Mycoremediation Protocols (Springer Protocols Handbooks) 1071620053, 9781071620052
This volume provides a wide range of aspects related to mycoremediation, which can be applied for both basic and advance
121 17
English Pages 264 [251] Year 2022
Preface
Contents
Contributors
Chapter 1: Isolation, Enrichment, and Characterization of Fungi for the Degradation of Organic Contaminants
1 Introduction
2 Materials
3 Screening of Pesticide-Degrading Fungal Strains
3.1 Isolation of the Fungal Strains
3.2 Purification of Fungal Isolates
3.3 Morphological Characterization of Fungi
3.4 Microscopic Observation of Fungi by Temporary Wet Mount Technique
3.5 Mounting of Fungal Cells by Lactophenol Cotton Blue
3.6 Molecular Characterization of Fungi
3.6.1 Genomic DNA Extraction
3.6.2 Polymerase Chain Reaction (PCR)
4 Notes
References
Chapter 2: Protocol for the Assessment of Mycoremediation of Polycyclic Aromatic Hydrocarbons
1 Introduction
2 Materials
2.1 PAHs Extraction and Quantification by HPLC-PDA
2.2 Soil Enzymatic Activities
2.2.1 Dehydrogenase Activity (DHA)
2.2.2 Total Microbial Activity (TMA)
2.2.3 Urease Activity (UA)
2.3 Germinability Test
2.4 Test of Survival and Reproduction of Collembolan (Folsomia candida)
3 Methods
3.1 Assessment of the PAH Mycodegradation: PAH Extraction and Quantification
3.2 Recovery of Soil Microbial Functions
3.2.1 Dehydrogenase Activity
3.2.2 Total Microbial Activity
3.2.3 Urease Activity
3.3 Germination Test
3.4 Test of Survival and Reproduction of a Type of Springtails/Collembolan (Folsomia candida)
4 Notes
References
Chapter 3: Characterization and Screening of Pesticide-Degrading Indigenous Fungi from Soil and Water
1 Introduction
2 Materials
2.1 Pesticides, Reagents, and Solvents
2.2 Media
2.3 Soil Sampling/Water Sampling
3 Methods
3.1 Isolation of Pesticide-Degrading Fungi from Contaminated Soil/Water
3.2 Identification and Characterization of Fungal Isolates
3.3 Growth Inhibition Studies
3.4 Pesticide Biodegradation Assay for Screening of Potential Fungal Isolate
4 Notes
References
Chapter 4: Mycoremediation of Synthetic Textile Dyes by Fungi Isolated from Textile Wastewater Effluent and Soil
1 Introduction
2 Materials
2.1 Modified Kirk´s Medium
2.2 Potato Dextrose Broth (PDB)
3 Methods
3.1 Preparation of Textile Dyes and Quantification
3.2 Sample Collection
3.3 Analysis of Physicochemical Parameters of the Collected Water Samples
3.4 Enrichment of Textile Wastewater and Soil Samples
3.5 Isolation of Textile Dye-Decolorizing Fungi
3.6 Screening of Textile Dye-Decolorizing Fungi
3.6.1 Solid Medium Screening
3.6.2 Liquid Medium Screening
3.7 Optimization of the Textile Dye Decolorization Processes by Selected Fungi
3.8 Assessment of the Toxicity of the Decolorized Dye Solutions
4 Notes
References
Chapter 5: Protocol for Screening Endophytic Fungi Against Heavy Metals
1 Introduction
2 Materials
2.1 Endophytic Fungus Isolation and Purification
2.2 Heavy Metals Screening of the Fungal Isolates
3 Methods
3.1 Isolation of Endophytic Fungi
3.2 Preparation of PDA Supplemented with the Heavy Metal Solution (Prior to Screening)
3.3 Screening of Fungal Isolates against Heavy Metals
4 Notes
References
Chapter 6: Screening of Phyllosphere Fungi Inhabiting in Urbanized Areas for Phylloremediation Capabilities of Polyaromatic Hy...
1 Introduction
2 Materials
2.1 Isolation of Phyllosphere Fungi
2.1.1 Chemical Composition of Bacto-Bushnell Hass (BBH) Broth Medium
2.2 Screening and Confirmation of PAH-Degrading Potential of Phyllosphere Fungi
2.2.1 Screening of PAH Degradation Potential Using Plate Assay
2.2.2 Determination of PAH Degradation Potential Using UV-Visible Spectroscopy
2.2.3 HPLC Analysis of PAH Degradation
2.3 Identification of PAH-Degrading Fungi
2.3.1 Molecular Analysis to Identify Them up to Species Level
Extraction of Fungal Genomic DNA
2.4 Enzyme Assay
3 Methods
3.1 Leaf Sample Collection
3.2 Isolation of Phyllosphere Fungi
3.2.1 Isolation of Phylloplane-Inhabiting Fungi
3.2.2 Isolation of Endophytic Phyllosphere Fungi
3.2.3 Microscopic Observations of Leaf Endophytes
3.2.4 Endophytic Fungal Observations Through SEM
3.3 Screening and Confirmation of PAH-Degrading Potential Phyllosphere Fungi
3.3.1 Plate Assay (Primary Screening)
3.3.2 Spectrophotometric Analysis (Secondary Screening)
3.3.3 HPLC Quantification of Polyaromatic Hydrocarbon (Confirmation Test)
3.4 Identification of Phyllosphere Fungi
3.4.1 Identification of Phyllosphere Fungi up to Genus Level
3.4.2 Molecular Identification of PAH-Degrading Phyllosphere Fungi
Extraction of Fungal Genomic DNA
Amplification of Fungal 18S rRNA Region of Genomic DNA by PCR Method
Agarose Gel Electrophoresis
DNA Sequencing
GenBank Search
3.5 Enzymatic Degradation of PAH by Extracellular Enzymes
3.5.1 Manganese-Dependent Peroxidases (MnP) Activity
3.5.2 Lignin Peroxidases (LiPs) Activity
3.5.3 Laccases Activity
3.5.4 Calculation of Enzyme (MnP, LiP, and Laccases) Activities
4 Notes
References
Chapter 7: Methods for Design and Bioremediation Applications of Reactors Based on Immobilized Fungi
1 Introduction
2 Materials
2.1 Trickle-Bed Bioreactors
2.1.1 WRF-Based Bioreactor for Degradation of Textile Dyes
Scaling-Up of Trickle-Bed Reactors
2.1.2 Spent Mushroom Substrate-Based Bioreactor for Degradation of Endocrine-Disrupting Compounds (EDCs)
2.1.3 SMS-Based Bioreactor for Degradation of Polychlorinated Biphenyls (PCBs)
2.2 Rotating Drum Biological Contactor (RDBC) for Biodegradation of Textile Dyes
2.3 Rotating Disk Reactor (RDR) for Biodegradation of Textile Dyes
2.3.1 Scaling-Up of Rotating Biological Contactors
3 Methods
3.1 White-Rot Fungus-Based Bioreactor for Degradation of Textile Dyes
3.1.1 Examples of Application
3.2 SMS-Based Bioreactor for Degradation of EDCs
3.2.1 Examples of Application
3.3 SMS-Based Bioreactor for Degradation of PCBs
3.3.1 Examples of Application
3.4 Rotating Drum Biological Contactor (RDBC) for Biodegradation of Textile Dyes
3.4.1 Examples of Application
3.5 Rotating Disk Reactor (RDR) for Biodegradation of Textile Dyes
3.5.1 Examples of Application
4 Notes
References
Chapter 8: Denaturing Gradient Gel Electrophoresis (DGGE) Analysis of the Fungi Involved in Biodegradation
1 Introduction
2 Materials
2.1 Fungal DNA Extraction
2.2 For DGGE PCR
2.3 Gel Casting/Loading/Running
2.4 Staining and Visualization
3 Methods
3.1 DNA Extraction by the CTAB Method
3.2 DGGE PCR
3.3 Gel Casting/Loading/Running
3.4 Staining and Visualization
4 Notes
References
Chapter 9: Optimization of Mycoremediation Process for the Isolated Fungi
1 Introduction
2 Materials
3 Methods
3.1 Preparation Media and Stock Solutions for the Mycoremediation
3.2 Mycoremediation of the Xenobiotics
3.3 High-Performance Liquid Chromatography of the Extracted Sample
3.4 Calculation of the Mycoremediation Kinetics
4 Notes
References
Chapter 10: Protocol for Assessing Mycoremediation of Acidic Radioactive Wastes
1 Introduction
2 Materials
2.1 Media and Solutions
2.1.1 Sodium Dodecyl Sulfate-Polyacrylamide Gel Electrophoresis (SDS-PAGE) Solutions
2.1.2 Solution for DNA Detection Using Agarose Gel Electrophoresis
3 Methods
3.1 Elemental Analysis of Solid and Solution
3.2 Samples Collection and Preparation
3.3 Isolation of Active Fungi in Acidic Radioactive Wastes Remediation
3.4 Screening of Fungal Isolates for Radioactive Wastes Remediation
3.5 Efficiency of Filamentous Aquatic Fungi to Uptake Radioactive Cs-137 and Co-60 from Exposing Wastes
3.6 Efficiency of Fungi to Uptake Radioactive Cs-137 and CO-60 from Exposing Wastes
3.7 Factors Affecting Radionuclide Uptake
3.8 Effect of Pigments in Radionuclide Uptake
3.9 Identification of the Most Active Fungal Isolate
3.10 Transmission Electron Microscope for Characterization
3.11 Phenotype Characterization
3.12 Protein Analysis of Fungal Strains and/or Supernatants Free Cells Using SDS-PAGE
4 Notes
References
Chapter 11: Protocol for Screening Low-Density Polyethylene (LDPE)-Degrading Soil Fungi Isolated from Urban Waste Dumping Sites
1 Introduction
2 Materials
2.1 Culture Media
2.2 Test Material
2.3 Other Chemicals
3 Methods
3.1 Pretreatment, Sterilization, and Disinfection of LDPE
3.1.1 UV Pretreatment
3.1.2 Thermal Pretreatment
3.1.3 Presterilization of LDPE Films and Pellets
3.1.4 Disinfection of Fungal-Treated LDPE Films and Pellets
3.2 Sampling of Fungi
3.3 Isolation of Fungi
3.3.1 Spread Plate Method
3.3.2 Single Spore Isolation
3.3.3 Hyphal Tip Isolation
3.4 Screening of LDPE-Degrading Fungi
3.4.1 Screening of Fungi Using mPDB Medium
3.4.2 Screening of Fungi Using BHB Medium
3.4.3 Recovery of Pellets and Determinations of the Average Percentage of Weight Loss (WLP)
3.5 Assessment of Biodegradation Potential of LDPE Films by Selected Fungi
3.5.1 Incubation of LDPE Films on Solid Medium
3.5.2 Incubation of LDPE Films in the Liquid Medium
3.6 Quantitative Assessment of LDPE Biodegradation
3.6.1 Calculation of Percentage of Weight Loss (WLP)
3.6.2 Calculation of Carbonyl Index (CI)
3.6.3 Calculation of Percentage of Crystallinity
3.6.4 Calculation of Relative Elongation and Relative Tensile Strength
3.7 Qualitative Analysis of LDPE Biodegradation: Scanning Electron Microscopy (SEM)
4 Notes
References
Chapter 12: Production of Laccases from Agricultural Wastes: Strain Isolation and Selection, Enzymatic Profiling, and Lab-Scal...
1 Introduction
1.1 Occurrence of Laccase in Fungal Systems
2 Materials
2.1 Culture Media
3 Methods
3.1 Isolation of Promising Fungal Strains
3.2 Qualitative Assays for Laccase Production
3.3 Laccase Production in Solid-State Fermentation (SSF)
3.4 Quantitative Laccase Activity and Protein Content Determination
3.5 Conditioning of Enzyme Extracts: Partial Laccase Purification
3.5.1 Aqueous Two-Phase Extraction (ATPE)
3.5.2 Foam Fractionation (FF)
3.5.3 Cross-Linking of Enzyme Aggregates (CLEAs)
3.6 Concentration
3.7 Affinity Chromatography: Final Laccase Purification
3.8 Electrophoresis
4 Notes
References
Chapter 13: Bioremediation of Sugarcane Vinasse by Fungi-Based Biological Methods
1 Introduction
2 Materials
2.1 Sugarcane Vinasse
2.2 Fungi
2.3 Fungi Cultivation
2.4 Toxicity Tests
3 Methods
3.1 Sampling and Characterization of Sugarcane Vinasse
3.2 Preparation of Fungi Spores
3.3 Selection of Fungi Culture Conditions
3.4 Vinasse Treatment by Fungi-Based Biological Methods
3.5 Toxicity Assessed of Raw and Treated Vinasse Using Vegetable Cells (Wheat Seeds) as the Indicator Organism
3.6 Toxicity Assessed of Raw and Treated Vinasse Using Mammalian Cells (Caco-2 Cell Line) as the Indicator Organism
4 Notes
5 Conclusion
References
Chapter 14: Two-Dimensional Gel Electrophoresis: Discovering Isoenzymes for Mycoremediation
1 Introduction
2 Materials
2.1 Fungal Growth and Supernatant Obtention
2.2 Sample Preparation and Two-Dimensional Gel Electrophoresis
2.3 Staining Step
3 Methods
3.1 Microbial Growth and Protein Extraction
3.2 Preparation of Molecular Weight Marker
3.3 Second-Dimensional Gel Electrophoresis on SDS-PAGE
3.4 Staining Step
3.5 Measurement of Protein Sizes
4 Notes
References
Chapter 15: Measurement of Ligninolytic Enzymes of Soil Treated with Bioaugmentation
1 Introduction
2 Materials
2.1 Fungal Maintenance
2.2 Bioaugmentation
2.3 Enzymatic Extraction from Bioaugmented Soil
2.4 Oxidative Activity Determination
2.5 Hydrolytic Activity Determination
3 Methods
3.1 Bioaugmentation
3.2 Enzymatic Extraction
3.3 Oxidative Activity Determination
3.4 Hydrolytic Activity Determination
4 Notes
References
Chapter 16: Whole Shotgun Proteomics and Its Role in Mycoremediation
1 Introduction
2 Materials
2.1 Sample Preparation
2.2 Protein Quantification
2.3 HPLC-MS/MS Analysis
3 Methods
3.1 Sample Preparation
3.2 Protein Quantification
3.3 Mass Spectrometry and Analysis
4 Notes
References
Chapter 17: Isolation and Selection of Tolerant Fungal Strains from Soil Polluted with Heavy Metals
1 Introduction
2 Materials
2.1 Soil Sample Collection
2.2 Determination of Total Chromium in Contaminated Soil
2.3 Isolation of Chromium-Tolerant Fungi
2.3.1 Preparation of Antibiotic Supplemented Isolation Medium: Modified Lee´s Minimal Medium
2.3.2 Fungal Isolation from Soil
2.4 Morphological Identification of Isolated Fungal Strains
2.5 Fungal Tolerance Analysis
2.5.1 Plate Screening Technique to Measure Fungi Tolerance
3 Methods
3.1 Soil Sample Collection
3.2 Determination of Total Chromium in Polluted Soil
3.3 Isolation of Chromium-Tolerant Fungi
3.3.1 Preparation of Antibiotic Supplemented Isolation Medium: Modified Lee´s Minimal Medium
3.3.2 Fungal Isolation from Soil
3.4 Morphological Identification of Isolated Fungal Strains
3.5 Fungal Tolerance Analysis
3.5.1 Plate Screening Technique to Measure Fungi Tolerance
3.5.2 Fungal Tolerance Determination
4 Notes
References
Chapter 18: Mycoremediation of Wastewater by Fungal Lipases
1 Introduction
2 Materials
2.1 Obtaining Supernatant Rich in Lipase Enzyme
2.2 Application of Lipase-Rich Supernatant in Water Contaminated with Fats and Oils
3 Methods
3.1 Reactivation of Fungal Strain on Agar
3.2 Obtaining Supernatant Rich in Lipase Enzyme from Penicillium Fungus
3.3 Effluent Treatment with Penicillium Lipase-Rich Supernatant
3.3.1 Sampling and Preservation
3.3.2 Mycoremediation Treatment
3.3.3 Determination of Oils and Fats
4 Notes
References
Chapter 19: Bioaugmentation of Biomixtures with Consortia of Actinobacteria and Fungi for Improving Pesticides Removal
1 Introduction
2 Materials
2.1 Microorganisms
2.2 Culture Media
2.3 Components of the Biomixture
2.4 Pesticide
3 Methods
3.1 Evaluation of Antagonism Among Actinobacteria Strains and Fungi
3.2 Inoculum Preparation
3.3 Biomixtures Preparation
3.4 Microbial Counts
3.5 Lindane Analysis
3.6 Kinetic Parameters of Lindane Removal
4 Notes
References
Chapter 20: Predictive Mycology for the Screening of White-Rot Fungi
1 Introduction
2 Materials
2.1 Maintenance of Fungi
2.2 Tolerance to PCBs
2.3 Ligninolytic Enzyme Assay
3 Methods
3.1 Tolerance to PCBs
3.2 Ligninolytic Enzyme Assay
3.3 Modelling and Statistical Analysis
4 Notes
References
Chapter 21: Enzyme Biosensors for the Detection of Environmental Contaminants
1 Introduction
2 Materials
2.1 Obtaining the Laccase Enzyme
2.2 Conformation of the Enzymatic Biosensor
3 Methods
3.1 Reactivation of the Fungus on Agar
3.2 Obtaining the Fungus Supernatant
3.3 Preparation of the Enzyme To Be Used in the Biosensor
3.4 Preparation and Loading of Screen-Printed Electrodes for Measurement
3.5 Electrochemical Measurements
4 Notes
References
Index
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Dhanushka Udayanga Pankaj Bhatt Dimuthu Manamgoda Juliana Maria Saez Editors
Mycoremediation Protocols
SPRINGER PROTOCOLS HANDBOOKS
For further volumes: http://www.springer.com/series/8623
Springer Protocols Handbooks collects a diverse range of step-by-step laboratory methods and protocols from across the life and biomedical sciences. Each protocol is provided in the Springer Protocol format: readily-reproducible in a step-by-step fashion. Each protocol opens with an introductory overview, a list of the materials and reagents needed to complete the experiment, and is followed by a detailed procedure supported by a helpful notes section offering tips and tricks of the trade as well as troubleshooting advice. With a focus on large comprehensive protocol collections and an international authorship, Springer Protocols Handbooks are a valuable addition to the laboratory.
Mycoremediation Protocols Edited by
Dhanushka Udayanga Department of Biosystems Technology, Faculty of Technology, University of Sri Jayewardenepura, Pitipana, Homagama, Sri Lanka
Pankaj Bhatt Integrative Microbiology Research Centre, South China Agricultural University, Guangzhou, China
Dimuthu Manamgoda Department of Botany, Faculty of Applied Sciences, University of Sri Jayewardenepura, Gangodawila, Nugegoda, Sri Lanka
Juliana Maria Saez Universidad Nacional de Tucumán, Tucumán, Argentina
Editors Dhanushka Udayanga Department of Biosystems Technology, Faculty of Technology University of Sri Jayewardenepura Pitipana, Homagama, Sri Lanka Dimuthu Manamgoda Department of Botany, Faculty of Applied Sciences University of Sri Jayewardenepura Gangodawila, Nugegoda, Sri Lanka
Pankaj Bhatt Integrative Microbiology Research Centre South China Agricultural University Guangzhou, China Juliana Maria Saez Universidad Nacional de Tucuma´n Tucuma´n, Argentina
ISSN 1949-2448 ISSN 1949-2456 (electronic) Springer Protocols Handbooks ISBN 978-1-0716-2005-2 ISBN 978-1-0716-2006-9 (eBook) https://doi.org/10.1007/978-1-0716-2006-9 © The Editor(s) (if applicable) and The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022 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 Fungi are a highly diverse and ubiquitous group of microorganisms with a wide range of applications in biotechnology. The species of fungi have been traditionally used in sustainable agriculture, medicine, and the food industry since ancient times. Mycoremediation is the form of bioremediation, which primarily exploits fungi to remove environmental pollutants. Fungi are well known as the major group of microorganisms that naturally degrade organic matter in the environment. However, during the past few decades, the use of fungi to restore polluted environments has emerged as a promising approach in environmental biotechnology. Mycoremediation is an ecofriendly and cost-effective process to remediate a vast range of environmental pollutants. The selection of an appropriate fungus as well as an effective method of application is equally important for the success of a remediation effort. Therefore, the studies on mycoremediation essentially focus on fungal isolation, selection, process optimization, and effective implementation. However, it has been difficult to find comprehensive protocols for researchers to screen suitable fungal species and methods of effective implementation. Therefore, this book aims to provide a well-grounded practical approach for aspiring researchers toward the advancement of mycoremediation studies. The first six chapters (1–6) cover the screening of fungi that can be used to remediate organic contaminants such as polyaromatic hydrocarbons (PAHs), textile dyes, pesticides, and heavy metals. In the next eight chapters (7–14), the protocols for advanced aspects of mycoremediation are provided, including the use of bioreactors, molecular methods to estimate the diversity, and the use of fungi to remediate complex contaminants such as polyethylene and radioactive waste. The latter part of the book which includes chapters 15–20 aim to provide insights into predictive mycology and proteomics approaches to select fungi and elucidating biological mechanisms in the mycoremediation process. The final chapter (Chapter 21) describes the use of fungal laccase enzyme-based biosensors for the detection of environmental contaminants. Accordingly, this book covers a wide range of aspects related to mycoremediation which can be applied for both basic and advanced multidisciplinary research. We greatly appreciate all the authors around the world for their invaluable contribution to complete this project. We hope that the techniques included in this book will be useful toward a comprehensive understanding of this innovative area in science ensuring better and sustainable living on earth through scientific research. We invite the users of this book to follow these protocols accordingly in their endeavors in teaching and research. Pitipana, Homagama, Sri Lanka Guangzhou, China Nugegoda, Sri Lanka Tucuma´n, Argentina
Dhanushka Udayanga Pankaj Bhatt Dimuthu Manamgoda Juliana Maria Saez
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Contents Preface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Contributors. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
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1 Isolation, Enrichment, and Characterization of Fungi for the Degradation of Organic Contaminants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1 Saurabh Gangola, Pankaj Bhatt, Samiksha Joshi, Narendra Singh Bhandari, Saurabh Kumar, Om Prakash, Avikal Kumar, Samarth Tewari, and Deepa Nainwal 2 Protocol for the Assessment of Mycoremediation of Polycyclic Aromatic Hydrocarbons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 13 ˜ a Mayans, Carlos Garcı´a-Delgado, Raquel Camacho-Are´valo, Begon Rafael Anto n-Herrero, and Enrique Eymar 3 Characterization and Screening of Pesticide-Degrading Indigenous Fungi from Soil and Water . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 25 Geeta Bhandari 4 Mycoremediation of Synthetic Textile Dyes by Fungi Isolated from Textile Wastewater Effluent and Soil. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 31 Manavi S. Ekanayake, Dhanushka Udayanga, and Pathmalal M. Manage 5 Protocol for Screening Endophytic Fungi Against Heavy Metals . . . . . . . . . . . . . . 45 ¨ ller Jenny Choo, Changi Wong, and Moritz Mu 6 Screening of Phyllosphere Fungi Inhabiting in Urbanized Areas for Phylloremediation Capabilities of Polyaromatic Hydrocarbon Pollutants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 55 L. J. S. Undugoda, R. B. N. Dharmasiri, Dhanushka Udayanga, and N. N. R. N. Nugara 7 Methods for Design and Bioremediation Applications of Reactors Based on Immobilized Fungi . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 71 ˇ eneˇk Novotny´, Kamila Sˇre´dlova´, Toma´ˇs Cajthaml, and Pavel Hasal C 8 Denaturing Gradient Gel Electrophoresis (DGGE) Analysis of the Fungi Involved in Biodegradation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 93 Saurabh Kumar, Divya Joshi, Prasenjit Debbarma, Manali Singh, Ajar Nath Yadav, Nasib Singh, Deep Chandra Suyal, Ravindra Soni, and Reeta Goel 9 Optimization of Mycoremediation Process for the Isolated Fungi. . . . . . . . . . . . . 101 Pankaj Bhatt and Shaohua Chen 10 Protocol for Assessing Mycoremediation of Acidic Radioactive Wastes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 109 Yehia A. -G. Mahmoud and Osama M. Darwesh 11 Protocol for Screening Low-Density Polyethylene (LDPE)–Degrading Soil Fungi Isolated from Urban Waste Dumping Sites . . . . . . . . . . . . . . . . . . . . . . . 123 Gimhani D. Ramanayake, Dhanushka Udayanga, L. J. S. Undugoda, N. N. R. N. Nugara, A. H. L. R. Nilmini, and Pathmalal M. Manage
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Contents
Production of Laccases from Agricultural Wastes: Strain Isolation and Selection, Enzymatic Profiling, and Lab-Scale Production. . . . . . . . . . . . . . . . Pablo M. Ahmed, Hipolito F. Pajot, and Pablo M. Ferna´ndez Bioremediation osf Sugarcane Vinasse by Fungi-Based Biological Methods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Luciana Melisa Del Gobbo, Macarena Marı´a Rulli, Liliana Beatriz Villegas, and Vero nica Leticia Colin Two-Dimensional Gel Electrophoresis: Discovering Isoenzymes for Mycoremediation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . ˜ oski, Laura Ester Ortellado, Marı´a Isabel Fonseca, Marcela Alejandra Sadan Silvana Florencia Benı´tez, and Pedro Darı´o Zapata Measurement of Ligninolytic Enzymes of Soil Treated with Bioaugmentation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . ˜ oski, Ana Silvia Tatarin, Marcela Alejandra Sadan and Laura Lidia Villalba Whole Shotgun Proteomics and Its Role in Mycoremediation . . . . . . . . . . . . . . . . Anibal Sebastian Chelaliche, Adriana Elizabet Alvarenga, Pedro Darı´o Zapata, and Marı´a Isabel Fonseca Isolation and Selection of Tolerant Fungal Strains from Soil Polluted with Heavy Metals . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . ˜ oski, Marta Alejandra Polti, Ana Silvia Tatarin, Marcela Alejandra Sadan and Marı´a Isabel Fonseca Mycoremediation of Wastewater by Fungal Lipases . . . . . . . . . . . . . . . . . . . . . . . . . Laura Ester Ortellado, Laura Lidia Villalba, Pedro Darı´o Zapata, and Marı´a Isabel Fonseca Bioaugmentation of Biomixtures with Consortia of Actinobacteria and Fungi for Improving Pesticides Removal . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Enzo E. Raimondo, Ana L. Bigliardo, Samanta K. Gonza´lez, Juliana M. Saez, Marta A. Polti, and Claudia S. Benimeli Predictive Mycology for the Screening of White-Rot Fungi . . . . . . . . . . . . . . . . . . ˜ oski, Juan Ernesto Vela´zquez, Marcela Alejandra Sadan and Laura Lidia Villalba Enzyme Biosensors for the Detection of Environmental Contaminants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Alan Rolando Ayala Schimpf, Daniela Rodrı´guez, Marı´a Isabel Fonseca, and Pedro Darı´o Zapata
Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
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Contributors PABLO M. AHMED • ITANOA–CONICET, EEAOC, Las Talitas, Tucuma´n, Argentina ADRIANA ELIZABET ALVARENGA • Laboratorio de Biotecnologı´a Molecular, Instituto de Biotecnologı´a Misiones, CONICET, Facultad de Ciencias Exactas, Quı´micas y Naturales, Universidad Nacional de Misiones, Posadas, Misiones, Argentina RAFAEL ANTO´N-HERRERO • Department of Agricultural Chemistry and Food Sciences, Universidad Autonoma de Madrid, Madrid, Spain CLAUDIA S. BENIMELI • Planta Piloto de Procesos Industriales Microbiologicos (PROIMICONICET), Avenida Belgrano y Pasaje Caseros, Tucuma´n, Argentina; Facultad de Ciencias Exactas y Naturales, Universidad Nacional de Catamarca, Catamarca, Argentina SILVANA FLORENCIA BENI´TEZ • Facultad de Ciencias Exactas, Quı´micas y Naturales, Laboratorio de Biotecnologı´a Molecular (BIOTECMOL), Instituto de Biotecnologı´a Misiones “Dra. Marı´a Ebe Reca” (INBIOMIS), Universidad Nacional de Misiones, Posadas, Misiones, Argentina; CONICET, Misiones, Argentina GEETA BHANDARI • Department of Biochemistry and Biotechnology, Sardar Bhagwan Singh University, Dehradun, Uttarakhand, India NARENDRA SINGH BHANDARI • School of Agriculture, Graphic Era Hill University, Bhimtal, India PANKAJ BHATT • State Key Laboratory for Conservation and Utilization of Subtropical Agrobioresources, Guangdong Province Key Laboratory of Microbial Signals and Disease Control, Integrative Microbiology Research Centre, South China Agricultural University, Guangzhou, China; Guangdong Laboratory for Lingnan Modern Agriculture, Guangzhou, China ANA L. BIGLIARDO • Planta Piloto de Procesos Industriales Microbiologicos (PROIMICONICET), Avenida Belgrano y Pasaje Caseros, Tucuma´n, Argentina TOMA´Sˇ CAJTHAML • Institute of Microbiology of the Czech Academy of Sciences, Laboratory of Environmental Biotechnology, Prague, Czech Republic; Institute for Environmental Studies, Faculty of Science, Charles University, Prague, Czech Republic RAQUEL CAMACHO-ARE´VALO • Department of Agricultural Chemistry and Food Sciences, Universidad Autonoma de Madrid, Madrid, Spain ANIBAL SEBASTIAN CHELALICHE • Laboratorio de Biotecnologı´a Molecular, Instituto de Biotecnologı´a Misiones, CONICET, Facultad de Ciencias Exactas, Quı´micas y Naturales, Universidad Nacional de Misiones, Posadas, Misiones, Argentina SHAOHUA CHEN • State Key Laboratory for Conservation and Utilization of Subtropical Agro-bioresources, Guangdong Province Key Laboratory of Microbial Signals and Disease Control, Integrative Microbiology Research Centre, South China Agricultural University, Guangzhou, China; Guangdong Laboratory for Lingnan Modern Agriculture, Guangzhou, China JENNY CHOO • Faculty of Engineering, Computing and Science, Swinburne University of Technology, Kuching, Sarawak, Malaysia VERO´NICA LETICIA COLIN • Planta Piloto de Procesos Industriales Microbiologicos (PROIMICONICET), Tucuma´n, Argentina
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Contributors
OSAMA M. DARWESH • Agriculture Microbiology Department, National Research Centre, Cairo, Egypt PRASENJIT DEBBARMA • School of Agriculture, Graphic Era Hill University, Dehradun, Uttarakhand, India R. B. N. DHARMASIRI • Department of Biosystems Technology, Faculty of Technology, University of Sri Jayewardenepura, Pitipana, Homagama, Sri Lanka MANAVI S. EKANAYAKE • Centre for Water Quality and Algae Research, Department of Zoology, University of Sri Jayewardenepura, Nugegoda, Sri Lanka; Faculty of Graduate Studies, University of Sri Jayewardenepura, Nugegoda, Sri Lanka ENRIQUE EYMAR • Department of Agricultural Chemistry and Food Sciences, Universidad Autonoma de Madrid, Madrid, Spain PABLO M. FERNA´NDEZ • PROIMI–CONICET, SM de Tucuma´n, Tucuma´n, Argentina; FACEN, UNCA, SFV de Catamarca, Catamarca, Argentina MARI´A ISABEL FONSECA • Laboratorio de Biotecnologı´a Molecular, Instituto de Biotecnologı´a Misiones, CONICET, Facultad de Ciencias Exactas, Quı´micas y Naturales, Universidad Nacional de Misiones, Posadas, Misiones, Argentina; Universidad Nacional de Misiones. Facultad de Ciencias Exactas Quı´micas y Naturales, Instituto de Biotecnologı´a Misiones “Dra. Marı´a Ebe Reca” (INBIOMIS). Laboratorio de Biotecnologı´a Molecular (BIOTECMOL), Misiones, Argentina; Facultad de Ciencias Exactas, Quı´micas y Naturales, Laboratorio de Biotecnologı´a Molecular (BIOTECMOL), Instituto de Biotecnologı´a Misiones “Dra. Marı´a Ebe Reca” (INBIOMIS), Universidad Nacional de Misiones, Posadas, Misiones, Argentina; CONICET, Buenos Aires, Argentina SAURABH GANGOLA • School of Agriculture, Graphic Era Hill University, Bhimtal, India CARLOS GARCI´A-DELGADO • Department of Geology and Geochemistry, Universidad Autonoma de Madrid, Madrid, Spain LUCIANA MELISA DEL GOBBO • Planta Piloto de Procesos Industriales Microbiologicos (PROIMI-CONICET), Tucuma´n, Argentina REETA GOEL • GLA University, Mathura, Chaumuhan, Uttar Pradesh, India SAMANTA K. GONZA´LEZ • Planta Piloto de Procesos Industriales Microbiologicos (PROIMICONICET), Avenida Belgrano y Pasaje Caseros, Tucuma´n, Argentina PAVEL HASAL • Department of Chemical Engineering, University of Chemistry and Technology, Prague, Prague, Czech Republic DIVYA JOSHI • Uttarakhand Pollution Control Board, Regional Office, Kashipur, Uttarakhand, India SAMIKSHA JOSHI • School of Agriculture, Graphic Era Hill University, Bhimtal, India AVIKAL KUMAR • School of Agriculture, Graphic Era Hill University, Bhimtal, India SAURABH KUMAR • Division of Crop Research, ICAR-Research Complex for Eastern Region, Patna, Bihar, India YEHIA A. -G. MAHMOUD • Botany Department, Mycology Research Lab., Faculty of Science, Tanta University, Tanta, Egypt PATHMALAL M. MANAGE • Centre for Water Quality and Algae Research, Department of Zoology, Faculty of Applied Sciences, University of Sri Jayewardenepura, Nugegoda, Sri Lanka; Faculty of Graduate Studies, University of Sri Jayewardenepura, Nugegoda, Sri Lanka BEGON˜A MAYANS • Department of Agricultural Chemistry and Food Sciences, Universidad Autonoma de Madrid, Madrid, Spain MORITZ MU¨LLER • Faculty of Engineering, Computing and Science, Swinburne University of Technology, Kuching, Sarawak, Malaysia
Contributors
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DEEPA NAINWAL • School of Agriculture, Graphic Era Hill University, Bhimtal, India A. H. L. R. NILMINI • Department of Material and Mechanical Technology, Faculty of Technology, University of Sri Jayewardenepura, Pitipana, Homagama, Sri Lanka ˇ ENEˇK NOVOTNY´ • Institute of Microbiology of the Czech Academy of Sciences, Laboratory of C Environmental Biotechnology, Prague, Czech Republic; Czech University of Life Sciences, Prague, Prague, Czech Republic N. N. R. N. NUGARA • Department of Biosystems Technology, Faculty of Technology, University of Sri Jayewardenepura, Pitipana, Homagama, Sri Lanka LAURA ESTER ORTELLADO • Laboratorio de Biotecnologı´a Molecular, Instituto de Biotecnologı´a Misiones “Dra. Marı´a Ebe Reca” (INBIOMIS), CONICET, Facultad de Ciencias Exactas, Quı´micas y Naturales, Universidad Nacional de Misiones, Posadas, Misiones, Argentina; CONICET, Buenos Aires, Argentina HIPO´LITO F. PAJOT • PROIMI–CONICET, SM de Tucuma´n, Tucuma´n, Argentina; FACEN, UNCA, SFV de Catamarca, Catamarca, Argentina MARTA ALEJANDRA POLTI • Planta Piloto de Procesos Industriales Microbiologicos (PROIMI), CONICET, Tucuma´n, Argentina; Facultad de Ciencias Naturales e Instituto Miguel Lillo, Universidad Nacional de Tucuma´n, Tucuma´n, Argentina OM PRAKASH • Department of Chemistry, GB Pant University of Agriculture and Technology, Pantnagar, India ENZO E. RAIMONDO • Planta Piloto de Procesos Industriales Microbiologicos (PROIMICONICET), Avenida Belgrano y Pasaje Caseros, Tucuma´n, Argentina; Facultad de Bioquı´mica, Quı´mica y Farmacia, Universidad Nacional de Tucuma´n, Tucuma´n, Argentina GIMHANI D. RAMANAYAKE • Department of Biosystems Technology, Faculty of Technology, University of Sri Jayewardenepura, Pitipana, Homagama, Sri Lanka; Centre for Water Quality and Algae Research, Department of Zoology, Faculty of Applied Sciences, University of Sri Jayewardenepura, Nugegoda, Sri Lanka; Faculty of Graduate Studies, University of Sri Jayewardenepura, Nugegoda, Sri Lanka DANIELA RODRI´GUEZ • Laboratorio de Biotecnologı´a Molecular, Instituto de Biotecnologı´a Misiones, Facultad de Ciencias Exactas, Quı´micas y Naturales, Universidad Nacional de Misiones, Posadas, Misiones, Argentina MACARENA MARI´A RULLI • Planta Piloto de Procesos Industriales Microbiologicos (PROIMICONICET), Tucuma´n, Argentina MARCELA ALEJANDRA SADAN˜OSKI • Laboratorio de Biotecnologı´a Molecular, Instituto de Biotecnologı´a Misiones “Dra. Marı´a Ebe Reca” (INBIOMIS), CONICET, Facultad de Ciencias Exactas, Quı´micas y Naturales, Universidad Nacional de Misiones, Posadas, Misiones, Argentina; Universidad Nacional de Misiones. Facultad de Ciencias Exactas Quı´micas y Naturales, Instituto de Biotecnologı´a Misiones “Dra. Marı´a Ebe Reca” (INBIOMIS). Laboratorio de Biotecnologı´a Molecular (BIOTECMOL), Misiones, Argentina; CONICET, Buenos Aires, Argentina JULIANA M. SAEZ • Planta Piloto de Procesos Industriales Microbiologicos (PROIMICONICET), Avenida Belgrano y Pasaje Caseros, Tucuma´n, Argentina; Facultad de Ciencias Naturales e Instituto Miguel Lillo, Universidad Nacional de Tucuma´n, Tucuma´n, Argentina ALAN ROLANDO AYALA SCHIMPF • Laboratorio de Biotecnologı´a Molecular, Instituto de Biotecnologı´a Misiones, Facultad de Ciencias Exactas, Quı´micas y Naturales, Universidad Nacional de Misiones, Posadas, Misiones, Argentina
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Contributors
MANALI SINGH • Department of Biotechnology, Invertis University, Bareilly, Uttar Pradesh, India NASIB SINGH • Department of Microbiology, Akal College of Basic Sciences, Eternal University, Baru Sahib, Himachal Pradesh, India RAVINDRA SONI • Department of Agricultural Microbiology, College of Agriculture, Indira Gandhi Krishi Vishwa Vidyalaya, Raipur, Chhattisgarh, India KAMILA SˇRE´DLOVA´ • Institute of Microbiology of the Czech Academy of Sciences, Laboratory of Environmental Biotechnology, Prague, Czech Republic; Institute for Environmental Studies, Faculty of Science, Charles University, Prague, Czech Republic DEEP CHANDRA SUYAL • Department of Microbiology, Akal College of Basic Sciences, Eternal University, Baru Sahib, Himachal Pradesh, India ANA SILVIA TATARIN • Laboratorio de Biotecnologı´a Molecular, Instituto de Biotecnologı´a Misiones “Dra. Marı´a Ebe Reca” (INBIOMIS), CONICET, Facultad de Ciencias Exactas, Quı´micas y Naturales, Universidad Nacional de Misiones, Posadas, Misiones, Argentina; CONICET, Buenos Aires, Argentina; Universidad Nacional de Misiones. Facultad de Ciencias Exactas Quı´micas y Naturales, Instituto de Biotecnologı´a Misiones “Dra. Marı´a Ebe Reca” (INBIOMIS). Laboratorio de Biotecnologı´a Molecular (BIOTECMOL), Misiones, Argentina SAMARTH TEWARI • School of Agriculture, Graphic Era Hill University, Bhimtal, India DHANUSHKA UDAYANGA • Department of Biosystems Technology, Faculty of Technology, University of Sri Jayewardenepura, Pitipana, Homagama, Sri Lanka L. J. S. UNDUGODA • Department of Biosystems Technology, Faculty of Technology, University of Sri Jayewardenepura, Pitipana, Homagama, Sri Lanka JUAN ERNESTO VELA´ZQUEZ • Laboratorio de Biotecnologı´a Molecular, Instituto de Biotecnologı´a Misiones “Dra. Marı´a Ebe Reca” (INBIOMIS), CONICET, Universidad Nacional de Misiones, Posadas, Misiones, Argentina; CONICET, Buenos Aires, Argentina LAURA LIDIA VILLALBA • Laboratorio de Biotecnologı´a Molecular, Instituto de Biotecnologı´a Misiones “Dra. Marı´a Ebe Reca” (INBIOMIS), CONICET, Universidad Nacional de Misiones, Posadas, Misiones, Argentina; CONICET, Buenos Aires, Argentina LILIANA BEATRIZ VILLEGAS • Instituto de Quı´mica San Luis (INQUISAL-CONICET), San Luis, Argentina CHANGI WONG • Faculty of Engineering, Computing and Science, Swinburne University of Technology, Kuching, Sarawak, Malaysia AJAR NATH YADAV • Department of Biotechnology, DKSG Akal College of Agriculture, Eternal University, Baru Sahib, Himachal Pradesh, India PEDRO DARI´O ZAPATA • Laboratorio de Biotecnologı´a Molecular, Instituto de Biotecnologı´a Misiones “Dra. Marı´a Ebe Reca” (INBIOMIS), CONICET, Universidad Nacional de Misiones, Posadas, Misiones, Argentina; CONICET, Buenos Aires, Argentina; Laboratorio de Biotecnologı´a Molecular, Instituto de Biotecnologı´a Misiones, CONICET, Facultad de Ciencias Exactas, Quı´micas y Naturales, Universidad Nacional de Misiones, Posadas, Misiones, Argentina
Chapter 1 Isolation, Enrichment, and Characterization of Fungi for the Degradation of Organic Contaminants Saurabh Gangola, Pankaj Bhatt, Samiksha Joshi, Narendra Singh Bhandari, Saurabh Kumar, Om Prakash, Avikal Kumar, Samarth Tewari, and Deepa Nainwal Abstract Fungal metabolism is more effective and efficient than bacterial metabolism for bioremediation of organic pollutants. The exploitation of fungi for the degradation of organic pollutants is an economic and eco-friendly technique. Fungi produces several oxidative enzymes or secondary metabolites, which help in the biodegradation of organic contaminants from the contaminated areas. This study is based on the isolation and purification of indigenous fungal isolates from contaminated environments followed by acclimatization of fungal isolates at different increasing concentrations of contaminants to enhance the metabolic potential of fungi. In addition, morphological and molecular characterization of isolated fungal strain is also explained, which is useful for the proper utilization of identified organisms in contaminated environments and in their value addition. Key words Fungi, Isolation, Biodegradation, Pesticide, Electrophoresis
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Introduction Rapid industrialization process and indiscriminate application of agricultural chemicals (pesticides and fertilizers) has been polluting our environment continuously for decades of years. Generally, a specific group of chemicals such as chlorinated aliphatic hydrocarbons and aromatic hydrocarbons are more recalcitrant in nature and polluting the environment. Since the start of the industrial revolution, these xenobiotic compounds come to contact with the natural environment by leakage, improper means of disposal, or by accidental [1, 2]. Due to their accumulation, persistence, and toxic nature, these compounds are very harmful to the environment and also cause many human health problems. Recently, the United States Environmental Protection Agency (EPA) has released a list of 129 compounds that are potentially hazardous for the
Dhanushka Udayanga et al. (eds.), Mycoremediation Protocols, Springer Protocols Handbooks, https://doi.org/10.1007/978-1-0716-2006-9_1, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022
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environment and kept them in the priority pollutant category. In that list, mainly chlorinated aliphatic and aromatic chemical compounds are enlisted. To maximize crop production for the fulfilment of fastgrowing population food demand, the farmers are using pesticides for managing the different groups of pests in modern agricultural practices. Statistical data of the accurate usage of agricultural pesticides are much harder to obtain for many of the developing countries comprising the bulk of the tropical landmass compared to well-regulated North Europe and Japanese markets. Therefore, it is crucial to study the fate of pesticides in that area where these are applied regularly. Contamination of the environment due to excessive use of pesticides in agricultural fields or in households now is of great concern today [2]. Because of the toxic nature of pesticides that affect nontargeted organisms, offsite mobility negatively affects soil microbial diversity, and soil fertility, which has become a great matter of environmental concern to study the fate of pesticides in the soil as well as in water ecosystems [3]. The presence of pesticide residues, such as endosulfan, HCH (Hexachlorocyclohexane), and DDT (DichloroDiphenylTrichloroethane), are also observed in mineral water and in soft drinks: it is a matter of serious concern. Generally, an ideal pesticide should be nontoxic to the nontargeted organisms, should not contaminate the groundwater by leaching from the soil, and should be biodegradable. Unfortunately, the widespread indiscriminate use of pesticides in modern agriculture is of great concern [4]. Besides the benefits of pesticides, these are producing several kinds of toxic effects, which become potential threats to the surrounding environment. The excessive use of these agricultural chemicals causes severe consequences because of their potential runoff and leaching into surface and groundwater. Therefore, the contamination of the environment with chemicals is a major issue of concern. Cleaning of pesticide-contaminated sites has been a direct concern of the environment since the establishment of the industrial era. Some natural processes are employed for the removal of organic pollutants from the environments like biological transformation, dilution, volatilization, sorption, and photolysis. However, all the mentioned natural processes can reduce the level of environmental risk to some extent, but only chemical and biological transformation can alter the structure and nature of the pollutants and convert it into the nontoxic form or environmentally accepted form. Under existing environmental conditions, chemical transformation is a very slow process that takes several decades of years [2]. Indiscriminate use of pesticides resulted in the accumulation of the pesticide residues in the ecosystem, causing serious health problems to human beings and other environmental organisms. Therefore, the utilization of microbial systems for the removal of toxic pollutants from the environment can be the possible solution
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by means of biodegradation and bioremediation. The microbial systems reduce the concentration of toxic xenobiotic compounds by cleaning and convert them into nontoxic chemical compounds and help to restore the natural condition [5]. The ultimate goal of bioremediation is to remediate or clean the toxic environment, protect biodiversity (soil habitats and microbial composition) and human resources for their consumption (maintain soil fertility and groundwater purity). Nature has many natural remediation processes like hydrolysis, accumulation of toxic compounds, sorption, volatilization that are actively involved in the degradation of toxic compounds and respond differently at different environmental conditions. By considering microbial systems, it is possible to develop potential strategies for pesticide removal and their derivative residue from the contaminated area. Microorganisms present in polluted environments generally perform well by utilizing these toxic chemicals as a source of energy, carbon, and nitrogen for their growth. Generally, biodegradation is defined as the breakdown of any complex chemical compound into simple chemical compounds by the means of living microorganisms. Biodegradation is totally based on the growth and cometabolism of microorganisms. The activity of microbes can be enhanced in the natural environment by adding some amendments in soil or by adding some stimulant for increasing the rate of degradation within a limited period of time. Several numbers of bacteria and fungi are reported, which are actively involved in the pesticide degradation. Organic compounds along with the growth substrate are metabolized by the process of cometabolism, where the growth substrate is used as the primary source of carbon and energy. A group of microorganisms having different genetic bases serve as highly efficient microbial degradative tools because mixed microbial community can degrade the intermediate metabolites of the others, hence at the last complete mineralization occurs. Certain factors such as genetic potential of the microorganism and environmental factors like pH, temperature, available nitrogen, and phosphorus can affect the rate of pesticide degradation. In nature, the biodegradation generally proceeds by the indigenous microorganisms; however, the process can be stimulated by adding some nutrients or electron acceptors (biostimulation) or by adding some specific degrader microorganism (bioaugmentation) [6]. In case of bioaugmentation process, the big challenge for the introduced microorganism includes saving themselves from the predators and compete with the existing microbial community. Therefore, the success of bioaugmentation process is not only relying on the degradation ability of microorganisms but also depends on the survival, growth, and distribution of introduced microorganisms. Due to the fast metabolic activity, fungi alone or in combination with bacteria could be a better approach to overcome such challenges. The application of fungi for biodegradation
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purposes is known as mycoremediation. Several fungal species are reported for their high concentration tolerance and fast rate of pesticide metabolism [7]. Most of the biodegradation studies are associated with potential of specific bacterial isolates. Although, the filamentous fungi having the important characteristic can be applied in heterogeneous environments and advantageous for biodegradation study. However, the fungi are nonmotile in nature, but they can respond and adapt themselves according to change in the environmental condition quickly and survive [8]. There are some abiotic factors such as pH, environmental temperature, water potential, oxygen accessibility, and nutrient status, which mainly affect the growth of fungal mycelium [9]. Fungal hyphae are capable enough to penetrate solids and can access the microhabitat such as water-filled micropores in soil [10]. Therefore, the use of fungal isolate for biodegradation study is very advantageous than the bacteria, because the hyphae of the fungi can reach much better to the nutrients or to the pollutants, which are distributed heterogeneously or inaccessible to the bacteria [11]. Moreover, the fungal hyphae also help in the growth of fungi by transporting nutrients in different regions of mycelia [12].
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Materials Agar, potato dextrose broth (PDB), pesticides, double distilled water (DDW), PCR primers, DNTPs, lactophenol cotton blue, DNA polymerase, MgCl2, agarose, ethidium bromide (EtBr), stock solution of pesticide (1 mg/mL), ethanol (70%), glass slides, Petri plates, coverslips, cork borer, mounting needles, spirit lamp or Bunsen burner, compound light microscope, laminar air flow, centrifuge, thermocycler (PCR machine), electrophoresis apparatus, sampling bags, and autoclave.
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Screening of Pesticide-Degrading Fungal Strains
3.1 Isolation of the Fungal Strains
1. Collect soil samples from pesticide-contaminated agricultural land in polythene sampling bags and store them at 20 C till further use. 2. Prepare microbial inoculums by shaking 20 g of soil in 100 mL potato dextrose broth and incubate overnight at 28 C and 150 rev min1. Filter the soil suspension and collect the supernatant in a sterile flask. 3. Make the required concentration (generally 10 or 20 ppm) of pesticides from the stock solution. 4. Prepare potato dextrose agar (250 mL) in a 500 mL flask.
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5. In the next step, add 1 mL of supernatant in different Petri plates (autoclaved). 6. Pour autoclaved PDA (lukewarm) to all the Petri plates and mix gently. 7. After that add the required concentration of pesticide (10–20 ppm) to each plate and mix it properly for homogenization of the mixture. Further, leave the plates for solidification. 8. Prepare two sets of control, the first set as negative control having medium supplemented with pesticide and without inoculation of supernatant while another set as a positive control with medium inoculated with supernatant and without the addition of pesticide. 9. Incubate the inoculated plates for 5 to 10 days at 28 2 C. 10. Obtain fungal colonies further enriched by transferring to a higher concentration of pesticides in the same medium. Recovered fungal isolates are further purified by growing them on Czapek–Dox medium. The composition of Czapek–Dox medium is (g L1): NaNO3, 3; MgSO4, 0.5; Sucrose, 30; KCl, 0.5; K2HPO4, 1 and FeSO4, 0.01. Maintain the pH (6) of the medium [13]. The source of carbon is pesticide instead of sucrose. Or 1. 1 g of soil sample is suspended in 10 mL of double-distilled water. 2. Make the dilution of microbial suspensions up to 105. 3. Dilution of 103, 104, and 105 can be used to isolate fungi. 4. Add 1 mL of microbial suspension of each dilution to sterile Petri dishes (triplicate of each dilution). 5. Add 15 mL of sterile potato dextrose agar and Czapek–Dox agar supplemented with a required concentration of pesticides in the Petri dishes. 6. One percent streptomycin solution can be added to the medium before pouring into Petri plates for preventing bacterial growth. 7. Incubate the Petri dishes at 28 2 C in dark. 8. Observe the Petri dishes everyday up to 5 days. 9. Obtained fungal colonies are further enriched by transferring to a higher concentration of pesticides in the same medium. Recovered fungal isolates are further purified by growing them on Czapek–Dox medium. The source of carbon is pesticide instead of sucrose.
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3.2 Purification of Fungal Isolates
1. Purify the recovered fungal isolates by inoculating them on solid Czapek–Dox medium. 2. Add 10 ppm of pesticide aseptically to the medium after autoclaving (once the medium has become lukewarm). 3. Pour the molten Czapek–Dox agar media equally into Petri dishes. 4. Cut the piece of agar with the help of a cork borer in which the fungal isolate is growing. 5. Inoculate the fungal disc (6 mm) to the solidified media. Further, incubate the plates under aerobic conditions at 28 C for 48 h. 6. Isolate and purify the fungi showing luxuriant growth by transferring on fresh pesticide supplemented Czapek–Dox medium [14].
3.3 Morphological Characterization of Fungi
The desirable information can be obtained by microscopic observation of examining microorganisms. For the morphological study of microbial cells, generally, two methods are employed: (i) Microscopic observation of living cells (ii) Microscopic observation by staining the cells To study the shape, size, and arrangement of microbial cells, living cells are examined under the microscope. But the observation of living cells sometimes becomes difficult, because the microbial cells are semitransparent in nature; hence, in the absence of a proper microscope, it is difficult to observe.
3.4 Microscopic Observation of Fungi by Temporary Wet Mount Technique
1. Place one drop of double distilled water on the center of a clean glass slide. 2. Inoculate active fungal mycelium to the water drop on the center of the glass slide with the help of inoculating needle. 3. Place the coverslip on the drop of water from the edge of the slide. 4. Gently press the coverslip from over with the help of a pencil. 5. Observe the microbial cells under a microscope by using objective lenses of 20 and 40. 6. Note the observations like shape, size, and characteristics of examined fungal cells include spores, hyphae, conidiospores, septate or aseptate, asexual or sexual stage.
3.5 Mounting of Fungal Cells by Lactophenol Cotton Blue
For the microscopic examination of fungal cells generally, lactophenol cotton blue stain is used. It provides a light blue background by staining fungal cytoplasm, against which the fungal hyphae wall can readily be seen. Mainly this fungal stain has four constituents: (i) phenol, which acts as a fungicide; (ii) lactic acid, which serves as a clearing agent, (iii) cotton blue, which helps to stain blue the cytoplasm of the fungal cell (iv) glycerine, which gives
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a semipermanent preparation. In place of glycerine, by adding polyvinyl alcohol in the mounting medium the permanent preparation can be made. For the examination of dark-colored fungi, lactophenol alone (without the use of cotton blue) can also be used. 1. Place one or two drops of lactophenol cotton blue on a clean glass slide. 2. Inoculate a small amount of well-sporulated fungi into the drop of lactophenol cotton blue on a slide by using a sterilized inoculating needle. 3. Gently the fungal cells are mixed properly in the stain. 4. By avoiding bubbles, place the coverslip over the preparation gently. 5. Press the coverslip gently with the help of a pencil to expel the air bubble and for the spreading of the fungal structure. 6. Observe the preparation under the microscope by using the objective lens of 20 and 40. 7. To save the preparation for a long time, seal the lactophenol mount with coverslip by using nail clear varnish as follows: (a) By providing gentle heat or pressure or by adding additional lactophenol cotton blue, the entrapped air bubble can be removed. (b) Use the 70% ethanol for removing the over flooded stain around the coverslip with the help of blotting paper or cotton swab. (c) Seal the coverslip from the edge by applying a thin layer of nail varnish and allow it to dry overnight. (d) Over the first coat, apply the second coat of nail varnish. The fungal cytoplasm is seen as a lightly stained blue region forming a layer inside the unstained cell wall of hyphae, conidiophores, and conidia that is surrounded by light blue background on the slide. Precautions: (a) Aseptic conditions should be maintained. (b) No air bubbles should be present in the sample preparation. 3.6 Molecular Characterization of Fungi 3.6.1 Genomic DNA Extraction
Extraction of genomic DNA from selected fungal isolates can be done by using the methods of Samarrai et al. [15]. 1. Grind the 30 mg of fungal mycelium with liquid nitrogen in a prechilled mortar pestle and transfer the fine powder into the Eppendorf tube. 2. Resuspend the powdered fungal mycelium into 500 μL of the lysis buffer (Tris-acetate, 40 mmol/L; Sodium acetate, 20 mmol/L; EDTA, 1 mmol/L; SDS, 1% w/v; pH 7·8)
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[16, 17] by pipetting with 1000 μL pipette till the suspension viscosity is reduced significantly and the froth formation is the indication of a detachment of DNA from polysaccharide. 3. Repeat the pipetting process at least five times. 4. Add 2 μL of RNaseA (10 mg/mL); then incubate the suspension at 37 C for 5 min. 5. Add 165 μL of NaCl solution (5 mol/L) and mix by gently inverting the tube several times (>50 times), which allows the precipitation of protein, polysaccharides, and cell debris. 6. Precipitate the unwanted material by centrifuging (13,000 rpm) the suspension at 4 C for 20 min. 7. After centrifugation, immediately transfer the supernatant into a fresh Eppendorf tube and add phenol (400 μL) and chloroform (400 μL) in the ratio of 1:1. For proper mixing of the solution, gently invert the tube (50 times) till the whole solution becomes milky. 8. Centrifuge the solution at 12,000 rpm for 20 min, separate the supernatant or aqueous phase. 9. To precipitate DNA from the aqueous phase, add twice volumes of ethanol (95%). 10. Add the 500 μL of lysis buffer to the precipitated DNA and mix by gentle pipetting (by using cut tips) to make DNA free from the contaminants (polysaccharides). 11. Add 165 μL of NaCl (5 mol/L) and mix the suspension by inverting the tube gently several times. 12. Centrifuge (12,000 rpm) the solution for 10 min, discard the aqueous phase. Further, add 95% of ethanol for the precipitation of DNA. 13. If the aqueous phase still looks hazy, then re-extract it with one volume of chloroform before the DNA precipitation. 14. Wash the DNA three times with 70% prechilled ethanol and air-dry at room temperature. After that, dissolve the DNA in 50 mL of TE buffer (Tris–HCl, 10 mmol/L; EDTA, 0·1 mmol/L; pH 7·8) and store at -20 C till the further use. 15. Measure the concentration of genomic DNA by UV/VIS spectrophotometer (Perkin Elmer), using TE buffer as blank. Take the optical density of extracted DNA at 260 nm, which is used to calculate DNA concentration using the following equation: DNA Conc:ðμg=μLÞ ¼
O:D260 50 Dilution factor 1000
To check the purity of extracted DNA, take the optical density of DNA at a ratio of OD260/OD280 with the help of
Fungal Mediated Biodegradation of Pesticide
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a UV/VIS spectrometer. A value of 1.8 is optimum for best DNA preparation and if the value comes below 1.8, it signifies the contamination of proteins in the preparation, whereas if the value comes above 1.8, it indicates the preparation is contaminated with RNA. 16. Further, the purity can also be checked by electrophoresis. Load the extracted DNA on agarose gel (0.8% W/V) (with ethidium bromide 0.5 μg/mL) in 1X TAE buffer (pH 8.0) at 80 V (5 V/cm being optimal) for 40 min. Finally, for the visualization of the DNA band, observe the gel under UV light using a gel documentation system [16, 17]. 3.6.2 Polymerase Chain Reaction (PCR)
First, prepare the master mix. Perform the preparation of PCR reaction according to the final volume remains 50 μL. In addition to the template DNA (4 μL), it contains 5 μL of 10 PCR buffer, 1.5 μL of 10 mmol/L dNTP, 2.5 μL of 10 mmol/L stock solution of each primer (Table 1) (ITS-1 forward primer and NL-4 reverse primer), and 0.5 U of DNA polymerase. Perform the PCR reactions in a thermocycler machine and run with a temperature profile of 2 min at 94 C followed by 35 cycles of 15 s at 94 C, 30 s at 60 C and 1 min at 72 C. After 35 cycles, perform the last cycle for 5 min at 72 C. Perform each PCR reaction with two controls, a positive control (with DNA) and negative control (No DNA). After the completion of the reaction, a 5 μL aliquot of each PCR product runs on a 1% agarose gel and visualized by ethidium bromide staining to confirm amplification. A PCR reaction is considered to be successful if an amplicon generated from the template preparation, or if an additional manipulation of the template, such as second centrifugation or dilution, yielded a product. Gel results can be documented with a Gel Doc imaging system. Purified positive PCR products further are used for sequencing [18]. ITS sequences of test fungal isolates can be analyzed for homology with known 18S ITS sequences available at NCBI (National Centre for Biotechnology Information) database using BLAST (Basic Local Alignment Search Tool) [19]. Align the sequences by multiple sequence alignment using the Clustal W algorithm program. To deduce the evolutionary relatedness of the aligned sequences, construct the phylogram using the neighbor-joining method [20] with MEGA 4.0 (molecular evolutionary genetic analysis) software [21]. The neighbor-joining method generates a phylogenetic tree on the basis of the distance matrix, calculated from sequence data. The percent identity, divergence, and conserved sequence analysis can be done by MEGA alignment. Further, carry out the statistical evolution of tree topologies by bootstrapping [22]. Calculate the bootstrap confidence values from 1000 repeats.
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Table 1 List of primers used for the amplification of fungal ITS (nuclear ribosomal internal transcribed spacers 1, 2, and 5.8S) region Primer forward
Primer reveres
0
ITS1-5 -TCCGTAGGTGAACCTGCGG-3
0
0
NL4-5 -GGTCCGTGTTTCAAGACGG-3 0
Reference 0
[18] 0
ITS150 CTTGGTCATTTAGAGGAAGTAA-30
ITS1-5 -TCCTCCGCTTATTGATATGC-3
[18]
ITS8650 -GTGAATCATCGAATCTTTGAA-30
ITS8650 -TTCAAAGATTCGATGATTCAG-30
ITS3-50 -GCATCGATGAAGAACGCAGC30
ITS4-50 -TCCTCCGCTTATTGATATGC-30 [18]
[18]
ITS1-50 -TCCGTAGGTGAACCTGCGG-30 ITS4-50 -TCCTCCGCTTATTGATATGC-30 [16, 17]
4
Notes 1. Use sterilized gloves during the collection of samples. Preused or unsterilized gloves can be contaminated with the native microbial population. 2. Do not use old stock solutions of the chemicals, because in old stock solutions, the ionic balance becomes disturbed and it can give false results. 3. Maintain the appropriate pH of the fungal growth medium. High pH inhibits fungal growth. 4. Micropipette should be calibrated for the measurement of small volumes; otherwise, the results can be failed or give false results. 5. Wear gloves while handling EtBr and dispose of EtBr-stained agarose gel properly, because it is a mutagenic agent that can cause serious diseases such as cancer. 6. A PCR reaction that did not generate an amplicon is considered a failed attempt after three tries that utilized new template preparations. 7. While handling the fungal culture, always wear the face mask. Exposure to fungal spores can give an allergic response. 8. While making the serial dilution of the fungal sample, always use new micropipette tips for each dilution. 9. Switch on UV light of laminar airflow before 20 min of the use for complete sterilization. 10. While extracting the DNA from fungal cells, use an actively growing culture of fungi. From old culture, the yield of DNA can be reduced.
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References 1. Smidt H, de Vos WM (2004) Anaerobic microbial dehalogenation. Annu Rev Microbiol 58: 43–73 2. Gangola S, Sharma A, Bhatt P, Khati P, Chaudhary P (2018) Presence of esterase and laccase in Bacillus subtilis facilitates biodegradation and detoxification of cypermethrin. Sci Rep 8(1):1–11 3. Schuster E, Schro¨der D (1990) Side-effects of sequentially-applied pesticides on non-target soil microorganisms: field experiments. Soil Biol Biochem 22(3):367–373 4. Johnsen K, Jacobsen CS, Torsvik V, Sørensen J (2001) Pesticide effects on bacterial diversity in agricultural soils–a review. Biol Ferti Soils 33(6):443–453 5. Ahemad M, Khan MS, Zaidi A, Wani PA (2009) Remediation of herbicides contaminated soil using microbes. In: Microbes in sustainable agriculture. Nova Science Publishers, New York, pp 261–284 6. El Fantroussi S, Agathos SN (2005) Is bioaugmentation a feasible strategy for pollutant removal and site remediation? Curr Opin Microbiol 8(3):268–275 7. Singh H (2006) Mycoremediation: fungal bioremediation. Wiley 8. Read ND (2007) Environmental sensing and the filamentous fungal lifestyle. Fungi in the environment. Cambridge University Press, Cambridge, pp 38–57 9. Lynne B, Jones TH (2007) Mycelial responses in heterogeneous environments: parallels with macroorganisms. Fungi Environ 25:112–158 10. Gadd GM (2007) Geomycology: biogeochemical transformations of rocks, minerals, metals and radionuclides by fungi, bioweathering and bioremediation. Mycol Res 111(1):3–49 11. Harms H, Schlosser D, Wick LY (2011) Untapped potential: exploiting fungi in bioremediation of hazardous chemicals. Nat Rev Microbiol 9(3):177–192
12. Lindahl BD, Olsson S (2004) Fungal translocation–creating and responding to environmental heterogeneity. Mycologist 18(2):79–88 13. Siddique T, Okeke BC, Arshad M, Frankenberger WT (2003) Enrichment and isolation of endosulfan-degrading microorganisms. J Environ Qual 32(1):47–54 14. Hussain S, Arshad M, Saleem M, Zahir ZA (2007) Screening of soil fungi for in vitro degradation of endosulfan. World J Microbiol Biotechnol 23(7):939–945 15. Al-Samarrai TH, Schmid J (2000) A simple method for extraction of fungal genomic DNA. Lett Appl Microbiol 30(1):53–56 16. Gangola S, Khati P, Sharma A (2015) Mycoremediation of imidacloprid in the presence of different soil amendments using Trichodermalongibrachiatum and Aspergillusoryzae isolated from pesticide-contaminated agricultural fields of Uttarakhand. J Bioremed Biodegr 6(5):1 17. Gangola S, Pankaj GN, Srivastava A, Sharma A (2015) Enhanced biodegradation of endosulfan by Aspergillus and Trichoderma spp. isolated from an agricultural field of Tarai Region of Uttarakhand. Pestic Res J 27(2):223–230 18. Romanelli AM, Fu J, Herrera ML, Wickes BL (2014) A universal DNA extraction and PCR amplification method for fungal rDNA sequence-based identification. Mycoses 57(10):612–622 19. Altschul SF, Gish W, Miller W, Myers EW, Lipman DJ (1990) Basic local alignment search tool. J Mol Biol 215(3):403–410 20. Saitou N, Nei M (1987) The neighbor-joining method: a new method for reconstructing phylogenetic trees. Mol Biol Evol 4(4):406–425 21. Tamura K, Dudley J, Nei M, Kumar S (2007) MEGA4: molecular evolutionary genetics analysis (MEGA) software version 4.0. Mol Biol Evol 24(8):1596–1599 22. Felsenstein J (1985) Confidence limits on phylogenies: an approach using the bootstrap. Evolution 39(4):783–791
Chapter 2 Protocol for the Assessment of Mycoremediation of Polycyclic Aromatic Hydrocarbons Carlos Garcı´a-Delgado, Raquel Camacho-Are´valo, Begon˜a Mayans, Rafael Anto´n-Herrero, and Enrique Eymar Abstract Mycoremediation of soils polluted with polycyclic aromatic hydrocarbons (PAHs) is an effective approach in environmental biotechnology to restore ecosystems. In this protocol, we describe selected experimental procedures to assess the mycoremediation process of PAH-polluted soils using chemical and ecotoxicological methodologies. An analytical procedure is described to quantify the removal of PAHs. Additionally, a battery of ecotoxicological assays focused on soil microbial activity, survival and reproduction of invertebrates, and seed germination is also described. Key words Organic pollutants, Soil remediation, Ecotoxicity, Soil health
1
Introduction Comprehensive assessment of soil ecotoxicity is a difficult task due to the intrinsic complexity of soil pollution. The behavior of pollutants in the soil system is complex due to multiple factors that are implicated in the mobility and bioavailability of them [1]. The nature of pollutants and their physicochemical properties limit or favor the interaction with a solid matrix of soil [1, 2], which also impact the bioavailability of pollutants and their toxic effects. For example, pollutants with low water solubility (or high hydrophobicity) tend to be retained in soils and potentially produce lower ecotoxicity and vice versa [3]. In the same way, soil characteristics such as texture, pH, or organic carbon are the key factors to govern the mobility and bioavailability of pollutants [2, 3]. In general, soils with high clay and organic carbon content tend to retain pollutants, decreasing their mobility and bioavailability. However, all of the above factors that reduce the ecotoxicity, bioavailability, and mobility of pollutants limit the effectiveness of bioremediation technologies. This fact is clear in hydrophobic
Dhanushka Udayanga et al. (eds.), Mycoremediation Protocols, Springer Protocols Handbooks, https://doi.org/10.1007/978-1-0716-2006-9_2, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022
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pollutants such as polycyclic aromatic hydrocarbons (PAHs) that tend to be strongly adsorbed in soil [4]. PAHs are ubiquitous hazardous pollutants and highly resistant to natural degradation. PAHs are commonly found in sites associated with petroleum, gas production, and wood processing industries [1]. PAHs represent a broad group of different individual molecules made of two or more unsubstituted benzene rings fused. The PAHs persistence in the environment increases with increasing molecular weight [5], because of high molecular weight, lower bioavailability, among other factors. In addition, some PAHs present toxic, carcinogenic, and mutagenic properties [6]. They were, perhaps, the first recognized environmental carcinogens [5]. Mycoremediation is a promising approach in biotechnology to restore PAH-polluted soils, because ligninolytic fungi are able to biodegrade low bioavailable PAHs [5, 7, 8]. This group of fungi produces extracellular enzymes such as laccases, manganeseperoxidases, other versatile peroxidases, and lignin peroxidases, with low substrate specificity, which enable them to degrade a wide range of organic pollutants, including PAHs [9]. These extracellular enzymes are able to diffuse into the soil matrix and potentially oxidize PAHs with low bioavailability. Multiple studies are available that report the effectiveness of ligninolytic fungi to biodegrade PAHs and remediate PAH-polluted soils [9–15]. These fungi oxidize PAHs to produce oxygenated derivatives such as quinones, hydroxyl- and dihydroxyPAH, between other metabolites [5, 16]. Generally, these oxygenated PAHs are more polar, water soluble, and bioavailable than the parent compound. Therefore, such properties facilitate their further biodegradation by the soil microbiota. However, several oxygenated PAHs metabolites can be accumulated in the environment producing a negative impact in soil as well as human health due to their toxic and carcinogenic properties [4, 17, 18]. For this reason, only the determination of PAHs reduction after a mycoremediation process is not enough to assess the soil remediation process [19]. Hence, it is recommended to conduct a proper chemical analysis in combination with ecotoxicological assessment of mycoremediated soil by bioassays with different soil organisms (microorganisms, invertebrates, and plants) for a comprehensive assessment of the PAHs mycoremediation results. However, a large number of published articles are available about the mycoremediation of PAH-polluted soils or other related applications in biotechnology, which did not include ecotoxicological assays. Instead, many studies only determine the success of bioremediation by the reduction of the soil PAHs content. In contrast, some other related studies provide a comprehensive assessment of the bioremediation of PAH-polluted soil using ecotoxicological tests. Several ecotoxicological tests have been used to determine the recovery level of soil by mycoremediation such as
Assessment of the Mycoremediation of Polycyclic Aromatic Hydrocarbons
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acute and reproduction toxicity test [9, 13, 20, 21], soil enzymatic activity [9, 11, 12, 22], germination test [10, 12, 13, 15, 19, 21, 23, 24], genotoxicity test [14], or inhibition of bioluminescence test [19]. Thus, this chapter describes several methodologies to assess the results of mycoremediation of PAH-polluted soils at chemical and ecotoxicological levels with an analytical methodology for PAHs extraction and quantification. In addition, a battery of bioassays is provided to determine the recovery of soil functions. The ecotoxicological tests are based on three soil enzymatic activities that reflect the soil microbial activity, and the two bioassays are provided to determine the habitability of soil.
2
Materials
2.1 PAHs Extraction and Quantification by HPLC-PDA
2.2 Soil Enzymatic Activities 2.2.1 Dehydrogenase Activity (DHA)
2.2.2 Total Microbial Activity (TMA)
l
Pure quartz sand.
l
Cellulose filters.
l
Diatomaceous earth.
l
Dichloromethane HPLC grade.
l
Acetone HPLC grade.
l
Acetonitrile HPLC grade.
l
Ultrapure water (18 MΩ cm1 at 25 C).
l
16 US EPA PAHs standard mixture (Naph), acenaphthene (Ace), acenaphthylene (Acy), fluorene (Flu), phenanthrene (Phe), anthracene (Ant), fluoranthene (Fla), pyrene (Pyr), benzo[a]anthracene (BaA), chrysene (Chr), benzo[b]fluoranthene (BbF), benzo[k]fluoranthene (BkF), benzo[a]pyrene (BaP), dibenzo[a,h]anthracene (DBahA), benzo[ghi]perylene (BghiP), and indeno[1,2,3-cd]pyrene (IcdP).
l
M tris–HCl buffer at pH 7.5.
l
Solution of 1.5% of 2,3,5-tripheniltetrazolium chloride dissolved in 0.1 M tris–HCl buffer at pH 7.5.
l
Acetone analytical grade.
l
1,3,5-triphenylformazan.
l
Fluorescein diacetate 1 mg mL1 in acetone.
l
Sodium fluorescein.
l
Potassium phosphate buffer 60 mM pH 7.6.
l
Acetone analytical grade.
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2.2.3 Urease Activity (UA)
l
Borate buffer 0.1 M pH 8.8.
l
Urea 0.1 M.
l
KCl 1.35 M in HCl 0.1 M.
l
l
2.3 Germinability Test
2.4 Test of Survival and Reproduction of Collembolan (Folsomia candida)
Buffer of analysis: 50 g L1 of sodium potassium tartrate tetrahydrated, 14 g L1 of K2HPO4, 24 g L1 of NaOH. Salicylate reactive: 150 g L1 of sodium salicylate and 0.30 g L1 of sodium nitroprusiate.
l
Sodium hypochlorite 5.25%.
l
Seeds of plant species sensitive to PAHs such as Lactuca sativa, Brassica alba, Brassica chinensis, Lepidium sativum, and Trifolium pretense [12, 13, 25, 26].
l
Petri dishes Ø14 cm.
l
Deionized water.
l
Fine sand (