS used for blank, B = mL of FAS used for the sample, M = molarity of FAS (approximately in 0.25 M), and 8000 = the milliequivalent weight of oxygen. 3.1.5

Alkalinity [5]

Color Change

*For the alkalinity essay, some parts are for different stages of the method; these parts are indicated before the starting instructions. 1. Take a known volume of the sample and write down the normality from the titrant. Care should be taken that the sample is brought to room temperature before analysis. 2. Take the sample up in a pipette and discharge it into an Erlenmeyer flask. If the sample has some chlorine present, add

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0.05 mL/1 drop of 0.1 M of NaS2O3 solution. An alternative is to destroy the chlorine with UV radiation. 3. Add five drops/0.2 mL of indicator solution, and then titrate over the surface that has a white color until a color change is maintained, depending on the indicator and its equivalent point. Check whether commercial indicator solutions can set the pH to values from 3.7 to 8.3, and check the color and endpoint by adding the same concentration of indicator used with the sample (see Note 18) and by using a buffer for the required pH. Potentiometric Titration Curve

1. For the potentiometric titration curve, first select one titration vessel and wash it out, then wash out the electrodes and the drain. Select the sample known volume and the normality of titrant solution on the basis of the criteria. Adjust it to an environmental temperature; after that, take up the sample in a pipette, and put it into the vessel. 2. Continue to measure the sample pH, and add the standard acid solution in portions of 0.5 mL or even less, such that the pH will change an estimated 0.2 pH units per portion added; every time that standard acid solution is added, mix with a magnetic stirrer; avoid losing any volume from the mixing. Keep adding titrant solution until a pH of 4.5 is reached. 3. Construct a titration curve by plotting observed pH values and comparing them against the value of the total volume (mL) of the titrant used. It is important to consider that a smooth curve shows few points (one or more), which are known as inflections; on the other hand, an erratic curve might indicate that an equilibrium never existed between successive acid additions. It is recommended to determine an alkali relative to a particular pH by using another curve.

Potentiometric Titration to Preselected pH

1. For potentiometric titration, to preselect the pH, adhere to the following instructions: The determinate depends on the endpoint (concentration of carbon dioxide; see Note 18), as in the before steps (potentiometric curve titration). Prepare the sample and all equipment for titration; titrate until the endpoint pH has been reached. For this test, ignore the intermediate pH, and avoid delays. 2. Make some smaller additions of acid standard solution; once that endpoint pH has been obtained, check that pH equilibrium has been achieved before adding more titrant.

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1. Before starting, please see Note 19 to learn about low alkalinity and a what is occurring in this part. 2. Titrate 100–200 mL by following the same procedure as that for preselecting the pH for potentiometer titration, but instead, use a 10 mL microburet and 0.02 N standard acid solution because they are necessary to stop titration in the range of 4.3–4.7 pH. Note the pH and the volume of titrant used. Then add titrant to reduce the pH to exactly 0.3 pH, and note the volume. Calculations

For Potentiometric Titration to Endpoint pH

Alkalinity, mg CaCO3 =L =

A  N  50 000 mL of sample

where A = mL standard acid used and N = normality of standard acid. Another equation to determine alkalinity is Alkalinity, mg CaCO3 =L =

A t 1000 mL of sample

where A = the same as in the previous equation and t = titer of standard acid, mg CaCo3/mL. For Potentiometric Titration of Low Alkalinity

Alkalinity, mg CaCO3 =L =

ð2B‐CÞ  N 50 000 mL of sample

where B = mL used of titrant to first recorded pH, C = total mL titrant to reduce 0.3 pH unit, and N = normality of acid. 3.1.6 Total Volatile Fatty Acids (VFAs) [6]

1. First, use a filter aid to filter the sample and thus obtain more clarity; in some cases, clarification is obtained by spinning the sample in a centrifuge. 2. Take 0.5 mL of the liquor into one dry test tube, add 1.7 mL of acidic ethylene-glycol reagent, and mix from top to bottom (an alternative is to add 1.5 of ethylene-glycol reagent and 0.2 mL of diluted sulfuric acid). 3. Heat the tube in a boiling water bath for at least 3 min, avoiding direct contact between the tube and the boiling water container. After that, cool the tube by using water, add 2.5 mL of hydroxylamine reagent (another way adds 0.5 mL of hydroxylammonium sulfate solution and 2 mL of NaOH solution at 4.5 N), and mix. 4. Take 10 mL of acid ferric chloride reagent and put it into a 25 mL volumetric flask, add solution from the tube, fill the flask to the mark with water, shake the flask, and let rest for 5 min without the top of the flask— this way, dissolved gasses escape

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more easily. If it is expected that the volatile acid concentration will exceed 500 ppm, put the tube away for at least 1 min. 5. Measure OD (optical density) at 500 nm. Bubbles and gas in the optical cell can affect the determination. Take the reading at the first sign of color development. Discount the blank value that is obtained with 0.5 mL of distilled water. 6. The results must be verified with a previous graphic of acetic acid. Make sure that the organic acid concentration of a previous graph is prepared with solutions of pure acetic acid (see Note 17). 3.1.7 Volatile Fatty Acids Profile and Quantification [7]

1. Take 8 mL of the sample growing culture from the biodigester, and add 0.1–0.2 mL of hydrochloric acid (HCl) or orthophosphoric acid (H3PO4) to acidify the sample to a pH of 2 or below. 2. Take 4 mL of acidified sample, and put it into a screw-cap tube; add 1 mL of ether, gently invert the tube for mixing for 20 min, and then centrifuge at 2000 rpm for 5 min. 3. Put the tube in a freezer at 10 °C or lower until the bottom phase (liquid fraction) is frozen. Remove the top portion, and put the ether portion into a screw-cap tube. Then add magnesium sulfate-anhydrous (MgSO4) until it is equal to half the ether portion’s volume, and let it sit for 10 min. 4. Inject 1 μL into the column of the chromatograph, where the chromatograph is fitted with 20 M of 3% Carbomax column and flame ionization detector (FID), under the conditions mentioned earlier [31]. 5. To identify and quantity the volatile acids of the sample, compare the elution times and the area of the peaks obtained with known standard acids’ chromatographic runs.

3.1.8 Methane Measurement [7]

1. By using a gas-tight syringe, inject 1 μL of the sample into the chromatograph. 2. The column and the conditions for measurement were mentioned previously. 3. To calculate the methane concentration, prepare a calibration curve at different concentrations, using methane at 99.9% as the standard sample.

3.1.9 Enzymatic Assays Amylase [8]

1. Take 1.5 mL of sample and centrifuge at 10,000 rpm for 10 min. 2. Precipitate the proteins on supernatant with 80% (w/v) ammonium sulfate. 3. Centrifuge again and resuspend the precipitates in 20 mM of sodium phosphate buffer, pH 6 (see Note 23).

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4. Add 125 μL of the resuspended precipitates and 125 μL of 1% starch solution prepared in 20 mM of sodium phosphate buffer, pH 6 (see Note 24). 5. Incubate for 10 min at 45 °C (see Note 25). 6. Stop the reaction by adding 250 μL of DNS reagent and boiling for 5 min. 7. Let the samples cool, and add 2.5 mL of distillate water. 8. Measure the released reducing sugars at 540 nm. 9. One unit of amylase activity was determined as the amount of enzyme required to release 1 μmol of reducing sugars as glucose equivalent per mL in 1 min (U mL-1/min-1). 3.1.10

Exoglucanase [9]

1. Take 1.5 mL of sample and centrifuge at 10,000 rpm for 10 min. 2. Precipitate the proteins on supernatant with 80% (w/v) ammonium sulfate. 3. Centrifuge again and resuspend the precipitates, and resuspend them in 0.1 M of citrate buffer, pH 4 (see Note 23). 4. Add 125 μL of the resuspended precipitates and 125 μL of 1% (w/v) crystalline cellulose in 0.1 M of citrate buffer, pH 4 (see Note 24). 5. Incubate for 60 min at 60 °C (see Note 25). 6. Stop the reaction by adding 250 μL of DNS reagent and boiling for 5 min. 7. Let the samples cool, and add 2.5 mL of distillate water. 8. Measure the reducing sugars released at 540 nm. 9. One unit of exoglucanase activity was determined as the amount of enzyme required to release 1 μmol of reducing sugars as glucose equivalent per mL in 1 min (U mL-1/min-1).

3.1.11 Endoglucanase [10]

1. Take 1.5 mL of sample and centrifuge at 10,000 rpm for 10 min. 2. Precipitate the proteins on supernatant with 80% (w/v) ammonium sulfate. 3. Centrifuge again and resuspend the precipitates, and resuspend them in 0.05 M of phosphate buffer, pH 7.5 (see Note 23). 4. Add 125 μL of the resuspended precipitates and 125 μL of 0.1% (w/v) carboxymethyl cellulose (CMC) in 0.05 M of phosphate buffer, pH 7.5 (see Note 24). 5. Incubate for 60 min at 60 °C (see Note 25). 6. Stop the reaction by adding 250 μL of DNS reagent and boiling for 5 min.

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7. Let the samples cool, and measure the reducing sugars released at 540 nm. 8. One unit of endoglucanase activity was determined as the amount of enzyme required to release 1 μmol of reducing sugars as glucose equivalent per mL in 1 min (U mL-1/min-1). 3.1.12

Cellobiase [10]

1. Take 1.5 mL of sample and centrifuge at 10,000 rpm for 10 min. 2. Precipitate the proteins on supernatant with 80% (w/v) ammonium sulfate. 3. Centrifuge again and resuspend the precipitates, and resuspend them in 0.05 M of phosphate buffer, pH 7.5 (see Note 23). 4. Add 125 μL of the resuspended precipitates and 125 μL of 0.2% (w/v) cellobiose in 0.05 M of phosphate buffer, pH 7.5 (see Note 24). 5. Incubate for 60 min at 60 °C (see Note 25). 6. Stop the reaction by adding 250 μL of DNS reagent and boiling for 5 min. 7. Let the samples cool, and add 2.5 mL of distillate water. 8. Measure the reducing sugars released at 540 nm. 9. One unit of β-glucosidase activity was determined as the amount of enzyme required to release 1 μmol of reducing sugars as glucose equivalent per mL in 1 min (U mL-1/min-1).

3.1.13

Lipase [11]

1. Take 1.5 mL of sample and centrifuge at 10,000 rpm for 10 min. 2. Precipitate the proteins on supernatant with 80% (w/v) ammonium sulfate. 3. Centrifuge again and resuspend the precipitates, and resuspend them in 50 mM of Tris buffer, pH 8.5 (see Note 23). 4. Add 50 μL of resuspended precipitates and 75 μL of 20 mM p-nitrophenyl palmitate (p-NPP) in 50 mM of Tris buffer, pH 8.5 (see Note 24). 5. Incubate for 20 min at 45 °C (see Note 25). 6. Stop the reaction by adding 20 μL of 20% (w/v) SDS. 7. Measure the released p-nitrophenols at 410 nm. 8. One unit of lipase activity was determined as the amount of enzyme required to release 1 μmol of p-nitrophenol per mL in 1 min (U mL-1/min-1).

3.1.14

Chitinase [12]

1. Take 1.5 mL of sample and centrifuge at 10,000 rpm for 10 min.

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2. Precipitate the proteins on supernatant with 80% (w/v) ammonium sulfate. 3. Centrifuge again and resuspend the precipitates, and resuspend them in 0.1 M of phosphate buffer, pH 7 (see Note 23). 4. Add 300 μL of the resuspended precipitates and 300 μL of 0.1% (w/v) colloidal chitin (see Note 24). 5. Incubate for 10 min at 55 °C (see Note 25). 6. After incubation, centrifuge the samples at 10,000 rpm for 5 min. 7. Take 200 μL of the supernatant with 500 mL of distilled water and 1 L of Schales’s reagent, boiling for 10 min. 8. Let the samples cool, and measure the reducing sugars released at 420 nm. 9. One unit of the chitinase activity was determined as the amount of enzyme required to release 1 μmol of reducing sugar as N-acetyl-D-glucosamine (GlcNAc) equivalent per mL in 1 min (U mL-1/min-1). 3.1.15

Xylanase [13]

1. Take 1.5 mL of sample and centrifuge at 10,000 rpm for 10 min. 2. Precipitate the proteins on supernatant with 80% (w/v) ammonium sulfate. 3. Centrifuge again and resuspend the precipitates, and resuspend them in 0.1 M of sodium acetate buffer, pH 5 (see Note 23). 4. Add 125 μL of resuspended precipitates and 125 μL of 2% (w/v) birchwood xylan solution in 100 mM of sodium acetate buffer, pH 5 (see Note 24). 5. Incubate for 5 min at 50 °C (see Note 25). 6. Stop the reaction by adding 250 μL of DNS reagent and boiling for 5 min. 7. Let the samples cool, and add 2.5 mL of distillate water. 8. Measure the reducing sugars released at 540 nm. 9. One unit of xylanase activity was determined as the amount of enzyme required to release 1 μmol of reducing sugars as xylose equivalent per mL in 1 min (U mL-1/min-1).

3.1.16

Protease [9]

1. Take 1.5 mL of sample and centrifuge at 10,000 rpm for 10 min. 2. Precipitate the proteins on supernatant with 80% (w/v) ammonium sulfate. 3. Centrifuge again and resuspend the precipitates, and resuspend them in 50 mM of glycine-NaOH buffer, pH 9 (see Note 23).

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4. Add 250 μL of the resuspended precipitates and 250 μL of 1% (w/v) casein in 50 mM of glycine-NaOH buffer, pH 9 (see Note 24). 5. Incubate for 10 min at 60 °C (see Note 25). 6. Stop the reaction by adding 125 μL of 10% (w/v) trichloroacetic acid (TCA). 7. After 20 min, centrifuge the samples at 10,000 g, 4 °C for 10 min. 8. Add 125 μL of 0.5 M Na2CO3 to 250 μL of the supernatant. 9. Keep the samples in dark conditions, and add 125 μL of Folin and Ciocalteu’s phenol reagent. 10. Measure the tyrosine released at 660 nm. 11. One unit of protease activity was determined as the amount of enzyme required to release 1 μmol of tyrosine per mL in 1 min (U mL-1/min-1). 3.1.17 Pullulanases [14, 15 ]

1. Take 1.5 mL of sample and centrifuge at 10,000 rpm for 10 min. 2. Precipitate the proteins on supernatant with 80% (w/v) ammonium sulfate. 3. Centrifuge again and resuspend the precipitates, and resuspend them in 0.1 M of sodium acetate buffer, pH 5 (see Note 23). 4. Add 125 μL of resuspended precipitates and 125 μL of 1% (w/v) pullulan in 0.1 M of sodium acetate buffer, pH 5 (see Note 24). 5. Incubate for 30 min at 60 °C (see Note 25). 6. Stop the reaction by adding 250 μL of DNS reagent and boiling for 5 min. 7. Let the samples cool, and add 2.5 mL of distillate water. 8. Measure the reducing sugars released at 540 nm. 9. One unit of pullulanase activity was determined as the amount of enzyme required to release 1 μmol of reducing sugars as glucose equivalent per mL in 1 min (U mL-1/min-1).

3.1.18 Benzoyl-CoA Reductase [16]

1. Take 1.5 mL of sample and centrifuge at 10,000 rpm for 10 min. 2. Precipitate the proteins on supernatant with 80% (w/v) ammonium sulfate. 3. Centrifuge again and resuspend the precipitates, and resuspend them in 460 μL of Mops/KOH reaction buffer (see Note 23). 4. Add 5 μL of ATP, 5 μL of 20 mM of benzoyl-CoA, and 5 μL of 0.1 M of methyl viologen to the resuspended precipitates.

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5. Incubate for 10 min at 37 °C (see Note 25) under anaerobic conditions (see Note 26). 6. Measure methyl viologen oxidation at 730 nm. 7. One unit of benzoyl-CoA reductase was determined as the amount of enzyme required to oxidate 1 μmol of methyl viologen per mL in 1 min (U mL-1/min-1). 3.1.19 Phloroglucinol Reductase [17, 18]

1. Take 1.5 mL of sample and centrifuge at 10,000 rpm for 10 min. 2. Precipitate the proteins on supernatant with 80% (w/v) ammonium sulfate. 3. Centrifuge again and resuspend the precipitates, and resuspend them in 460 μL of KH2PO4-EDTA reaction buffer (see Note 23). 4. Add 10 μL 50 mM phloroglucinol and 5 μL 10 mM NADPH to the resuspended precipitates. 5. Incubate for 10 min at 30 °C (see Note 25). 6. Measure the dihydrophloroglucinol released at 730 nm. 7. One unit of phloroglucinol reductase was determined as the amount of enzyme required to release 1 μmol of dihydrophloroglucinol per mL in 1 min (U mL-1/min-1).

3.1.20 Resorcinol Reductase [19]

1. Take 1.5 mL of sample and centrifuge at 10,000 rpm for 10 min. 2. Precipitate the proteins on supernatant with 80% (w/v) ammonium sulfate. 3. Centrifuge again and resuspend the precipitates, and resuspend them in 462 μL of KH2PO4 reaction buffer (see Note 23). 4. Add 7.5 μL of 40 mM resorcinol and 5 μL of 40 mM methyl viologen to the resuspended precipitates. 5. Incubate for 10 min at 30 °C (see Note 25). 6. Measure methyl viologen oxidation at 578 nm. 7. One unit of resorcinol reductase was determined as the amount of enzyme required to oxidate 1 μmol of methyl viologen per mL in 1 min (U mL-1/min-1).

3.1.21 Resorcinol Hydrolase [20]

1. Take 1.5 mL of sample and centrifuge at 10,000 rpm for 10 min. 2. Precipitate the proteins on supernatant with 80% (w/v) ammonium sulfate. 3. Centrifuge again and resuspend the precipitates, and resuspend them in 450 μL of Tris–HCl reaction buffer (see Note 23).

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4. Add 12.5 μL of 40 mM of resorcinol and 12.5 μL of 40 mM of K3[Fe (CN)6] to the resuspended precipitates. 5. Incubate for 10 min at 30 °C (see Note 25) under anaerobic conditions (see Note 26). 6. Measure K3[Fe (CN)6] oxidation at 578 nm. 7. One unit of resorcinol hydrolase was determined as the amount of enzyme required to oxidate 1 μmol of K3[Fe (CN)6] per mL in 1 min (U mL-1/min-1). 3.2

Metagenomics

3.2.1 Sample Preparation for DNA Extraction [21]

All steps require a sterile, nuclease-free condition to avoid DNA contamination and degradation. Wear gloves to avoid contaminating the nucleases, and keep them away from dangerous solvents. Samples from anaerobic biodigesters normally require a prewashing step where the cells are washed until waste and PCR/sequencing interfering materials have been removed. 1. Collect the equivalent of 1–1.5 mL from the biodigester in a microtube of 2 mL. 2. Centrifuge for 15 min at 10,000 rpm to precipitate cells; discard the supernatant by using a pipette or by inverting the tube (see Note 27). 3. Wash the pellet, filling the microtube with the equivalent of 1–1.5 mL of phosphate buffer at a pH of 8, and vortex to dissolve the pellet. 4. Repeat step 2 and 3 three times until the cells are clean and ready for DNA extraction. 5. Keep the pellet resuspended in 1.5 mL of phosphate buffer, pH 8.

3.2.2 Enzymatic Disruption of DNA Extraction [21]

DNA extraction is a protocol where cells are disrupted to obtain the DNA from cells for further techniques. Several case-dependent methods are applied. In this sense, enzymatic disruption has been proven to perform better for the samples from anaerobic biodigesters. 1. Centrifuge the pellets from the sample preparation step for 10 min at 10,000 rpm. Remove the supernatant. 2. Add 1.5 mL of extraction buffer to the same microtube with the pellet and 20 μL of snailase solution (20 mg/mL); solubilize cells by gently vortexing them until solubilized. 3. Incubate the mixture in a water bath at 37 °C for 1 h of enzymatic action. 4. After incubation, add 100 μL of 10% SDS to the microtube and gently vortex.

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5. Incubate the mixture in a water bath at 65 °C for 1 h and gently carry out end-over-end inversions every 15–20 min during incubation. 6. After incubation, sequentially add 20 μL of lysozyme (50 mg/ mL) and 10 μL of proteinase K (10 mg/mL) to the mixture. Carry out another incubation at 37 °C for 30 min of enzymatic reaction. 7. Centrifuge the incubated mixture for 10 min at 10,000 rpm, and transfer the supernatant into a new microtube. Wash the remaining pellet of the previous microtube with 1.5 mL of sterile deionized water, solubilize the cells, and centrifuge for 10 min at 10,000 rpm. Combine the supernatant with the first supernatant collected via pipette transfer. 8. Add an equal volume of chloroform/isoamyl alcohol (24:1) to the combined supernatants and gently vortex to separate the phases (see Note 28). Centrifuge for 5 min at 10,000 rpm to obtain better separation. 9. Transfer the upper phase (Aqueous phase) to a new, sterile microtube (see Note 29). Add 0.6 μL of volume (in the aqueous phase) to the upper phase collected in the new microtube that has the previously stored -20 °C isopropanol, and incubate the mixture for 30 min at 4 °C. 10. After incubation, centrifuge the mixture for 10 min at 13,000 rpm. 11. Discard the isopropanol from the precipitated DNA by pipetting it out, and add 700 μL of the previously stored -20 °C 70% ethanol for washing (see Note 30). Centrifuge for 5 min at 10,000 rpm. This washing step is repeated twice. 12. Open the tube and air-dry the DNA pellet at room temperature for 1 h or until fully dried. 13. Resuspend the dried pellet in 20–50 μL of ultrapure nucleasefree distilled water for library preparation. (See Note 31). 3.2.3 16S rRNA Amplicon [22]

PCR [22]

Based on the sequencing of 16S hypervariable regions, generally v3–v4, 16S metagenomics can be compared to identify and classify microorganisms from the bacteria and archaea domains. Thus, a PCR amplification of this region is required to obtain sequences for subsequent steps. 1. In a PCR 96-well plate, set up the PCR mixture in each well of the experiment, composed of 12.5 μL of 2X KAPA HiFi HotStart Ready Mix, 5 μL of Amplicon PCR Reverse Primer at 1 μM, Amplicon PCR Reverse Primer at 1 μM, and 2.5 μL of total genomic DNA at 5 μg/μL—for a final volume of 25 μL (see Note 32).

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2. Set the thermal cycler conditions with the following program: 95 °C for 3 min; 25 cycles of 95 °C for 30 s, 55 °C for 30 s, 72 °C for 30 s, and 72 °C for 5 min; and hold at 4 °C. After PCR, the oligonucleotides and oligonucleotide dimers are often left untouched, so a further PCR cleanup step is required. 2.1 After PCR, condensate might appear. Centrifuge the PCR 96-well plate for 1 min at a low rpm; 3000 rpm is generally sufficient at room temperature. 2.2 Vortex the AMPure XP beads for 30 s before its usage, ensuring that the beads are evenly dispersed. Add 20 μL of AMPure XP beads to each well of a 96-well plate, and gently pipette up and down for the entire mixture, then incubate at room temperature for 5 min (see Note 33). 2.3 Place tubes on a magnetic stand until the supernatant has cleared (2 min approximately). 2.4 Afterward, in the same magnetic stand, pipette to remove and discard supernatant. Wash the beads with recent freshly 80% ethanol by adding 200 μL to each experimental well of the 96-well plate. Incubate on the magnetic stand for 30 s, then carefully remove and discard the supernatant. Repeat this step twice. Finally, use a 10 μL pipette or a fine-tip pipette to move the excess ethanol to the used wells of the PCR 96-well plate. 2.5 With the PCR 96-well plate still on the magnetic stand, air-dry it for 10 min, then remove it from the magnetic stand (see Note 34). 2.6 Add 52.5 μL of 10 mM of Tris–HCl at pH 8.5 to the wells of the PCR 96-well plate, and mix it by pipetting up and down until the beads are fully resuspended. Incubate at room temperature for 2 min, and then place the tube on the magnetic stand until the supernatant has cleared— approximately 2 min. 2.7 Transfer 50 μL of the supernatant from the wells used on the PCR 96-well plate to another PCR 96-well plate. (See Note 35). 3.2.4 16S rRNA Library Preparation [22]

The library preparation indicates that indexing is necessary before sequencing DNA through next generation sequencing (NGS), to adhere sequences to the flow cell and identify each DNA sequence. 1. Collect only 5 μL of the previous PCR 96-well plate, and transfer to a well in a new PCR 96-well plate. 2. Using Index Primers 1 and 2, set up the PCR mixture into the well composed of 5 μL of Nextera XT Index Primer 1 (N7xx), 5 μL of Nextera XT Index Primer 2 (S5xx), 25 μL of 2× KAPA

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HiFi HotStart ReadyMix, and 10 μL of PCR-grade water—for a final volume of 50 μL. Pipette up and down to mix the components. Cover the plate with sealing tape, and centrifuge at 3000 rpm for 1 min at 20 °C to eliminate bubbles (see Note 32). 3. Set the thermal cycler conditions with the following program: 95 °C for 3 min; 8 cycles of 95 °C for 30 s, 55 °C for 30 s, 72 °C for 30 s, and 72 °C for 5 min; and hold at 4 °C. After PCR, oligonucleotides and oligonucleotide dimers are often left untouched, so a further PCR cleanup step is required. 1. After indexing the PCR plate, a condensation might appear, so centrifuge the PCR 96-well plate for 1 min at 1000 rpm at 20 ° C to collect condensation. 2. Vortex the AMPure XP beads for 30 s before use, ensuring that the beads are evenly dispersed. Add 56 μL of AMPure XP beads to each well of a PCR 96-well plate and gently pipette up and down to mix them, then incubate at room temperature for 5 min (see Note 33). 3. Place the plate on a magnetic stand until the supernatant has cleared (approximately 2 min). 4. Afterward, in the same magnetic stand, pipette the supernatant to remove and discard it. Then wash the beads with 80% ethanol by adding 200 μL to each well of the 96-well plate; incubate on the magnetic stand for 30 s; then carefully remove and discard the supernatant. Repeat this step twice. Lastly, use a 10 μL pipette or a fine-tip pipette to remove excess ethanol from the wells. 5. With the PCR 96-well plate still on the magnetic stand, air-dry the wells for 10 min, then remove them from the magnetic stand (see Note 34). 6. Add 27.5 μL of 10 mM of Tris–HCl at pH 8.5 to each well of the PCR 96-well plate, and mix by pipetting up and down until the beads are fully resuspended. Incubate at room temperature for 2 min, then place the PCR 96-well plate onto the magnetic stand until the supernatant has cleared (approximately 2 min). 7. Transfer 25 μL of the supernatant from the PCR 96-well plate to a new PCR 96-well plate. (See Note 35) 3.2.5 Quantification, Normalization, and Pooling [22]

The quantification, normalization, and pooling of libraries is necessary to ensure good case-dependent data reliability. For 16S metagenomics, more than 100,000 reads are necessary to adequately classify bacterial compositions. Illumina suggests using a fluorometric quantification method that uses dsDNA binding dyes. It is recommended to dilute

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libraries to 4 nM. The DNA concentration is calculated in nM on the basis of the size of DNA amplicons and by using the following formula: (concentration in ng/μL) (660 g/mol × average library size) × 106 = concentration in nM For example: 15 ng/μL (660 g/mol × 500) × 106 = 45 nM 1. Dilute the library using 10 mM of Tris–HCl at pH 8.5. Up to 96 libraries can be pooled for one MiSeq run. 2. Take 5 μL of a diluted library then mix libraries with unique indexes for pooling. A MiSeq run allows more than 20 million reads, for a maximum of 96 libraries. Sequencing Preparation

Clustering on the Illumina platform requires a DNA denaturing step for further sequencing and the addition of PhiX, a reliable library derived from PhiX genome that functions as a control for sequencing runs in Illumina.

DNA Denaturation [22]

1. In a nuclease-free sterile microtube of at least 1.7 mL of a mix of 5 μL of 4 nM of pooled library and 5 μL of 0.2 N of NaOH, gently vortex the mix and centrifuge it for 1 min at 3000 rpm at room temperature to remove bubbles and condensates. Incubate the mixture for 5 min at room temperature to let the DNA library to obtain ssDNA. 2. Add 990 μL of previously cooled HT1 solution to the tube containing the denatured DNA solution, for a final concentration of 20 pM of denatured library in 1 mM of NaOH. Keep the denatured DNA solution on ice and ignore the final dilution step (see Note 36). 800–1000 K/mm-2 raw cluster density is recommended by the Illumina platform, starting with concentrations of 4 pM and then adjusting accordingly. 3. Dilute the denatured library to 4 pM by pipetting 120 μL of 20 pM of denatured library and 480 μL of previously cooled HT1, inverting and centrifuging it for 1 min at 3000 rpm at room temperature, then store on ice until ready for use (see Note 37).

Denature and Dilution of PhiX Control [22]

The loading concentration of the library must be equal to that of the amplicon library and that of the one containing 5% of PhiX. 1. Prepare PhiX library at 4 nM by mixing 2 μL of 10 nM of PhiX library and 3 μL of 10 mM of Tris–HCl at pH 8.5. 2. In a nuclease-free sterile microtube of at least 1.7 mL of a mix of 5 μL of 4 nM of PhiX library and 5 μL of 0.2 N of NaOH,

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gently vortex the mix and centrifuge it for 1 min at 3000 rpm at room temperature to remove bubbles and condensates. Incubate the mixture for 5 min at room temperature to let the PhiX DNA library to obtain ssDNA. 3. Add 990 μL of previously cooled HT1 solution to the tube containing the PhiX denatured libraray (DNL) solution, for a final concentration of 20 pM of denatured library in 1 mM of NaOH. 4. Dilute the PhiX library to 4 pM by pipetting 120 μL of 20 pM of denatured library and 480 μL of previously cooled HT1, inverting and centrifuging for 1 min at 3000 rpm at room temperature, then store on ice until ready for use (see Note 36). Combine Amplicon Library and PhiX Control [22]

Running Sequencing on Illumina’s MiSeq Platform [22]

1. In a nuclease-free sterile microtube, mix 30 μL of the denatured and diluted PhiX and 570 μL of the denatured and diluted amplicon library, then keep it on ice in a heat block for an incubation period of 2 min at 96 °C. Invert twice to mix the microtube after the heat incubation and set it in an ice-water bath for 5 min. Running sequencing refers to the loading of prepared DNA samples on the sequencing platform. 1. Make a hole in the foil seal y using a sterile nuclease-free 1 mL tip. Charge the 600 μL of the combination of the amplicon and the PhiX control into the reservoir loading sample, then set up the sequencer control parameters directly in the software. Make sure not to touch the foil seal of the loading site. 2. Set up a run by using the MiSeq control software.

Bioinformatic 16S Amplicon Metagenomic Analysis Pipelines [22]

After sequencing, raw data in the FastQC format from the sequencers normally require a preprocessing quality analysis and trimming to ensure reliable results and no interferences, allowing only highquality reads for downstream analysis. Common tools for these tasks are FastQC, which assesses the quality of sequences on the basis of Phred scores, and Trimmomatic, which removes adapters and low-quality regions. Next, a quantification stage, where chimeras are removed and sequences are clustered to generate operational taxonomic units (OTUs) or amplicon sequence variants (ASVs), takes place, which can be further used during downstream analysis, where the Quantitative Insights into Microbial Ecology (QIIME) and Mothur pipelines are common applied. Alpha diversity and beta diversity can be performed with Shannon and NMDS, respectively, in R packages such as phyloseq. Taxonomy can be carried out throughout the mapping to the SILVA and RDP databases. Phylogenetic Investigation of communities by Reconstruction of Unobserved States (PICRUSt) is normally used to study the

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functionality of microbial communities on the basis of classification. MG-RAST is an easy-to-use web server that can conduct 16S rRNA analysis as previously mentioned without bioinformatic background researchers. 3.3 Gene Expression of mcrA

All the steps require sterile nuclease-free conditions to avoid RNA contamination and degradation. All the surfaces must be cleaned, and handling RNAse inhibitors requires wearing gloves. RNA is sensitive to environmental conditions, so work in a closed room under controlled conditions. During the process, use an ice cube to keep the reagents cold.

3.3.1 Sample Preparation for RNA Extraction [23, 24]

Samples of anaerobic digesters need to undergo a washing process before RNA extraction to remove interfering materials. 1. Collect 10–15 mL from the biodigester in a conical tube. 2. Centrifuge for 15 min at 500 rpm to precipitate the cells, and discard the supernatant by inverting the tube. 3. Wash the pellet by filling the tube with 10 mL of sterile double distilled water and cortex to dissolve the pellet. 4. Repeat steps 2 and 3 three times to clean the cells for RNA extraction. 5. Once the clean pellet is obtained, immediately resuspend it in 3–5 mL of lysis buffer in a 50 mL centrifuge tube. 6. Flash-freeze it in liquid nitrogen and store at -80 °C until ready for RNA extraction.

3.3.2 Physicochemical Cell Disruption for RNA Extraction [23, 24]

1. Quickly add 800 μL of acid saturated phenol and 800 μL of lysis buffer. 2. Vortex it until the pellet is suspended. 3. Place tubes in a heated shaker (preheated to 65 °C), and vortex at maximum intensity for 45 min at 65 °C. 4. Transfer lysate to a 2 mL microtube (sterile and RNAse-free). 5. Centrifuge the tube for 10 min at 10,000 rpm and 4 °C. 6. Transfer the top aqueous layer into another sterile and RNAsefree microtube containing 800 μL of chloroform. 7. Repeat steps 4, 5, and 6 one more time.

3.3.3 RNA Isolation [23, 24 ]

1. Add 75 μL of sodium acetate (3 M) and 1.8 mL of 100% ethanol. 2. Place tubes at -20 °C to precipitate RNA overnight. 3. Pellet RNA at 15,000 rpm for 30 min at 4 °C. 4. Cautiously decant supernatant, and add 1 mL of 70% ethanol to wash the pellet.

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5. Pellet RNA at 15,000 rpm for 30 min at 4 °C. 6. Remove ethanol by using a 1 mL micropipette. 7. Centrifuge again under the most recent conditions to collect any residue of ethanol and remove it with a 10 μL pipette. 8. Resuspend pellet in 50 μL of TE buffer, and mix, pipetting up and down. 9. Store RNA at -80 °C. 3.3.4 Gene Expression of mcrA with qRT-RNA

1. Conduct reverse transcription with iScript cDNA synthesis kit (Bio-Rad) in 20 μL of reaction volume containing 1 μg of total RNA incubated at 25 °C for 5 min.

Reverse Transcription [23, 24 ]

2. Transcript at 42 °C for 30 min, and terminate heating at 85 °C for 5 min. 3. The synthesized cDNA must be stored at -20 °C until ready for use.

Real-Time PCR [24]

1. In a PCR 96-well plate, set up the PCR mixture in each well of the experiment, composed of 10 μL of SYBR Green qPCR Master Mix, 0.4 μL of 10 μM of forward primer, 0.4 μL of 10 μM of reverse primer, 2 μL of cDNA, and 7.2 μL of RTPCR-quality H2O. 2. Set thermal cycler to be under the following conditions: 94 °C for 5 min; 40 cycles of 95 °C for 30 s, 57 °C for 45 s, 72 ° C for 30 s, and 72 °C for 5 min; and hold at 4 °C until ready for use.

Quantification and Interpretation [24]

1. Put out the tubes from real-time PCR instrument, and conduct a dissociation curve analysis with the saved copy of the setup file in Bio-Rad CFX Manager software. 2. Check to determine whether there is any bimodal dissociation curve or abnormal amplification plot. 3. Export the results to Excel software, and calculate the gene expression by using the 2-△Ct method.

3.4

Proteomics

3.4.1 Sample Extraction and Quantification [27]

1. Take 40 mL of the sample. 2. Centrifuge the sample at 10,000 rpm for 10 min to separate it via extracellular fraction (see Note 38). 3. Take the supernatant for the protein analysis. 4. Quantify protein concentration with Bradford assay (see Note 39).

3.4.2

SDS-Page [27]

1. For the resolving gel (10%, 1 mm), mix 2 mL of 30% (w/v) acrylamide–bis acrylamide, 1.5 mL of 1.5 M Tris–HCl at pH 8.8, 2.4 mL of H2O distillate, 60 μL of 10% (w/v) SDS, 30 μL of 10% (w/v) APS, and 3 μL of TEMED (see Note 40).

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2. Put the mixture between the spacer plate and the short plate, and fill in the space with water or isopropanol (see Note 41). 3. Wait until gel forms, then throw out the water. 4. For the stacking gel (4%, 1 mm), mix 0.396 mL of 30% (w/v) acrylamide–bis acrylamide, 0.756 mL of 0.5 M Tris–HCl at pH 6.8, 1.8 mL of distilled H2O, 30 μL of 10% (w/v) SDS, 15 μL of 10% (w/v) APS, and 3 μL of TEMED (see Note 42). 5. Pour the stacking gel mix over the resolving gel, and carefully place the gel comb. 6. Wait until gel forms, then take off the gel comb. 7. Fill the electrophoresis cell with running buffer 1× (see Note 43). 8. Prepare samples by maintaining the following the ratio of 1:1 sample: lysis buffer. 9. Load the molecular weight standard and the samples onto the SDS-PAGE gel. 10. Connect the electrophoresis cell, and run at 120 V for 120 min. 11. After the electrophoresis is complete, turn off the power supply and remove the gel. 3.4.3

1D-Page [28]

1. To prepare the sample, pipet 200 μL of each sample and 200 μL of rehydration buffer as a line along the edge of the channel in the rehydration/equilibration tray (see Note 44). The sample is included in the hydration solution, and the sample is passively taken up into the IPG strip during rehydration (passive hydration). Take care not to introduce any bubbles. 2. Gently place the IPG strips, gel-side down, onto the sample. The “+” and the pH range marked on the IPG strips should be legible. Overlay each of the IPG strips with 200 μL of mineral oil to prevent evaporation during the rehydration process. 3. Cover the rehydration tray with the plastic lid, and leave the tray sitting on a level bench overnight (16 h) to rehydrate the IPG strips and load the protein sample. 4. For all IPG strip lengths, use the default temperature of 20 °C, with a maximum current of 50 uA/IPG for 12 h. 5. When the electrophoresis run has been completed, remove the IPG strips from the focusing tray; transfer them gel-side up into a new, clean, dry disposable rehydration tray; and let the mineral oil drain from the IPG strips for ~5 s before transferring them. 6. After the first dimension, take the strips and take off the excess mineral oil. Take care not to introduce any bubbles, which may

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interfere with the even distribution of the sample in the IPG strips. 7. If the equilibration step is not to be directly proceeded to, cover the tray containing the IPG strips, wrap it in plastic wrap, and place it in freezer storage at -70 °C. 3.4.4

2D-Page [28]

1. Add 4 mL of equilibration buffer I to each IPG strip (see Note 45). 2. Place the tray on an orbital shaker for 10 min (see Note 46). 3. After the incubation period, remove the buffer, carefully decanting the liquid from the tray. 4. Add 4 mL of equilibration buffer II to each IPG strip. Return the tray to the orbital shaker for 10 min (see Note 46). 5. During the incubation, melt the overlay agarose. 6. Discard the equilibration buffer II by decanting at the end of the incubation, as described in step 4 above. 7. Take off the strip, and place it on the top of the SDS-PAGE (7.5–15%). 8. Add molten agarose solution to cover the strip and ensure contact with the gel, taking care not to introduce any bubbles. 9. Fill the electrophoresis cell with running buffer. 10. Connect the electrophoresis cell, and run for 30 min at 50 V. 11. After the first 30 min, run at 120 V for 90 min. 12. After the electrophoresis is complete, turn off the power supply and remove the gel. 13. Run the electrophoresis under similar conditions for SDSPAGE.

3.4.5

Gel Staining [28]

1. Wash the gel three times with distilled water for 5 min each, changing the water each time. 2. Add 50 mL of Coomassie stain solution, and agitate for 1 h at room temperature (see Note 47). 3. After that 1 h, remove the staining solution and add 50 mL of destaining solution until the protein bands are visible. 4. The intensity of each band and its molecular weight can be determined by using Gel Analyzer software (http://www. gelanalyzer.com).

3.4.6

LC-MS/MS [29, 30]

1. Excise the SDS-PAGE gel protein bands and/or the 2D-PAGE spots of interest, washing the gel with NH4HCO3 and acetonitrile. 2. Swell the gel in digestion buffer for 45 min at 4 °C.

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3. Aspirate the supernatant, and replace it with 5–10 μL of digestion buffer without trypsin. 4. Incubate overnight at 37 °C. 5. Extract the peptides through three changes of 5% (w/v) formic acid and acetonitrile (20 min for each change) at room temperature. 6. Dry down the samples, and resuspend them in 15 μL of 5% acetonitrile with 0.1% formic acid. 7. Inject 5 μL of them sample into a C18 HPLC column for preconcentration and washing. 8. Elute peptides with a 120 min gradient of 5% acetonitrile with 0.1% formic acid to 40% acetonitrile with 0.1% formic acid at a nominal post-split flow rate of 250 μL per min-1. 9. Program a spray voltage of 2 kV, and set the capillary temperature at 170 °C. 10. Samples can be scanned when within the range of 400–1500 m/z. 3.4.7 Bioinformatic Analysis of LC-MS/MS

4

For peptide identification, convert the obtained data to SEQUEST format by using the most recent version of the Mascot software (www.matrixscience.com) and compare them against protein databases such as UnitprotKB, NCBI, and Swiss-Prot.

Notes 1. When assaying the total solids, the dish material should be either porcelain, platinum, or high-silica glass and the volume should be 100 mL, because the temperature will be in a range of 103–105 °C, and those materials can resist heat at these temperatures. 2. The materials and apparatuses for the total suspended solid (TSS) assay at 103–105 °C will be the same as those for the total solid (TS) assay, except for the evaporating dishes, the steam bath, and the drying oven. For this procedure, aluminum weighing dishes will be required. However, the materials and apparatuses of the fixed solids and volatile solids (FSs and VSs, respectively), ignited at 550 °C, are the same as those for the TS assay. 3. Acid ethylene-glycol reagent preparation requires taking 30 mL of ethylene-glycol and mixing it with 4 mL of diluted sulfuric acid. This reagent must be fresh, so it must be prepared each day. If the blank has a superior value of 200 ppm (specifically acetic acid), distill it to purify the ethylene-glycol from sodium hydroxide.

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4. For the preparation of 0.1 N of standard sulfuric acid or the same of hydrochloric acid, follow the instructions below. Prepare the acid solution at the indicated normality (0.1 N). Standardize it through titration, using 40 mL of 0.05 N of sodium carbonate solution with 60 mL of distilled water and a beaker as a container. The potentiometric titration must reach a pH of 5. Take out the electrodes, and start boiling the beaker for 3 to 5 min, covering the top of the beaker with a glass cover. Cool the beaker at room temperature, and finish the titration until the pH inflection point has been reached. To calculate normality, use the following equation: N=

A B 53:00 C

where N = normality; A = g of Na2CO3 weighted into a flask; B = mL of Na2CO3 solution used for titration; and C = mL of acid needed. 5. To prepare approximately 0.25 M of FAS solution, adhere to the following steps: add 98 g of Fe (NH4)2(SO4)2·6H2O to distilled water, add 20 mL of H2SO4 concentrate, cool, and dilute to 1 L. FAS solution needs to be standardized every day against K2Cr2O7 solution: Dilute 25 mL of K2Cr2O7 standard to 100 mL, add 30 mL of H2SO4 concentrate, and cool. Finally, titrate with FAS solution by using 0.10–0.15 mL (2 or 3 drops) of ferroin indicator. Molarity should be approximately 0.25 mol/L, but for accuracy, calculate with it by using the following equation: FAS ðstandard ferrous ammonium sulfateÞ molarity volume 0:041 K2Cr2O7 solution =

titrated, mL Volume of FAS used



0:25

in titration, mL 6. Slowly add chitin powder to HCl and keep at 30 °C for 60 min, where the colloidal chitin should precipitate thanks to the slow addition of 2 L of water at 4 °C. 7. SDS is not easy to dissolve and generate foam from, so mix it gently until fully dissolved and wait until little or no foam is formed before storing it. Storing it in fridge results in a solid solution. If it is not frequently used, it can be stored frozen and be thawed 1 min before an experiment.

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8. Use a lab fume hood when opening chloroform, to prevent any problems or symptoms of exposure, such as dizziness and vomiting. 9. This entire process must proceed under a vape hood to avoid accidents. 10. The pH of the solution will not change in the first wash with citrate buffer, so wash it multiple times, and frequently measure the pH until a pH of ~4.3 has been obtained. 11. Use concentrated HCl to obtain the required pH. 12. APS must be prepared at the time of use and never stored. 13. Before use, the dish must be weighed. The temperature to which it will be exposed varies depending on what is to be measured. If volatile solids are the target, the dish must be exposed in a muffle furnace at 550 °C for 1 h. On the other hand, if the target is total solids, the temperature must be set at 103–105 °C for 1 h, after which the dish must be cooled and stored in a desiccator until it is needed again. Before using any dish, measure its weight. 14. Constant weight is when weight is kept the same or is no longer than 4% of the previous weight. When weighing the sample, changes could be attributed to air exposure or sample degradation, so to obtain better accuracy for all the samples, at least 10% must be analyzed in duplicate. The duplicate must agree within 5% of the average weight. 15. TSS is an assay used to measure the solids that are captured in the filter, compared against the dissolve solids. One manner to calculate this is to determine the difference between the total solids and the dissolved solids; another is to determine the difference by using the filter weight. 16. An assay has been conducted on the data from three laboratories on four samples, performing 10 replicates. The standard deviation was 11 mg/L at 170 mg/L of volatile total solids [14]. 17. For the curve graph of acetic acid, use the following concentrations; 500, 1000, 1500, 2000, and 2500 ppm of acetic acid. Also, this assay is carried out to detect organic acids and their salts in samples from a biodigester (sewage sludge liquor). The results are used to produce a graph of volatile acid concentrations. In the distillation method for determining organic acids, using acetic acid offers more precise results than using phosphoric acid. 18. When alkalinity is due to the presence of carbonates or bicarbonates, pH has an equivalence point in the titration, where this point is due to the concentration of CO2 at this stage. Nevertheless, this can change for certain reasons, such as some

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of the original total carbonates were lost during titration. Thus, use the endpoint as a reference. Depending on the indicator, the color will change, but the value will remain the same, at pH 8.3. To determine the alkalinity of the titration for this assay, phenolphthalein or metacresol purple may be used (because the alkalinity of the titration is at pH 8.3), but other indicators can be also used, such as bromocresol green or mixed bromocresol green-methyl red indicator, for a pH of 4.5. 19. Low alkalinity is defined less than 20 mg of CaCO3/L or other alkalinities, such as bicarbonates and hydroxides, so the procedure will vary depending on the concentration of alkalinity. This titration will be working with a different range of pHs, from 4.3 to 4.7, and the unit will be reduced to a 0.30 pH unit. 20. The method of determining COD will vary depending on the concentration of O2. Samples with low COD values (samples below 50 mg O2/L) require some variations to the assay process that are in part B of the method; on the other hand, for samples with COD values higher than 50 mg O2/L, the instructions for the procedure are in part A of the method. 21. In a COD assay, it is important to know the concentration from the samples, because if the samples are overconcentrated with organic matter and if that COD is more than 900 mg O2/L, the sample must be diluted. On the other hand, the color will change, and measuring it will not be possible. In this case, the exact amount of COD cannot be measured, but it can interpreted from the samples and the method, depending on the sample. 22. For an alternative procedure for low-COD samples, take care to prevent contamination, and a larger volume of the sample will be need to be taken just before this one has been digested by reflux. As was mentioned, this is a better strategy to increase the accuracy and precision of the assay. All the reagents must be added to a sample with a volume larger than 50 mL but with a total volume of 150 mL. This volume is obtained from the boiling in the refluxing flask, with an open atmosphere, because the condenser will not be joined. Calculate an amount of HgSO4 that must be added, on the basis of a weight ratio of 10:1 for HgSO4:Cl-, using the present amount of Cl- in the original sample (volume), and with a blank reagent, following the same procedure from before. One of the advantages of this technique is obtaining the concentration of the sample without a significant loss of volatile materials. On the other hand, there are some volatile materials that are hard to digest (such as volatile acids) that are lost in the process; nevertheless, this loss also leads to an improvement in the ordinary evaporative concentration. If the duplicate experiments are not expected to

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produce precise results, use samples with COD values higher than 50 mg O2/L. 23. Resuspended precipitates can be stored at -20 °C until ready for use, preferably for no longer than 1 week. 24. The volume of the substrate and resuspended precipitates can be modified as needed. 25. For all the enzymatic assays, optimize the pH, temperature, and time for an optimal activity. 26. Anaerobic conditions can be perforated using Hungate tubes or cuvettes flushed with N2 and closed using butyl septa. 27. Pellet normally remains stuck in the bottom of the tube, but the tube can be freely inverted so that the supernatant can be discarded; otherwise, use a pipette for the removal. 28. Chloroform/isoamyl alcohol (24:1) is a highly dense reagent that does not stay in the tip that well. Slowly pipette it to obtain the appropriate volume, and quickly serve it in the tube. 29. The organic and aqueous phases must be fully separated. When pipetting the aqueous phase, take care to not touch the organic phase and the middle foaming phase, to avoid DNA contamination and lowing the purity. It is recommended to pipette in the middle of the microtube. 30. The DNA pellet is tough to see, depending on the concentration, but it normally remains attached to the bottom part of the microtube. 31. For better resuspension, incubate the resuspended pellet at 65 °C for 30 min. If it is not meaning to be used in DNA extraction, then to prevent DNA degradation, store the tubes at -20 °C for up to 1 week. 32. Make sure that pipettes are well calibrated to avoid high differences in amplicon sizes. When working on many samples, prepare the reagents for multiple samples; include PCR master mix in all the samples to avoid changes in volumes from small errors caused by pipetting. 33. Ensure that the beads are at room temperature. Please do not reuse beads, to avoid contamination. Before usage, vortex them until they are well dispersed and until the liquid’s color becomes homogenous. Beads are viscous, so gently and slowly pipette and dispense them to keep the required quantity. When removing the supernatant, take care not to disturb the beads. Once the solution has cleared, keep the plate on the magnetic stand and aspirate it slowly to prevent the beads from sliding down the tips and sides of the wells.

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34. A longer drying time might be needed for full evaporation. Let it dry out for longer than 10 min, as required. Ethanol can be removed by using a 10 μL pipette. 35. If the tubes are to be used later, store them at -20 °C for up to 1 week to prevent DNA degradation. 36. Place the denatured DNA on ice after thawing it at room temperature, but do not expose it to temperatures above 30 ° C. 37. Make sure that the tubes are fully sealed and that the liquid is down after mixing; otherwise, the concentration and the volume will widely vary. 38. To obtain the intercellular fraction, resuspend the pellet in 0.1 M of phosphate buffer at a pH of 7.5 and sonicate it at 100% amp for 5 min and for 30 pulses. Next, centrifuge the suspension under the same conditions to obtain the extracellular fraction. 39. If the sample has less than 3 g/L of protein, concentrate the samples with membranes of 10 kDa. 40. Because APS and TEMED are catalyzers, they are the last reagents to be added. 41. Water and isopropanol alcohol allow gel to form, erecting a barrier for the oxygen that inhibits polymerization. 42. The respective amounts of the reagents are for one gel. For two or more gels, the amounts must be increased. 43. Running buffer 1× can be achieved through dilution with running buffer 5×. The latter can be reused several times and until gel quality is preserved. 44. Using rehydration and an equilibration tray is a convenient method to prevent cross contamination among protein samples. Take care not to introduce any bubbles during the rehydration, which might interfere with the even distribution of samples in the IPG strips. 45. Two-step equilibration also ensures that cysteines are reduced and alkylated, which minimizes or eliminates the vertical streaking that may be visible after staining the second dimensions of the gels. 46. The IPG strips require 10–15 min to thaw. It is best not to leave the thawed IPG strips for longer than 15–20 min because the diffusion of proteins can reduce the sharpness of the protein spots. 47. The staining time could depend on the quality of the Coomassie stain solution. If necessary, another kind of stain could be used, depending on the sensitivity and the properties required. Stained gels can be stored in distilled water.

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INDEX B Bioactive compounds alkaloids .................................................................8, 20 carotenoids β-carotene .......................................................... 162 lycopene ............................................................. 162 essential oils ................................................24, 28, 116 glucosinolates ............................................................ 19 nutraceuticals................................................ 22, 28, 32 pectin .............................................................. 114, 116 phenolic compounds hesperidin ...............................................................7 saponins ................................................. 19, 20, 25, 28 terpenoids ..............................................................7, 28 xanthin alcaloids .......................................................... 8 Bioactivity anti-aging activity .................................. 11, 12, 14, 21 anti-inflammatory activity.................................... 7–12, 14–16, 18–21 antimicrobial activity ..............................17, 21–22, 86 antioxidant activity ..................................8–14, 17, 18, 20, 23, 25–28, 55, 56, 70, 71, 86, 87, 93, 148 antiproliferative activity..................... 9–11, 16, 19, 21 microbiota modulator............................................. 115 Biological control Trichoderma spore production antibiotic production ........................................ 178 mycoparasitism .................................................. 170 systemic immune response ............................... 170 Biomass ........................................ 3–5, 30, 67–76, 80, 81, 118, 120, 129–131, 134, 139, 165–167 Bioprocess biotechnological process......................................... 170 microbial biomass production anaerobic digestion ...................................187–225 solid-state fermentation ...........................147–158, 161–167 submerged fermentation ..........................177–184 Bioreactor Erlenmeyer flask ................................... 149, 152, 153, 157, 171, 173, 189, 201 petri dishes.....................................163, 165, 171, 173

plastic bags........................................................ 81, 179 Brewer’s spent grain (BSG) ............................... 29, 79–82

C Cancer.....................................................8, 16, 17, 20, 21, 24, 47, 86, 115, 162 Cardiovascular diseases (CVDs) ............................. 15, 18, 20, 23, 162 Chemical composition .................................................. 127 Chemical oxygen demand (COD) ..................... 188, 189, 200, 223, 224 Circular economy (CE) ............................. 33, 44–46, 63, 68, 105, 116, 117, 162 Coffea arabica .................................................... 13, 56, 57 Cosmetic ingredients ..................................................7, 25 Cosmetic products ....................................................24, 29 Crop protection ............................................................ 170

D Deep eutectic solvents (DESs) ............................... 28, 30, 67–76, 79–82 Diabetes ...............................................13, 20, 22, 47, 115 Drying process ................................................59, 62, 117, 150, 156, 172, 180

E Energy production ........................................................ 127 Environmental pollution prevention greenhouse gas (GHG) emissions prevention....... 127 Enzymes amylase................................................... 191, 204, 205 benzoyl-CoA reductase.................................. 192, 208 cellulase ...................................................................... 31 chitinase .......................................................... 192, 206 hemicellulose ...................................79, 126, 131, 137 lipase .............................................................. 179, 181, 182, 192, 206 phloroglucinol reductase ............................... 192, 209 protease.....................................................86, 192, 208 resorcinol hydrolase ....................................... 193, 210 resorcinol reductase ....................................... 193, 209 xylanase ........................................................... 192, 207

Cristobal N. Aguilar Gonzalez et al. (eds.), Food Waste Conversion, Methods and Protocols in Food Science, https://doi.org/10.1007/978-1-0716-3303-8, © The Editor(s) (if applicable) and The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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230 Index F

Food by-products........................... 4–22, 28, 30, 33, 122 Food packaging .................................................... 4, 25–28 Food products .............................. 4, 25–28, 32, 101, 105 Food waste (FW) coffee pulp ................................ 5, 56, 57, 60, 61, 102 fish by-products.............................................. 177–184 litchi by-products ................................ 56, 57, 61, 102 orange by-products ........................................ 114–118 tomato waste .................................................. 161–167 Food waste conversion ........................... vii, 3–33, 43–50, 55–63, 67–76, 79–82, 85–94, 97–109, 113–122, 125–139, 147–158, 161–167, 169–175, 177–184, 187–225

G Gene expression ......................... 188, 195–196, 216–217 Gravity technology............................................... 119, 120 Green chemistry ........................................................68, 71

H High-performance liquid chromatography (HPLC)...................72, 91–93, 165, 166, 220 High protein meals ....................................................... 179 Hydrothermal pretreatment autohydrolysis.......................................................... 130 hydrothermolysis..................................................... 130 liquid hot water ....................................................... 130 subcritical water treatment ..................................... 130

I Integrated valorisation .................................................. 116

L Lignin ............................23, 79–82, 86, 87, 91, 127, 137 Liquid Chromatography-Mass Spectrometry (LC-MS) .......................................72, 219–220

M Metabolic syndrome (MS)................................13, 14, 22, 23, 72, 76, 197 Metagenomics ............................ 188, 193–195, 210–216 Methane measurement ............................... 188, 191, 204 Microencapsulation ...................................................55–63 Microencapsulation ...................................................55–63 Microorganism bacteria strains Pediococcus acidilactici ...................................... 179 Staphylococcus warneri....................................... 179 fungal strains

Aspergillus niger GH1.............................. 148, 163 Rhizopus oligosporus........................................... 148 Rhizopus oryzae ................................................. 148 Trichoderma asperellum .................................... 171 Microwave hydrodiffusion................................... 118, 119 Molecular analysis ................................................ 187, 188

N Natural deep eutectic solvents (NADES) ................30, 68 Natural food additive ...................................................... 25 Neurological diseases ......................................... 20, 23, 24

P Physicochemical analysis ...................................... 189–191 Phytochemical ....................................................... 4, 8, 22, 68–70, 72, 74–75 Phytopathogens........................................... 169, 170, 174 Proteomics.................................. 188, 196–197, 217–220

R Real-time Polymerase Chain Reaction (RT-PCR) complementary deoxyribonucleic zcid (cDNA) ........................................................ 217 16S rRNA gene ......................................193, 211–212

S Secondary metabolites .......................................... 4, 7, 19, 47, 69, 99, 148, 151 Sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE).............................. 197, 218, 219 Solid-liquid extraction (SLE) ......................................... 29 Sustainable biorefinery production value added product biobased materials ............................................. 129 biofertilizers....................................................... 127 biofuels .............................................................. 130 chemicals............................................................ 129

U Ultrasonication................................................................ 74

V Valorization .................................................. 4, 55, 68, 80, 113–122, 126, 130, 177–184 Volatile fatty acids profile (VFA) ................ 188, 191, 204

Z Zero-waste approach....................................................... 45