Trichoderma reesei: Methods and Protocols [1st ed.] 9781071610473, 9781071610480

Table of contents :
Front Matter ....Pages i-xi
Ecological Genomics and Evolution of Trichoderma reesei (Komal Chenthamara, Irina S. Druzhinina, Mohammad J. Rahimi, Marica Grujic, Feng Cai)....Pages 1-21
Industrial Relevance of Trichoderma reesei as an Enzyme Producer (Amanda J. Fischer, Suchindra Maiyuran, Debbie S. Yaver)....Pages 23-43
The Potential of Synthetic Biology for Trichoderma reesei (Roland Martzy, Astrid R. Mach-Aigner)....Pages 45-54
Resistance Marker- and Gene Gun-Mediated Transformation of Trichoderma reesei (Monika Schmoll, Susanne Zeilinger)....Pages 55-62
Use of Auxotrophic Markers for Targeted Gene Insertions in Trichoderma reesei (Irene Tomico-Cuenca, Christian Derntl)....Pages 63-72
Open the Pores: Electroporation for the Transformation of Trichoderma reesei (Franziska Wanka)....Pages 73-78
Sexual Crossing of Trichoderma reesei (Rita B. Linke)....Pages 79-85
CRISPR/Cas9-Mediated Genome Editing of Trichoderma reesei (Gen Zou, Zhihua Zhou)....Pages 87-98
The Copper-Controlled RNA Interference System in Trichoderma reesei (Lei Wang, Weixin Zhang, Xiangfeng Meng, Weifeng Liu)....Pages 99-111
Batch Cultivation of Trichoderma reesei (Birgit Jovanović)....Pages 113-118
Image Analysis Method for the Characterization of Trichoderma reesei During Fermentations (Nicolas Hardy, Maxime Moreaud, Fadhel Ben Chaabane)....Pages 119-133
Measurement of Cellulase and Xylanase Activities in Trichoderma reesei (Qing-Shan Meng, Fei Zhang, Chen-Guang Liu, Feng-Wu Bai, Xin-Qing Zhao)....Pages 135-146
Flow Cytometry for Filamentous Fungi (Matthias G. Steiger)....Pages 147-155
Molecular Identification of Trichoderma reesei (Mohammad J. Rahimi, Feng Cai, Marica Grujic, Komal Chenthamara, Irina S. Druzhinina)....Pages 157-175
In Vivo Footprinting Analysis in Trichoderma reesei (Alice Rassinger)....Pages 177-189
RNA Characterization in Trichoderma reesei (Petra Till)....Pages 191-235
Proteomic Profiling of the Secretome of Trichoderma reesei (So Fong Cam Ngan, Siu Kwan Sze)....Pages 237-249
Transcriptomics in Trichoderma reesei (Amanda C. C. Antonieto, Roberto N. Silva)....Pages 251-269
The Comprehensive and Reliable Detection of Secondary Metabolites in Trichoderma reesei: A Tool for the Discovery of Novel Substances (Bernhard Seidl, Christoph Bueschl, Rainer Schuhmacher)....Pages 271-295
In Silico Gene Analysis and Oligonucleotide Design for the Construction of Expression Vectors (Bernhard Seiboth)....Pages 297-309
PacBio Long-Read Sequencing, Assembly, and Funannotate Reannotation of the Complete Genome of Trichoderma reesei QM6a (Wan-Chen Li, Ting-Fang Wang)....Pages 311-329
TSETA: A Third-Generation Sequencing-Based Computational Tool for Mapping and Visualization of SNPs, Meiotic Recombination Products, and RIP Mutations (Hou-Cheng Liu, Wan-Chen Li, Ting-Fang Wang)....Pages 331-361
Back Matter ....Pages 363-364

Citation preview

Methods in Molecular Biology 2234

Astrid R. Mach-Aigner Roland Martzy Editors

Trichoderma reesei Methods and Protocols

METHODS

IN

MOLECULAR BIOLOGY

Series Editor John M. Walker School of Life and Medical Sciences University of Hertfordshire Hatfield, Hertfordshire, UK

For further volumes: http://www.springer.com/series/7651

For over 35 years, biological scientists have come to rely on the research protocols and methodologies in the critically acclaimed Methods in Molecular Biology series. The series was the first to introduce the step-by-step protocols approach that has become the standard in all biomedical protocol publishing. Each protocol is provided in readily-reproducible step-bystep fashion, opening with an introductory overview, a list of the materials and reagents needed to complete the experiment, and followed by a detailed procedure that is supported with a helpful notes section offering tips and tricks of the trade as well as troubleshooting advice. These hallmark features were introduced by series editor Dr. John Walker and constitute the key ingredient in each and every volume of the Methods in Molecular Biology series. Tested and trusted, comprehensive and reliable, all protocols from the series are indexed in PubMed.

Trichoderma reesei Methods and Protocols

Edited by

Astrid R. Mach-Aigner Institute of Chemical, Environmental and Bioscience Engineering, TU Wien, Vienna, Austria

Roland Martzy Christian Doppler Laboratory for Optimized Expression of Carbohydrate-Active Enzymes, Institute of Chemical, Environmental and Bioscience Engineering, TU Wien, Vienna, Austria

Editors Astrid R. Mach-Aigner Institute of Chemical, Environmental and Bioscience Engineering, TU Wien Vienna, Austria

Roland Martzy Christian Doppler Laboratory for Optimized Expression of Carbohydrate-Active Enzymes Institute of Chemical, Environmental and Bioscience Engineering, TU Wien Vienna, Austria

ISSN 1064-3745 ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-0716-1047-3 ISBN 978-1-0716-1048-0 (eBook) https://doi.org/10.1007/978-1-0716-1048-0 © Springer Science+Business Media, LLC, part of Springer Nature 2021 This work is subject to copyright. All rights are reserved 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. Cover Illustration Caption: Untitled painting by the Danish/American artist Maria Dubin, commissioned by Novozymes in 2017. The painting depicts Trichoderma reesei filaments growing and forming conidia on a tent. This represents the first identification of this mold during World War II, in the Solomon Islands. T. reesei wreaked havoc by growing on and degrading tents and other cotton-based textiles used by the US Army. T. reesei was recognized for its potential and was further developed as a solution to the oil crisis of the 1970s. 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 This volume of the Methods in Molecular Biology book series covers both standard techniques and cutting-edge methods that are frequently used by the Trichoderma reesei research community. We hope to address a wide range of audiences, from students who want to familiarize themselves with basic research protocols to experienced scientists who are planning to establish a new method in their laboratories. To be able to realize this project, we are happy to have succeeded in bringing on board many of the leading experts in the field, both from the academic and the industrial sectors. The first chapters are intended to have an introductory character and will give the reader a broad overview on the evolution of the fungus and its use as a host for the large-scale production of industrially relevant enzymes. In addition, examples on the possible applications of T. reesei in the popular field of synthetic biology are outlined. The following methodological chapters are structured in a process-oriented way, starting with transformation techniques and gene editing methods up to protocols dealing with the fermentation of the fungus. The second part of the book addresses downstream-analytical applications, such as enzyme activity testing and flow cytometry, as well as general analytical techniques for the molecular identification of T. reesei. This part also includes chapters covering the characterization of macromolecules, such as (long noncoding) RNAs and DNA-protein interactions. The final chapters address the area of -omics analyses and the corresponding bioinformatics approaches, which have become increasingly important during recent years. At this point, we would like to thank all the authors for their willingness to contribute and to share their expertise and their experience in the respective fields, as well as for their patience and cooperation throughout the editorial process. We hope that this book will serve not only as a collection of methods and protocols but also as a synopsis of researchers around the world who are working with T. reesei and the concomitant opportunity to establish contacts with the corresponding people. Wien, Austria

Astrid R. Mach-Aigner Roland Martzy

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Contents Preface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Contributors. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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1 Ecological Genomics and Evolution of Trichoderma reesei. . . . . . . . . . . . . . . . . . . . Komal Chenthamara, Irina S. Druzhinina, Mohammad J. Rahimi, Marica Grujic, and Feng Cai 2 Industrial Relevance of Trichoderma reesei as an Enzyme Producer . . . . . . . . . . . . Amanda J. Fischer, Suchindra Maiyuran, and Debbie S. Yaver 3 The Potential of Synthetic Biology for Trichoderma reesei . . . . . . . . . . . . . . . . . . . . Roland Martzy and Astrid R. Mach-Aigner 4 Resistance Marker- and Gene Gun-Mediated Transformation of Trichoderma reesei . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Monika Schmoll and Susanne Zeilinger 5 Use of Auxotrophic Markers for Targeted Gene Insertions in Trichoderma reesei . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Irene Tomico-Cuenca and Christian Derntl 6 Open the Pores: Electroporation for the Transformation of Trichoderma reesei . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Franziska Wanka 7 Sexual Crossing of Trichoderma reesei. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Rita B. Linke 8 CRISPR/Cas9-Mediated Genome Editing of Trichoderma reesei . . . . . . . . . . . . . Gen Zou and Zhihua Zhou 9 The Copper-Controlled RNA Interference System in Trichoderma reesei . . . . . . . Lei Wang, Weixin Zhang, Xiangfeng Meng, and Weifeng Liu 10 Batch Cultivation of Trichoderma reesei . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Birgit Jovanovic´ 11 Image Analysis Method for the Characterization of Trichoderma reesei During Fermentations. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Nicolas Hardy, Maxime Moreaud, and Fadhel Ben Chaabane 12 Measurement of Cellulase and Xylanase Activities in Trichoderma reesei. . . . . . . . Qing-Shan Meng, Fei Zhang, Chen-Guang Liu, Feng-Wu Bai, and Xin-Qing Zhao 13 Flow Cytometry for Filamentous Fungi . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Matthias G. Steiger 14 Molecular Identification of Trichoderma reesei . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Mohammad J. Rahimi, Feng Cai, Marica Grujic, Komal Chenthamara, and Irina S. Druzhinina 15 In Vivo Footprinting Analysis in Trichoderma reesei . . . . . . . . . . . . . . . . . . . . . . . . . Alice Rassinger

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RNA Characterization in Trichoderma reesei. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Petra Till Proteomic Profiling of the Secretome of Trichoderma reesei . . . . . . . . . . . . . . . . . . So Fong Cam Ngan and Siu Kwan Sze Transcriptomics in Trichoderma reesei . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Amanda C. C. Antonieto and Roberto N. Silva The Comprehensive and Reliable Detection of Secondary Metabolites in Trichoderma reesei: A Tool for the Discovery of Novel Substances . . . . . . . . . . Bernhard Seidl, Christoph Bueschl, and Rainer Schuhmacher In Silico Gene Analysis and Oligonucleotide Design for the Construction of Expression Vectors. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Bernhard Seiboth PacBio Long-Read Sequencing, Assembly, and Funannotate Reannotation of the Complete Genome of Trichoderma reesei QM6a . . . . . . . . . . . . . . . . . . . . . . Wan-Chen Li and Ting-Fang Wang TSETA: A Third-Generation Sequencing-Based Computational Tool for Mapping and Visualization of SNPs, Meiotic Recombination Products, and RIP Mutations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Hou-Cheng Liu, Wan-Chen Li, and Ting-Fang Wang

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

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Contributors AMANDA C. C. ANTONIETO • Department of Biochemistry and Immunology, Ribeira˜o Preto Medical School, University of Sa˜o Paulo, Ribeira˜o Preto, SP, Brazil FENG-WU BAI • State Key Laboratory of Microbial Metabolism, Joint International Research Laboratory of Metabolic and Developmental Sciences, School of Life Sciences and Biotechnology, Shanghai Jiao Tong University, Shanghai, China FADHEL BEN CHAABANE • IFP Energies nouvelles, Rueil-Malmaison, France CHRISTOPH BUESCHL • Department of Agrobiotechnology (IFA-Tulln), Institute of Bioanalytics and Agro-Metabolomics, University of Natural Resources and Life Sciences, Vienna (BOKU), Tulln, Austria FENG CAI • Fungal Genomics Laboratory (FungiG), The Key Laboratory of Plant Immunity, Nanjing Agricultural University, Nanjing, China; Institute of Chemical, Environmental and Bioscience Engineering (ICEBE), TU Wien, Vienna, Austria KOMAL CHENTHAMARA • Fungal Genomics Laboratory (FungiG), The Key Laboratory of Plant Immunity, Nanjing Agricultural University, Nanjing, China; Institute of Chemical, Environmental and Bioscience Engineering (ICEBE), TU Wien, Vienna, Austria CHRISTIAN DERNTL • Institute of Chemical, Environmental and Bioscience Engineering, TU Wien, Vienna, Austria IRINA S. DRUZHININA • Fungal Genomics Laboratory (FungiG), The Key Laboratory of Plant Immunity, Nanjing Agricultural University, Nanjing, China; Institute of Chemical, Environmental and Bioscience Engineering (ICEBE), TU Wien, Vienna, Austria AMANDA J. FISCHER • Novozymes, Inc., Davis, CA, USA MARICA GRUJIC • Institute of Chemical, Environmental and Bioscience Engineering (ICEBE), TU Wien, Vienna, Austria NICOLAS HARDY • IFP Energies nouvelles, Rueil-Malmaison, France; DuPont Nutrition & Biosciences, Z.A. des Buxie`res, Dange´-Saint-Romain, France BIRGIT JOVANOVIC´ • Institute of Chemical, Environmental and Bioscience Engineering, TU Wien, Vienna, Austria WAN-CHEN LI • Taiwan International Graduate Program in Molecular and Cellular Biology, Academia Sinica, Taipei, Taiwan, Republic of China; Institute of Life Sciences, National Defense Medical Center, Taipei, Taiwan, Republic of China; Institute of Molecular Biology, Academia Sinica, Taipei, Taiwan, Republic of China RITA B. LINKE • Institute of Chemical, Environmental and Bioscience Engineering, Research Group of Environmental Microbiology and Molecular Diagnostics, TU Wien, Vienna, Austria CHEN-GUANG LIU • State Key Laboratory of Microbial Metabolism, Joint International Research Laboratory of Metabolic and Developmental Sciences, School of Life Sciences and Biotechnology, Shanghai Jiao Tong University, Shanghai, China HOU-CHENG LIU • Institute of Molecular Biology, Academia Sinica, Taipei, Taiwan, Republic of China WEIFENG LIU • State Key Laboratory of Microbial Technology, Shandong University, Qingdao, People’s Republic of China

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ASTRID R. MACH-AIGNER • Institute of Chemical, Environmental and Bioscience Engineering, TU Wien, Vienna, Austria SUCHINDRA MAIYURAN • Novozymes, Inc., Davis, CA, USA ROLAND MARTZY • Christian Doppler Laboratory for Optimized Expression of CarbohydrateActive Enzymes, Institute of Chemical, Environmental and Bioscience Engineering, TU Wien, Vienna, Austria QING-SHAN MENG • State Key Laboratory of Microbial Metabolism, Joint International Research Laboratory of Metabolic and Developmental Sciences, School of Life Sciences and Biotechnology, Shanghai Jiao Tong University, Shanghai, China XIANGFENG MENG • State Key Laboratory of Microbial Technology, Shandong University, Qingdao, People’s Republic of China MAXIME MOREAUD • IFP Energies nouvelles, Solaize, France SO FONG CAM NGAN • School of Biological Sciences, Nanyang Technological University, Singapore, Singapore MOHAMMAD J. RAHIMI • Institute of Chemical, Environmental and Bioscience Engineering (ICEBE), TU Wien, Vienna, Austria; Fungal Genomics Laboratory (FungiG), The Key Laboratory of Plant Immunity, Nanjing Agricultural University, Nanjing, China ALICE RASSINGER • ACIB GmbH, c/o BOKU, Vienna, Austria MONIKA SCHMOLL • Center for Health and Bioresources, AIT Austrian Institute of Technology GmbH, Tulln, Austria RAINER SCHUHMACHER • Department of Agrobiotechnology (IFA-Tulln), Institute of Bioanalytics and Agro-Metabolomics, University of Natural Resources and Life Sciences, Vienna (BOKU), Tulln, Austria BERNHARD SEIBOTH • Research Division Biochemical Technology, Institute of Chemical, Environmental & Bioscience Engineering, TU Wien, Vienna, Austria BERNHARD SEIDL • Department of Agrobiotechnology (IFA-Tulln), Institute of Bioanalytics and Agro-Metabolomics, University of Natural Resources and Life Sciences, Vienna (BOKU), Tulln, Austria ROBERTO N. SILVA • Department of Biochemistry and Immunology, Ribeira˜o Preto Medical School, University of Sa˜o Paulo, Ribeira˜o Preto, SP, Brazil MATTHIAS G. STEIGER • Institute of Chemical, Environmental and Bioscience Engineering, TU Wien, Vienna, Austria; Austrian Centre of Industrial Biotechnology (ACIB), Vienna, Austria SIU KWAN SZE • School of Biological Sciences, Nanyang Technological University, Singapore, Singapore PETRA TILL • Christian Doppler Laboratory for Optimized Expression of CarbohydrateActive Enzymes, Institute of Chemical, Environmental and Bioscience Engineering, TU Wien, Vienna, Austria IRENE TOMICO-CUENCA • Institute of Chemical, Environmental and Bioscience Engineering, TU Wien, Wien, Austria LEI WANG • State Key Laboratory of Microbial Technology, Shandong University, Qingdao, People’s Republic of China TING-FANG WANG • Institute of Molecular Biology, Academia Sinica, Taipei, Taiwan, Republic of China; Taiwan International Graduate Program in Molecular and Cellular Biology, Academia Sinica, Taipei, Taiwan, Republic of China FRANZISKA WANKA • Austrian Centre of Industrial Biotechnology (ACIB) GmbH c/o Research Division Biochemical Technology, Institute of Chemical, Environmental & Bioscience Engineering, TU Wien, Vienna, Austria

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DEBBIE S. YAVER • Novozymes, Inc., Davis, CA, USA SUSANNE ZEILINGER • Department of Microbiology, University of Innsbruck, Innsbruck, Austria FEI ZHANG • State Key Laboratory of Microbial Metabolism, Joint International Research Laboratory of Metabolic and Developmental Sciences, School of Life Sciences and Biotechnology, Shanghai Jiao Tong University, Shanghai, China WEIXIN ZHANG • State Key Laboratory of Microbial Technology, Shandong University, Qingdao, People’s Republic of China XIN-QING ZHAO • State Key Laboratory of Microbial Metabolism, Joint International Research Laboratory of Metabolic and Developmental Sciences, School of Life Sciences and Biotechnology, Shanghai Jiao Tong University, Shanghai, China ZHIHUA ZHOU • CAS-Key Laboratory of Synthetic Biology, CAS Center for Excellence in Molecular Plant Sciences, Institute of Plant Physiology and Ecology, Shanghai Institutes for Biological Sciences, Chinese Academy of Sciences, Shanghai, China GEN ZOU • CAS-Key Laboratory of Synthetic Biology, CAS Center for Excellence in Molecular Plant Sciences, Institute of Plant Physiology and Ecology, Shanghai Institutes for Biological Sciences, Chinese Academy of Sciences, Shanghai, China

Chapter 1 Ecological Genomics and Evolution of Trichoderma reesei Komal Chenthamara, Irina S. Druzhinina, Mohammad J. Rahimi, Marica Grujic, and Feng Cai Abstract The filamentous fungus Trichoderma reesei (Hypocreales, Ascomycota) is an efficient industrial cell factory for the production of cellulolytic enzymes used for biofuel and other applications. Therefore, researches addressing T. reesei are relatively advanced compared to other Trichoderma spp. because of the significant bulk of available knowledge, multiple genomic data, and gene manipulation techniques. However, the established role of T. reesei in industry has resulted in a frequently biased understanding of the biology of this fungus. Thus, the recent studies unexpectedly show that the superior cellulolytic activity of T. reesei and other Trichoderma species evolved due to multiple lateral gene transfer events, while the innate ability to parasitize other fungi (mycoparasitism) was maintained in the genus, including T. reesei. In this chapter, we will follow the concept of ecological genomics and describe the ecology, distribution, and evolution of T. reesei, as well as critically discuss several common misconceptions that originate from the success of this species in applied sciences and industry. Key words Ankyrins, Cellulolytic enzymes, Gene duplication, Gene loss, Lateral gene transfer, Mycoparasitism, Orphan genes, Phylogenomics, T. parareesei, Transcriptomics

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Introduction The genus of filamentous fungi Trichoderma (Hypocreales, Ascomycota) is best known for T. reesei—the industrial producer of cellulolytic and hemicellulolytic enzymes for biofuel and numerous other manufactured products. In suitable industrial fermentation conditions, genetically improved mutants of T. reesei can yield over 100 g of secreted protein per 1 liter of broth, making this fungus a primary choice for numerous commercial formulations [1]. Recently, advances in molecular biological and synthetic biological technologies have allowed the development of a T. reesei-based microbial cell factory for the production of heterologous proteins [2–6] and secondary metabolites [7]. However, the established role of T. reesei in the commercial production of cellulolytic enzymes has resulted in a frequently biased understanding of

Astrid R. Mach-Aigner and Roland Martzy (eds.), Trichoderma reesei: Methods and Protocols, Methods in Molecular Biology, vol. 2234, https://doi.org/10.1007/978-1-0716-1048-0_1, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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the biology of this fungus, where the valuable applied properties are extrapolated to the environmental adaptations. In this chapter, we will follow the concept of ecological genomics, which is an interdisciplinary area targeting the understanding of the gene and genome function in the natural environment [8]. We will also describe the ecology and evolution of this species and critically discuss several common misconceptions that originate from the success of T. reesei in applied sciences and industry.

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QM6a: Ever Since the Second World War T. reesei is unique among other model fungi because all of the strains used in industry have been derived by various genetic improvement techniques of a single wild-type isolate (for a review, see refs. 3, 5, 9). The discovery of the strongly cellulolytic Trichoderma strain dates back to the Second World War on Solomon Islands in Oceania, where it was isolated from rotting cotton fabric items of the US Army. The strain was first identified as T. viride and labeled QM6a by Mary Mandels and Elwyn T. Reese [10] in 1957. During the first oil crisis in the 1970s, Reese and Mandels, who were then both researchers at the US Army Quartermaster Research and Development Center at Natick, Massachusetts, initiated a study toward the commercial production of enzymes capable of hydrolyzing plant biomass. It was then that the QM6a strain came into light for its outstanding cellulase induction properties. They proposed the use of glucose from lignocellulose for bioethanol formation [11, 12]. Thus, QM6a became the parental strain of a pedigree of higher producing mutant strains. Consequently, all strains used in industry today have been derived from QM6a, which laid the foundation for genotype–phenotype studies based on the standardized genomic background. With the development of Trichoderma taxonomy, the species name of the QM6a isolate changed. Two decades after its initial naming, in the Second International Mycological Congress (1977), E.G. Simmons presented the distinction between QM6a strain and reference taxon for the genus T. viride [13]. He noticed that the strain’s morphology did not fit any of the then-described nine species of the genus known that time and proposed the species epithet “reesei” (in honor of Elwyn T. Reese) [14]. However, it was shortly discontinued because J. Bissett [15] attributed QM6a to T. longibrachiatum, the type species of the Longibrachiatum section within the genus Trichoderma. Consequently, several cellulolytic products still contain the “T. longibrachiatum” name in their annotations. However, the early molecular biological techniques— for example, the restriction fragment length polymorphism methods [16, 17] and combined morphometric and isozyme analyses [18, 19]—showed that T. reesei and T. longibrachiatum are

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taxonomically separable. Moreover, considerable morphological and isozyme differences between temperate and tropical collections of Hypocrea schweinitzii (now abandoned teleomorphic name for T. citrinoviride [20], vide infra) led to the recognition that the tropical collections were actually Hypocrea jecorina, the hypocrealean species described based on fruiting structures. Thus, the species name of T. reesei was recovered, and this taxon was proposed to be more closely related to Hypocrea jecorina than to either of other species of the section that were known by that time. Ten years later, the groups of Gary. J. Samuels (USDA, USA), Christian. P. Kubicek (TU Wien, Austria) and colleagues introduced molecular identification in Trichoderma taxonomy (which was then named DNA barcoding) and revealed the convincing molecular evolutionary evidence that QM6a was a clonal derivative of the holomorphic ascomycete Hypocrea jecorina. Since that time, the two names—the anamorphic species name T. reesei and the teleomorphic name H. jecorina—were applied to QM6a and other similar isolates. However, based on the }59 of the International Code of Botanical Nomenclature, which implied that for holomorphic fungi (those that reproduce sexually and asexually, such as T. reesei), the name of the teleomorph must be used as a single species name, the correct name for the organism was H. jecorina. It was followed in some publications but largely ignored in the area of applied science. Consequently, the existence of the two names for one organism was considered inconvenient. The situation with T. reesei and H. jecorina was not unique but illustrated the global trend in mycology when molecular methods allowed connections of numerous anamorph–teleomorph pairs and resulted in cases of two names per one fungal life cycle. To solve this taxonomic collision, in 2013 the International Commission on the Taxonomy of Fungi (ICTF, IUMS) agreed upon the Amsterdam Declaration on Fungal Nomenclature to use only a single name for a given fungus (i.e., either that of the ana- or teleomorph) and left it free which name the subcommittees for the individual genera would decide with a suggestion to give priority to the older name [21]. While this recommendation was criticized [22], results from a poll among researchers working with Trichoderma were in favor of using Trichoderma instead of Hypocrea (www.isth.info; [23]). Samuels [24] proposed the conservation of the well-known younger name, T. reesei, over H. jecorina. Thus, starting from January 1, 2013, the only correct species name for the QM6a strain and other co-specific isolates—irrespective of whether they have been isolated from sexual or asexual stages—is T. reesei [9, 20, 25]. The QM6a isolate that is also archived as IMI 192654, CBS 383.78, DSM 768, or ATCC 13631 in culture collections is the type strain of the species.

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The Origin of the First Trichoderma Superhero T. reesei is a member of the section Longibrachiatum of the genus Trichoderma [26, 27] that belongs to the family Hypocreaceae within the large order Hypocreales, which is a member of the class Sordariomycetes and the phylum Ascomycota. Hypocrealean fungi (those belonging to the order Hypocreales) share the common ancestor (monophyletic) approximately 200 Mya that most likely was associated with plants as either a pathogen or a mutualistic partner [28–30]. Extant members of this order are mainly biotrophs on plants (e.g., Fusarium spp.), insects (i.e., Cordyceps spp.), or fungi (i.e., Escovopsis spp.), and only some hypocrealean fungi are saprotrophs [31]. Phylogenetic and early phylogenomic studies revealed that the family Hypocreaceae combines such genera as Escovopsis and Hypomyces that were characterized by a strictly mycoparasitic lifestyle but also included a dozen of other genera (see NCBI Taxonomy browser). Sung et al. [28] proposed that shifts to fungicolous nutrition occurred several times during the evolution of hypocrealean fungi, since mycotrophs were present not only in Hypocreaceae but in Ophiocordycipitaceae and Bionectriaceae. Since the initial hosts of Hypocreaceae were supposed to be from the phylum Basidiomycota, this jump was best explained by the host-habitat hypothesis [32] (i.e., that new hosts were acquired due to their proximity in the environment), because arthropods including insects were often found on basidiocarps [32]. The attribution of the genus Trichoderma to the fungicolous family Hypocreaceae was confirmed by numerous phylogenetic and phylogenomic studies [29, 30, 33–35]. We analyzed more than 20 hypocrealean genomes [29, 30, 35] including a dozen of Trichoderma species and revealed that Trichoderma genus evolved from an ancestor with limited cellulolytic capabilities that likely fed on either fungi or arthropods. These analyses revealed that Trichoderma shared the last common ancestor with the genus Escovopsis and several entomopathogenic families such as Cordycipitaceae, Ophiocordycipitaceae, and Clavicipitaceae. The genus Trichoderma was formed in the time of the Cretaceous-Paleogene extinction event 66 (15) Mya, but the establishment of the Longibrachiatum section that contains T. reesei occurred in the Oligocene, 22 Mya [29, 30]. Thus, the fungicolous nutrition and mycoparasitism of Trichoderma spp. were described as an innate lifestyle of the genus [33, 36]. This conclusion was in agreement with a plethora of Trichoderma diversity and taxonomy studies performed over the last two decades which demonstrated that the fruiting bodies of other fungi or the dead wood colonized by other fungi were the most common habitats of Trichoderma spp. [36–38], while only a few (10–15%) Trichoderma spp. could establish in soil and rhizosphere [39, 40].

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In vitro, all Trichoderma species including T. reesei and related species could form abundant growth when fungal biomass was offered as the only source of nutrients (Fig. 1). However, this finding did not explain the superior cellulolytic activity of T. reesei QM6a that was shown not to be an exceptional property of a single strain or a species but reflected the overall high cellulolytic potential for the most common Trichoderma species such as T. longibrachiatum, T. citrinoviride, or T. harzianum (Fig. 1).

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An Exceptional Genome of T. reesei The understanding of the superior cellulolytic activity of T. reesei QM6a was the primary task for the initial genomic studies of Trichoderma spp. Owing to the potential mentioned above, T. reesei QM6a became the second hypocrealean fungus (after Fusarium graminearum PH-1 [41]) and the first Trichoderma whose genome was sequenced in 2006 and published in 2008 [42]. A total of 89 scaffolds were assembled, generating a genome with the size of 34 Mbp, and at that time, a total of 9129 genes were predicted in the genome. It was interesting to note that despite being the model organism for producing cellulolytic enzymes, the genome of T. reesei had fewer cellulases and hemicellulases encoding genes than the other sequenced plant cell wall degrading fungi like Magnaporthe grisea 70-15 (Magnaporthales, Ascomycota) [43] and F. graminearum PH-1 [41]. Shortly, after just 2 months, the genome of the genetically improved mutant of T. reesei QM6a, Rut-C30 strain, that had increased cellulolytic activity was published, revealing the genetic differences between it and the wild-type isolate [44]. Before this, only two differences were described between the two strains: a truncated carbon catabolite repressor protein CRE1 making Rut-C30 catabolite depressed and an additional frameshift mutation in the glycoprotein processing ß-glucosidase II gene. The Rut-C30 genome further revealed the deletion of a 0.085 Mbp fragment, including 29 open reading frames. This comparison also explained the differences between the phenotypes of the two strains. Later, genome-wide transcriptomic studies were also performed to reveal strategies employed by T. reesei to degrade lignocellulose in comparison with another model fungi, Aspergillus niger (Eurotiales, Ascomycota) [45]. In this study, all the genes induced in T. reesei when exposed to wheat straw were revealed using RNA-sequencing methods. It was shown that approximately 13% of the total mRNA was induced after 24 h period of exposure to wheat straw. The analysis revealed that enzymes from the same glycoside hydrolase families—mainly from GH11, GH7, GH3, GH30, and GH61 but different carbohydrate esterase (CE15 in T. reesei and CE8 and CE12 in A. niger)—were induced. Accessory proteins that had been shown to play a role in

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Fig. 1 Growth of T. reesei and related species on natural substrates. (a) Four Trichoderma spp. strains were cultivated for 8 days in darkness at 25  C on Petri plates (9 cm diameter) containing a solid medium plus one of the plant biomass-related substrates and agar or agarose. Images were taken using a Nikon D70 digital camera in ambient illumination. (b) Four Trichoderma spp. strains and a strain of fungicolous hypocrealean fungus Escovopsis weberi were cultivated on the agar medium containing fungal biomass prepared as described in [35]. The fungi were cultivated in 24-well plates for 8 days in darkness at 25  C. Images were taken using an Owl camera and a stereomicroscope. The scale bar corresponds to 5 mm

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enhancing carbohydrate deconstruction [46] were also revealed in the genome of T. reesei (also present in the genome of A. niger). Besides, the one-lipase encoding gene ceramidase (Transcript ID: 64397) was induced similarly as the GH- and CE-encoding genes in the presence of wheat straw, suggesting a role in plant cell wall degradation. Two hydrophobin encodings HFB2 (Transcript ID: 119989) and HFB3 (Transcript ID: 123967) and three cell wall proteins—that are, a QI74 orthologue (Transcript ID: 74282), a cell wall protein containing HsbA conserved domain (Transcript ID: 104277), and another one with a CFEM domain (Transcript ID: 124295)—all played a role in recognition of solid surfaces, which is an essential step in the fungal response to the plant cell wall. And they were also induced during a switch from glucose to the wheat straw. Induction of seven transporter genes belonging to the Major Facilitator Superfamily, one xylose transporter, two oligo-peptide transporters, and one iron transporter were highly induced in straw conditions, suggesting degradation of cellulose and hemicellulose fractions of wheat straw for producing simple sugars for the fungus. XYL1, a gene of the xylose utilization pathway, and XDH1, a gene of the xylitol dehydrogenase pathway, were also upregulated more than 20 folds when transitioning from glucose to straw, indicating the internalization of hemicellulosic sugars such as xylose by T. reesei. With the advancement in genome sequencing technologies, more and more mutants of T. reesei were sequenced. Subsequently, a comparison between the chromosome structures and sequences of eight cellulase mutant strains of T. reesei [3, 47–51] revealed more genes of regulatory relevance for cellulase induction. Also, the genome of one wild-type isolate—CBS 999.97 from a salt lake in French Guiana—was sequenced [52, 53], revealing the locus responsible for the female sterility of T. reesei [51]. Replacement of this locus (ham5) in the wild-type allele of T. reesei QM6a allowed sexual crossing with the parental strain QM6a [51]. Thus, today T. reesei is unique among the 16 Trichoderma spp. whose genome have so far been sequenced and annotated (https:// mycocosm.jgi.doe.gov/mycocosm/home, data retrieved September 29, 2019) because its seven chromosomes and their nt sequences are known [54–58], and its complete genome (34,922,528 bp; 10,877 genes) has been fully annotated [29, 55]. The breakthrough in the understanding of the cellulolytic activity of T. reesei and other Trichoderma species was achieved when the evolutionary analysis of all 122 individual genes encoding the plant cell wall degrading carbohydrate-active enzymes and auxiliary proteins (pcwdCAZymes) was performed [30, 35]. In that study, gene tree/species tree reconciliation methods were used to discover the impact of a massive (about 40%) lateral gene transfer (LGT) of such genes to Trichoderma genomes. Most surprisingly, it revealed that the donors of these laterally transferred

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pcwdCAZyme genes were fungi from different classes of Pezizomycotina fungi but none of the common Trichoderma hosts from the phylum Basidiomycota. This study showed that the majority of transfers occurred before the formation of the common ancestor of extant Trichoderma species—i.e. between 120 and 66 Mya [29, 30]—while several cases, including the transfer of the cbh1 (cel7a) gene that encodes the major industrially relevant cellulase CBH1, occurred before the divergence between the ancestor of modern Trichoderma and Escovopsis, i.e., more than 120 Mya [29, 30, 35]. It was a remarkable finding that explained the outstanding nutritional versatility of Trichoderma spp., which is only partially reflected in the ecophysiological profile of T. reesei (Fig. 1). Interestingly, the massive LGT was also restricted to pcwdCAZymes and only rarely observed in other gene families (K. Chenthamara, F. Cai, I. S. Druzhinina, unpublished). We believe that the recent evolutionary acquisition of the majority of pcwdCAZymes by Trichoderma spp., including T. reesei, partially explains the usability of these fungi in industry. Druzhinina et al. [30, 35] also showed that all known efficient cellulase regulatory proteins (XYR1, ACE2, and ACE3) [36] evolved vertically along with the evolution of the genus. Thus, LGT-derived pcwdCAZymes originating from different fungi were able to be controlled by a few innate regulatory proteins. We speculate that this resulted in a relatively simple and controllable regulation of T. reesei cellulases and hemicellulases compared to the regulatory network in the innate cellulolytic fungi such as Aspergillus spp. and Fusarium spp. Although the genomes of the later fungi encode at least two folds more cellulolytic enzymes compared to T. reesei, they were not selected as suitable cell factories for industrial production, likely because of a more evolutionary advanced and sophisticated regulatory system. Although most of LGT events were shown to occur before the formation of the last common ancestor of the genus Trichoderma, individual cases were also recorded in the more recent evolutionary history of species from the section Trichoderma (T. asperellum and T. atroviride), T. virens and species from the Harzianum clade (T. harzianum and T. guizhouense) [30, 35]. Interestingly, no LGT events were recorded to be specific for section Longibrachiatum. This correlates well with the overall small genome size of fungi from section Longibrachiatum that contains roughly 20% fewer genes compared to other species [29, 33]. The gene gain–gene loss analysis at the level of sections and individual species performed by Kubicek et al. [29] revealed that the significant loss of genes accompanied the formation of the section Longibrachiatum, which was the highest in the whole genus. However, individual species such as T. reesei and its sister species T. parareesei underwent a few single recent gene gain events since their divergence approximately 5 Mya. The gene gain estimated for these species was significantly

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lower compared to the species in other infrageneric groups, such as section Trichoderma, where individual species experienced up to 60 gene gain events, or the Virens clade, where T. virens reached almost one hundred [29]. Interestingly, the loss of genes was a unique feature of the evolution of the common ancestor of the section Longibrachiatum as it was not recorded for other groups: the gene losses were estimated to be an anticipated event only in the recent evolutionary history of all species except T. virens. The gene contraction in section Longibrachiatum took place in nearly all functional groups, which suggests that genetic drift (one driving force of genome contraction [59]) might not be the reason behind such a genome alteration. Instead, a more parsimonious lifestyle through conservation of energy for growth and development could be the driving force for genome reduction, since the genome of T. reesei does not lack genes for specific functions compared to other Trichoderma spp. [29] but contains only one or a few of those instead of several paralogs that are present in other species [35]. Sun and Blanchard [59] explained ecological advantages of having a smaller genome because these organisms would need to spend less energy for growth and development and thus may thrive relatively easier in a stable environment than their competitors with larger genomes. Another striking feature in the evolution of Trichoderma, including that of T. reesei, is the occurrence of a high number of orphan genes (i.e., genes that do not have homologs in species of adjacent clades). Orphan genes are theorized to be originating through gene duplication events, rearrangement processes and subsequent fast divergence or from de novo evolution out of non-coding genomic regions [60]. In the case of T. reesei, only a fifth of its orphan genes are shown to be occurring in clusters [29], which is a signal that points toward gene duplications. Only a tiny portion of orphans (clustered and non-clustered) are near the telomeres which are common area for gene duplication. Finally, these orphan genes are not preferred targets for repeat-induced point mutation (RIP) either, which inactivates duplicated genes. Therefore, the hypothesis of gene duplication as the principal mechanism for the emergence of orphan genes is not supported in this case. Published transcriptome data from T. reesei [61] showed that approximately 40% of the orphan genes are indeed expressed and therefore represent protogenes which are exposed to natural selection [62]. Chenthamara [30] verified this selection pressure in the case of orphan ankyrin genes present in the core genome by showing these genes evolved under purifying selection. The detection of the massive LGT of pcwdCAZymes to Trichoderma spp. from a narrow taxonomic group of other fungi (Pezizomycotina) strongly supported the long evolutionary history of Trichoderma mycoparasitism. It was widely demonstrated that all

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studied Trichoderma, including T. reesei QM6a, could parasitize distant species from Basidiomycota [63, 64] as well as their close neighbors, including the putative donors of acquired plant cell wall degrading genes [35]. This suggests Trichoderma’s ability to parasitize on taxonomically close neighbors (up to adelphoparasitism, parasitism on the members of the same family or the genus) might be the driving force behind the massive LGT.

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The Mycoparasitic Vigor of T. reesei: An Unpopular Fact Although an analysis involving the first three available genomes of Trichoderma (T. reesei, T. virens, and T. atroviride) suggested that mycoparasitism—that is the ability to parasitize on fungi—is an innate property of Trichoderma [33], only three Trichoderma species (i.e., T. atroviride, T. harzianum, and T. virens) have been extensively investigated for the characterization of genes involved in fungal–fungal interactions [65–76]. Chenthamara and Druzhinina [77] showed an comprehensive summary of genes studied over the past two decades involved in mycoparasitism, namely those Trichoderma genes that have the potential of influencing the pathogens of the plant. Most of these genes were involved in signal transduction, fungal cell wall degradation, and production of secondary antifungal metabolites. Mycoparasitism by Trichoderma is unique in the sense that they can parasitize even taxonomically close species (up to adelphoparasitism in the strict sense), unlike other mycoparasitism by species of genus Hypomyces spp. whose parasitism is restricted to Basidiomycota. Hence, this property of Trichoderma is primarily applied in the area of bioeffectors but also makes Trichoderma a devastating pest for mushroom farms [78–80]. Few studies exist presenting T. reesei as a bioeffector due to its overhype as an industrial cellulase producer. Atanasova et al. [63], through comparative transcriptomics studies, revealed interesting strategy in T. reesei interactions with other fungi as compared to T. atroviride and T. virens. Dual confrontation assays were set between Thanatephorus cucumeris (Rhizoctonia solani, Cantharellales, Basidiomycota) and the three Trichoderma species at 25  C. Transcriptional responses were observed in all three fungi even before contact between hyphae of two fungi on the assay plate. In the case of T. atroviride, an array of genes involved in the production of secondary metabolites, GH16 ß-glucanases, various proteases, and small secreted cysteine-rich proteins (SSCPs) were expressed. In T. virens, mainly the genes for biosynthesis of gliotoxin, respective precursors and also glutathione, which is necessary for gliotoxin biosynthesis, were expressed. In contrast, T. reesei increased the expression of genes encoding cellulases and hemicellulases and of the genes involved in solute transport. Thus, T. reesei

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Fig. 2 Mycoparasitism of GFP-labeled T. reesei TUCIM 4817 on Pestalotiopsis fici TUCIM 5788 observed on the surface of the glass slide, as described in [35]. The hyphal interaction was observed using a Nikon confocal laser scanning microscope (left) and a white light microscope (right). The scale bar corresponds to 10 μm

efficiently competed for resources instead of directly attacking the host. Druzhinina and Kubicek [9] speculated that exclusively tropical T. reesei, which has been rarely isolated from soil, was not able to recognize temperate soil-borne T. cucumeris as its host to prey. In confrontations with Alternaria alternata (Pleosporales, Ascomycota) and Botrytis cinereal (Helotiales, Ascomycota), T. reesei inhibited their growth, and in the case of B. cinerea, the mycelium was overgrown and killed [64]. Druzhinina et al. [35] investigated interactions between T. reesei and the lignocellulolytic Pestalotiopsis fici (Xylariales, Ascomycota) which was identified several times as one of the putative LGT donors. The latter species was also selected owing to its frequent isolation from similar ecosystem as that of T. reesei and also because they have comparable growth rates. Endoparasitism was revealed through confocal microscopy, showing the penetration of P. fici hyphae by that of T. reesei (Fig. 2). Dual confrontation assays were also set between T. reesei and some putative donors from the Eurotiales order, such as Penicillium spp. Although endoparasitism could not be seen in this case, T. reesei showed the capability of attacking these fungi as well [35]. T. reesei has also been shown to parasitize a fungus-like protist, Pythium ultimum (Oomycota) [78]. Although the latter is not a fungus, this demonstrates the endoparasitic capabilities of T. reesei. Hence, it is essential to note that just like other Trichoderma species, T. reesei can exploit a variety of mycoparasitic strategies depending on the hosts or interaction partner.

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Trichoderma reesei Is No Longer Rare Because the entire pedigree of industrial strains of T. reesei is based on the single isolate QM6a (vide supra), the species may be considered as rare. However, the recent study of Kubicek et al. [29] listed T. reesei among the most common Trichoderma species. For this chapter, we have updated the inventory of T. reesei isolates recorded in the public databases for nucleotide sequences that was initially performed by Druzhinina and Kubicek [9] (Fig. 3). We used the large, fourth intron of the gene encoding translation elongation factor 1-alpha, tef1 [81], that is a powerful DNA fragment suitable for molecular identification of Trichoderma by DNA barcoding. Maximum likelihood analysis revealed records for at least 48 strains that could be reliably identified as T. reesei (see Fig. 3) and at least 14 strains that belong to the sister species T. parareesei. Six strains that were monophyletic with T. parareesei sensu stricto may represent a still undiscovered taxon (including TUCIM 524—C.P.K. 524, GB Accession number GQ354349) that was already evident in Druzhinina et al. [26], but not recognized by Samuels et al. [27]. The resulting analysis of the biogeographic distribution of T. reesei and T. parareesei sensu lato (including the putative new taxon) still confirmed the earlier claims of Druzhinina et al. [64] concerning the most tropical occurrence of T. reesei. However, its detection in Japan and Argentina expands its possible occurrence to the latitudinal belt of 30–35 around the equator. The more ecologically versatile T. parareesei (Figs. 1 and 3) was initially described based on six isolates from soil [9, 64, 82], and the currently available diversity of this species confirms the sympatric occurrence with T. reesei on a large biogeographic scale. The reason for the affinity of both fungi to tropics is still not known [9]. We note, however, that T. reesei exhibits a high rate of exchange of genetic material over these vast geographic distances, without evidence for geographic segregation ([64]; Fig. 1); its biogeographic restriction can therefore not be due to a limited dispersal. Interestingly, the single strain of T. gracile that is known so far was isolated from Malaysia, while the newly recognized T. beinertii was detected in a marine sponge in the Mediterranean Sea near Israel and in soil in South Africa [83]. It is thus possible that this geographic specialization occurred already at the ancestor of several species related to T. reesei and T. parareesei because T. longibrachiatum and its neighboring species T. orientale, T. citrinoviride, and others are highly cosmopolitan and found at all latitudes [84]. Although T. reesei and T. parareesei share the same phylotype of the internal transcribed spacer 1 and 2 of the rRNA gene cluster (ITS1 and 2 of the rRNA) [64, 82], we checked all ITS1 and 2 rRNA sequences deposited for

Fig. 3 Maximum likelihood phylogenetic tree constructed based on a multiple sequence alignment of the partial tef1 locus from T. reesei, T. parareesei, and their closely related species. The tree was constructed by using IQ-TREE 1.6.12 and annotated using the online tool Interactive Tree Of Life (iTOL v4, https://itol.embl.de). Node labels indicate IQTree ultrafast bootstrap support values >90 (calculated from 1000 ultrafast replicates). The information regarding the origin and the substrate for strain isolation are given by the square and circle symbols, respectively. Sequence IDs in public databases (GenBank) are given for all strains unless indicated by an asterisk which indicates unpublished sequences from I. S. Druzhinina’s group. The tef1 sequences of the type strains are marked by a superscript T. Numbers in the insert show the records for individual substrates and origins for T. reesei

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these two species in GenBank (referring to an identity of >99% similarity to the sequence of T. reesei QM6a Z31016 deposited in GenBank). It revealed 38 sequences, all from tropical habitats over the world (October 13, 2019). The analysis of the habitat of T. reesei sensu stricto (Fig. 3) indicates deadwood or its derivatives (e.g., bark, logs, decorticated wood) as the most frequent ecological niche for T. reese.i And at least six strains were isolated from soil (Fig. 3) and one (CBS 999.97) from lake sediment. Interestingly, there are still no T. reesei isolates found on other fungi, and only one strain of T. parareesei is isolated from Lentinula edodes (Agaricales, Basidiomycota) in Japan [9]. It is interesting to note a sample of T. parareesei from UK air [85]. Although it may be an artifact, we also detected an ITS1 and 2 MOTU of either T. reesei or T. parareesei in air samples in Vienna (Austria [9]). These findings correspond well to the hypothesis of the efficient long-distance dispersal of these fungi. However, all attempts to cultivate T. reesei from those air filters failed (I.S. Druzhinina, unpublished). The inventory of habitats for the deposited ITS1 and 2 sequences of T. reesei or T. parareesei revealed several strains isolated from Bradypus variegatus (sloth) fur [86], several marine isolates from Thailand, Brazil, and Malaysia, one clinical sample from H. sapience in Malaysia. In this survey, we also detected a second record of the association of T. reesei–T. parareesei pair with fungi. The strain SPH-2010-9-132 was isolated from Ganoderma boninense (Polyporales, Basidiomycota) in Singapore and deposited as T. saturnisporum. However, the ITS1 and 2 sequences of this isolate (KY025558) allow its identification as either T. reesei or T. parareesei. Thus, T. reesei is observed in vitro as a species with a relatively weaker fitness compared to T. parareesei ([64, 82], Fig. 1), and at the same time, it is inherently more common and widespread. However, as it is not known from the whole range of habitats such as opportunistic environmental species (T. harzianum, T. longibrachiatum, T. asperellum, and some others), it is still considered to be relatively ecologically specialized. Unfortunately, little is still known about the occurrence of the T. reesei anamorph in natural environments. Further studies will ultimately reveal ecological niches occupied by all stages of the T. reesei life cycle, which will open new opportunities for the further domestication of this useful microorganism.

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Conclusions The superior cellulolytic activity of T. reesei has led to the scientifically advantageous situation for this species compared to other Trichoderma spp. because of a significant bulk of information that

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is available. Thus, in October 2019, T. reesei was cited in more than 2200 PubMed indexed publications, while the by far the most common Trichoderma species—T. harzianum [29]—was referred in only 1106 articles. The genome of T. reesei is best studied in the genus, and the molecular toolbox available for this species allows the broadest range of genetic manipulations (see ref. 5 for a review). However, ironically, instead of representing the model for Trichoderma research, T. reesei is an exception in the genus. Thus, although T. reesei can parasitize other fungi, this important generic trait is not expressed sufficiently strong for the studies of fungal– fungal interactions. Only a limited number of host fungi are known for T. reesei. In many such investigations, T. reesei is presented as a control saprotroph, while its mycoparasitism remains overlooked [28, 87]. Similarly, although T. reesei is a potent industrial cellulase producer, other Trichoderma species have essentially more pcwdCAZymes encoded genes in their genomes [35] and consequently secrete more efficient cellulolytic cocktails that may find their way to commercial production [88–92]. In this chapter, we did not review the in vitro mating behavior of T. reesei (see ref. 9 for a review), which is another unique feature of this species used for industrially relevant genetic improvement of the mutant strains [52, 93]. No other Trichoderma species are known to produce fertile fruiting bodies in laboratory conditions. It is likely that some or all of T. reesei features positively influenced the establishment of this species as an industrial microorganism, allowing efficient manipulation of its growth and development. However, care should be taken when the advanced technologies available for T. reesei are used for the investigations of the biology, physiology, and evolution of other Trichoderma species, the genus Trichoderma and other hypocrealean fungi.

Acknowledgments The authors are thankful to Christian P. Kubicek for the discussion of the topic and useful suggestions to the content of the chapter. This work was supported by grants from the Ministry of Science and Technology of Jiangsu Province (BK20180533), China, the National Natural Science Foundation of China (KJQN201920), and the Postdoctoral Science Foundation (198162), all to F.C. The work in Vienna (Austria) was supported by the Austrian Science Fund (FWF) P25613-B20 and P25745-B20, to I.S.D. and the Vienna Science and Technology Fund (WWTF), LS13-048, to I. S.D.

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Chapter 2 Industrial Relevance of Trichoderma reesei as an Enzyme Producer Amanda J. Fischer, Suchindra Maiyuran, and Debbie S. Yaver Abstract Trichoderma reesei’s potential as a rapid and efficient biomass degrader was first recognized in the 1950s when it was isolated from Army textiles during World War II. The microbe secreted cellulases that were degrading cotton-based tents and clothing of service members stationed on the Solomon Islands. In the 1970s, at the time of the first global oil crisis, research interest in T. reesei gained popularity as it was explored as part of the solution to the worlds growing dependence on fossil fuels. Much of this early work focused on classical mutagenesis and selection of hypercellulolytic strains. This early lineage was used as a starting point for both academic research with the goal of understanding secretion and regulation of expression of the complex mixture of enzymes required for cellulosic biomass decay as well as for its development as a host for industrial enzyme production. In 2001, at the onset of the second major oil crisis, the US Department of Energy supported research programs in microbial cellulases to produce ethanol from biomass which led to another surge in the study of T. reesei. This further accelerated the development of molecular biology and recombinant DNA tools in T. reesei. In addition to T. reesei’s role in bio-ethanol production, it is used to produce industrial enzymes with a broad range of applications supporting the bio-based economy. To date there are around 243 commercially available enzyme products manufactured by fermentation of microorganisms; 30 of these are made using Trichoderma as a host, 21 of which are recombinant products sold for use in food, feed, and technical applications including textiles and pulp and paper. Key words Protein production, Heterologous protein, Recombinant protein, Trichoderma reesei, Recombinant DNA technology, Cellulases, Bio-based economy

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Identification and Development of T. reesei as a Biotechnology Tool Trichoderma reesei’s potential was first recognized over 70 years ago during World War II when it was isolated from US Army textiles that were being degraded through the activity of this microbe in the Solomon Islands. This mold was isolated for its potential value as a rapid biomass degrader, and through screening of wild-type strains using crystalline cellulose as a substrate the type strain QM6a was identified [1, 2]. The initial isolate, and study of it, was carried out by the US Army Research Center with the purpose of finding ways

Astrid R. Mach-Aigner and Roland Martzy (eds.), Trichoderma reesei: Methods and Protocols, Methods in Molecular Biology, vol. 2234, https://doi.org/10.1007/978-1-0716-1048-0_2, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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to inhibit the cellulase enzymes that were degrading cotton clothing and tents [3]. The isolate was originally classified as Trichoderma viride and was renamed in 1977 to Trichoderma reesei [4]. Around this time, the first oil crisis occurred, and there was renewed focus on the utilization of microbial cellulases for the hydrolysis of cellulosic raw materials to sugars that could be used for fuel production. This interest initiated research to isolate hypercellulolytic mutants of QM6a using classical mutagenesis followed by screening and/or selection [5]. In 2001, the US Department of Energy supported research programs in microbial cellulases to produce ethanol from biomass which led to another surge in the study of T. reesei. The volume of research on T. reesei, as evident by the number of patents filed and scientific publications per year, tracks the increase in oil price over the last 50 years as well as the development of recombinant methods for T. reesei (Fig. 1). Fast forward 70 years from the isolation of QM6a and T. reesei is used widely for the production of native and recombinant enzymes enabling the bio-based economy. During the first oil crisis in the 1970s, improved T. reesei strains were chosen for industrial enzyme production [2, 6]. Many industrial strains have the common ancestor, RUT-C30, which is the cornerstone hypercellulolytic strain for both academic research and industrial production [7]. RUT-C30 was isolated at Rutgers University using a three-step mutagenesis campaign [5, 8, 9]. M7 was isolated following UV-light-induced mutagenesis of QM6a and screening for the ability to hydrolyze cellulose under carboncatabolite repressing conditions. Strain NG14 was obtained after NTG chemical mutagenesis of M7 using similar screening parameters. NG14 was exposed to UV mutagenesis, and mutants resistant to 2-deoxyglucose were selected and assayed for cellulase activity. Each step led to improved extracellular protein and cellulase activity as well as reduced catabolite repression [10, 11]. RUT-C30 was reported to produce 15–20 times more extracellular protein and cellulase activity than QM6a [11]. The lineages of hypercellulolytic mutants can be found in a review on biochemistry of T. reesei cellulases, strain improvement, process improvement, and molecular cloning [3]. These mutants of T. reesei have been both a starting point for academic research with the goal of understanding secretion and regulation of expression of the complex mixture of enzymes required for cellulosic biomass decay and for the development of hosts for industrial enzyme production. Classical mutagenesis remains a productive route for creating interesting mutants with altered abilities for cellulosic biomass decay and increased extracellular protein production. Studies to understand the physiological, metabolic, and genomic differences between QM6a and RUT-C30 have shed light on changes responsible for increased cellulase activity and increased

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Fig. 1 Inflation-adjusted crude oil prices (USD/barrel), number of academic journal publications, and number of patent applications published per year [67] (PatBase: Search in title, abstract, and claims: Trichoderma reesei or Hypocrea jecorina and enzymes ¼ 479 published applications. Web of Science Biosis article finder: Search in abstract: Trichoderma reesei OR Hypocrea jecorina AND Enzyme ¼ 3647 Academic Journal ARTICLES). Key developments in oil costs. 1973 is marked as the first oil crisis when prices increased 400% which coincides with the first phase of development of Trichoderma reesei for biofuel production. Again in 2003, prices started to rise quite sharply finally stabilizing in 2008 where they hovered around an all-time high. Since 2015, prices have fallen. Key milestones in T. reesei research. In the 1950s, T. reesei was identified and recognized for its cellulose degrading properties [2, 68]. Hypersecreting cellulase strain QM6a was isolated in 1977 from the US Army’s Quarter Major culture collection [4]. In 1979 RUT-C30 was isolated through mutagenesis having a 20-fold increase in secreted protein over QM6a [5, 9]. In the 1980s, the major cellulase gene cbhI was cloned, and recombinant DNA tools were developed for T. reesei [27, 35, 69]. In 2007, the US government awarded $385 million in grants aimed at jump-starting ethanol production from nontraditional sources including cellulosic biomass-based hydrolysis. In 2008, the genome sequence of QM6a was published [15]. In 2015, the first report on the use of CRISPR for genome editing in T. reesei was published [41]

secretory protein production in RUT-C30. However, it is still not understood if all mutations, indels and rearrangements responsible for the desired phenotype have been identified. Based on electron microscopy, it was observed that RUT-C30 contained an enrichment of ER content and was described as lacking typical Golgi bodies, instead containing many individual ER-associated vesicles. Many of these abnormal structures were consistent with cellular stress [12]. Unlike the type strain QM6a, RUT-C30 produces cellulases even under carbon catabolite conditions; isolation of the cre1 gene from RUT-C30 demonstrated that it has a truncation [13]. This truncation leads to a gene encoding only one of the two usual zinc finger regions of the CRE1 protein which explains the hypercellulolytic phenotype observed under carbon repression

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conditions. It was also observed that glycoproteins from RUT-C30 have an aberrant glycosylation pattern containing a terminal apha1,3-glucose residue which could be due to insufficient glucosidase II activity. It was shown that a frameshift mutation in the glucosidase II gene is responsible for this phenotype and that repairing this mutation led to a change in glycosylation although not back to that observed in wild-type strains, as well as a reduced capacity for protein secretion [14]. In 2008, the genome sequence for QM6a was published which provided a tool for further elucidation of genome changes that occurred in RUT-C30 [15]. An 85 kb genomic fragment containing 29 genes was found to be missing between RUT-C30 and its ancestor NG14; this fragment contains genes involved in primary metabolism, transporters, extracellular enzymes, and proteins involved in detoxification [16]. A comparison of the genomes of RUT-C30 and its parent NG14 found it to be missing over 100 kb of genomic DNA present in QM6a encompassing 18 large deletions, 15 small indels, and 223 single nucleotide variations [17]. The genome sequence and comparative genomics have provided many interesting scientific questions to elucidate all the single nucleotide polymorphisms and indels that contribute to the phenotypes displayed by the T. reesei mutants; however, there is still not complete clarity on all the genomic changes in RUT-C30 that impart its desirable phenotypes. Non-recombinant QM6a and its derivatives including RUT-C30 have been used commercially to produce native enzyme products that have been used in food, feed, and technical (including textiles and pulp/paper) applications (Table 1). T. reesei QM6a has been used for the commercial manufacturing of cellulases for textile applications since the 1960s [18]. In the late 1970s, the use of commercial microbial preparations of T. reesei cellulases for a variety of industrial applications expanded, including food industry for baking, malting, brewing, and grain alcohol production; animal feed for increasing digestibility and even in the pharmaceutical industry, where cellulases have been used as digestive aids [19– 22]. Cellulases have also been used in wood processing and textile applications [22, 23]. The birth of recombinant DNA technology and the development of tools to engineer T. reesei have expanded the number of enzyme products produced from QM6a and its derivatives (Table 1).

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Development of T. reesei as a Host for Recombinant Enzyme Production T. reesei was an obvious host to develop for recombinant protein production for a variety of reasons. Many decades of classical mutagenesis and screening for strains with high secretion capacity in both academic and industrial research laboratories led to the

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Table 1 A list of the 30 commercial enzyme products produced using Trichoderma as a host; 26 of these utilize T. reesei as a host of which 21 are recombinant enzymes Non-recombinant

Application

Production organism

Enzyme activity

Food Feed

Trichoderma reesei

Xylanase

X

X

None

Trichoderma reesei

Pentosanase

X

X

None

Trichoderma reesei

Hemicellulase

X

Trichoderma reesei

Glucanase (endo-1,3(4)- X beta)

X

X

None

Trichoderma reesei

Cellulase

X

X

None

X

Technical Enzyme donor

None

Total: 5 Recombinant

Application

Production organism

Enzyme activity

Food Feed

Technical Enzyme donor

Trichoderma reesei

Amylase (alpha)

X

X

Aspergillus

Trichoderma reesei

Catalase

X

Aspergillus

Trichoderma reesei

Cellulase

X

Staphylotrichum

Trichoderma reesei

Cellulase

X

X

X

Trichoderma

Trichoderma reesei

Glucanase (endo-1,3(4)- X beta)

X

X

Trichoderma

Trichoderma reesei

Glucoamylase or Amyloglucosidase

X

X

Trichoderma

Trichoderma reesei

Glucosidase (alpha)

X

Trichoderma reesei

Glucosidase (beta)

X

Trichoderma

Trichoderma reesei

Laccase

X

Thielavia

Trichoderma reesei

Mannanase (endo-1.4beta)

X

X

Trichoderma

Trichoderma reesei

Pectin lyase

X

X

X

Aspergillus

Trichoderma reesei

Pectin methylesterase or X Pectinesterase

X

X

Aspergillus

Trichoderma reesei

Phospholipase A2

X

X

X

Aspergillus

Trichoderma reesei

Phospholipase B

X

X

Aspergillus

Trichoderma reesei

Phytase

X

Aspergillus

Trichoderma reesei

Phytase

X

X

X

Buttiauxella (gram –ve bacteria)

Trichoderma reesei

Polygalacturonase or Pectinase

X

X

X

Aspergillus

Aspergillus

(continued)

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Table 1 (continued) Trichoderma reesei

Protease (incl. X Milkclotting enzymes)

Trichoderma reesei

Xylanase

Trichoderma reesei

Xylanase

X

Trichoderma reesei

Xylanase

X

X

Trichoderma Actinomadura (gram +ve bacteria)

X

Aspergillus X

X

Trichoderma Total: 21

Other Trichoderma sp.

Application

Production organism

Enzyme activity

Food Feed

Trichoderma viride

Xylanase

X

None

Trichoderma viride

Cellulase

X

None

Trichoderma harzianum Glucanase (endo-1,3(4)- X beta)

None

Trichoderma harzianum Glucosidase (exo-1.3beta)

None

X

Technical Enzyme donor

Total: 4 EFSA approved (post2015 AMFEP list)

Application

Production organism

Enzyme activity

Food Feed

Technical Enzyme donor

Trichoderma reesei

Trehalase glucohydrolase (alpha)

X

unknown unknown unknown

Trichoderma reesei

Muramidase

X

unknown unknown unknown

Trichoderma reesei

Xylanase (endo-1,4beta)

X

unknown unknown unknown

Trichoderma reesei

Amylase (alpha)

X

unknown unknown unknown

Trichoderma reesei

Lysophospholipase

X

unknown unknown unknown Total: 5

This list is adapted from the 2015 Association of Manufacturers and Formulators of Enzyme Products (AMFEP) list that contains 243 commercial enzymes manufactured by fermentation that have been approved for various food, feed, or technical applications [64]. There are five recent European Food Safety Authority (EFSA) food-approved enzymes made using T. reesei as a host; these approvals were published after publication of the AMFEP 2015 list [65, 70–73]

identification of strains producing upwards of 100 g/l of native cellulases and hemi-cellulases under controlled fermentation conditions [24]. In addition to robust secretion, cultivation under industrial scale conditions utilize cheap raw materials with enzyme recovery being vastly simplified as the product is secreted directly

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into the broth. Recovery of secreted proteins involves only a few, inexpensive steps: removal of cell debris, concentration, stabilization, sterile filtration, and formulation. In contrast, the recovery process for pharmaceutical products typically requires additional expensive steps including cell lysis and chromatographic separation. T. reesei has a long history of safe use for the production of enzymes for technical, feed, and food applications dating back to the 1960s which in part has led to its status as a microbe that is Generally Regarded as Safe (GRAS) for the production of ingredients as food additives by the US Food and Drug Administration [18–23, 25, 26]. Taking this framework or “chassis” with high potential for native protein production and enabling its evolution into a host for recombinant enzyme production required development of basic molecular biology tools including DNA transformation, selection for integrated recombinant DNA, and genome editing. The fungal cell wall poses a significant barrier to introduction of recombinant DNA into cells. Similar challenges arose for the development of recombinant DNA technology for plants and other fungi. In plant biotechnology, three common methods for overcoming the barrier to DNA insertion posed by the cell wall include the physical removal of the cell wall through the process of preparing protoplasts and use of polyethylene glycol (PEG)-mediated DNA uptake, transport of DNA through the cell wall using Agrobacterium as a DNA carrier, and the use of biolistic DNA-coated particle bombardment to break through the cell wall and introduce the DNA. These techniques have been adapted for use in T. reesei. Reports of PEG-mediated DNA uptake into T. reesei protoplast appeared in 1987. This process requires the digestion of the fungal cell wall of young germlings using different enzyme cocktails usually containing a mixture of chitinase and beta-glucanase activity and large amounts of recombinant DNA carrying an appropriate selection marker [27]. Although this process is functional, the efficiency is very low and varied depending on the method of DNA selection and mode of integration (homologous targeting, ectopic integration by non-homologous end joining (NHEJ), or recombinase-mediated integration systems like loxP/CRE). Reports on the use of Agrobacterium-mediated transformation into T. reesei using either protoplasts or conidia as the starting point came much later in 2007 [28]. Using Agrobacterium usually achieved higher transformation efficiency than PEG-mediated protoplast transformation; however, it came with other complicating factors including the added steps of transformation of the desired integrating-DNA into Agrobacterium and then ensuring the Agrobacterium was lost following transformation. Biolistic transformation was reported in 2000, but with lower transformation efficiency than PEG-mediated transformation and required coating of expensive gold or tungsten disks with micro-gram quantities of DNA that are then “shot” into the conidia or cells [29]. One advantage of

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biolistic transformation is the high rate of co-transformation where the carrier particle is coated with both DNAs to be integrated. It was also advantageous that biolistic transformation did not require preparation of protoplasts or a bacterial carrier strain. Spore electroporation has also been reported to be successful with similar efficiencies to PEG-mediated transformation, but the publication record is sparse with respect to this method [30]. Although reliable methods for transformation of recombinant DNA into T. reesei exist, transformation efficiency is still a bottleneck. To take full advantage of current synthetic biology tools to optimize recombinant DNA expression cassettes and genome editing tools for optimization of heterologous secretion, transformation efficiency needs to be improved by multiple orders of magnitude. Enabling approaches could include the enrichment for populations of protoplasts that are viable and competent combined with more efficient systems for recombinase-mediated DNA integration. Solutions to these challenges will propel the development and utility of T. reesei as a recombinant protein production host. Once the DNA has been introduced into the cell and integrated into the genome, efficient methods for selection are required to identify the transformed cells since they are rare, low efficiency events. A suite of dominant and auxotrophic selection markers has been developed for use in T. reesei. The first step in enabling an auxotrophic system is the deletion of an enzyme in the host, making it unable to synthesize a specific nutrient necessary for survival. This gene can then be included on the recombinant DNA or co-transformed with the DNA of interest, so that prototrophy is regained once the complementing DNA is integrated into the genome. Auxotrophic strains were first generated through spontaneous or random mutation, later through disruption using homologous recombination (HR). The commonly used auxotrophic systems in T. reesei include arginine biosynthesis (requiring the ornithine transcarbamylase gene, argB) and uridine biosynthesis (requiring orotidine 50 -monophosphate decarboxylase ura3/pyr4/ pyrG or orate phosphoryribosyltransferase ura5) [27, 31– 33]. Dominant selection markers are more versatile because they do not require engineering of an auxotrophic host as a prerequisite; however, they do require that the host is sensitive to the selection agent being used. Dominant selection markers commonly used in T. reesei include the E. coli hygromycin phosphotransferase gene hph that confers resistance to the antibiotic hygromycin and the A. nidulans acetamidase gene amdS which allows for survival on acetamide as the sole nitrogen source since T. reesei naturally lacks a copy of the amdS gene [34]. Once the cell takes up the DNA and there is a method for selecting cells containing the recombinant DNA, the next challenge is to achieve the desired level of transcription, translation, and secretion to harness T. reesei’s 100 g/l potential to produce a

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foreign, recombinant protein. Since approximately 60% of this total protein produced by high secreting strains is the result of the expression and secretion from one major cellulase protein, cellobiohydrolase I (CBHI), it made sense to utilize the cbhI gene’s key transcriptional and translational machinery as a starting point for recombinant gene expression. The cbhI gene was cloned, and its regulatory components were identified in 1983 [35, 36]. Soon after in 1989 the first report of recombinant protein production using T. reesei was published [34]. This work demonstrated the use of the T. reesei cbhI gene promoter and terminator for transcription, as well as a protein fusion to part of the mature cbh1 as a technique for effective secretion of bovine chymosin. Recombinant production of chymosin was a challenging starting point as it requires precise auto-catalytic, low-pH pro-peptide processing to form the active enzyme. At the time, calf chymosin had been demonstrated to be secreted at very low levels using other recombinant microbial protein production hosts including S. cerevisiae, A. nidulans, and E. coli. Interestingly, active chymosin was secreted from T. reesei at levels exceeding those of the other microbial hosts with proper preand pro-peptide processing. This was a very promising starting point for the development of T. reesei as a recombinant protein production host. Since then, a variety of different mammalian, bacterial, and fungal proteins have been successfully produced using T. reesei. Many of these reports describe using similar tools to those first developed for expression and secretion of bovine chymosin. Most of the examples of industrial scale recombinant protein secretion using T. reesei as a host are cases where Ascomycota gene donors are used and can be produced at yields sufficient to meet industrial-scale production economy targets (Table 1). Despite efforts to improve heterologous protein production using RUT-C30, all reported yields of recombinant protein products are considerably lower than that of the native cellulases [24, 37]. Additional technology development is required to fully realize the protein production potential of T. reesei. Tools have also been developed to better control integration locus and the number of copies of the recombinant gene integrated into the genome. Initial attempts at recombinant gene integration took advantage of random integration using the native NHEJ pathway and co-transformation with a selection marker. This method required intensive screening to identify transformed cells that contained both the recombinant gene of interest and the selection maker. Co-localizing the selection marker with the recombinant gene of interest on the same plasmid enabled selection though screening was still necessary as recombinant protein yields varied among transformants due to integration copy number and locus of integration [34]. Deletion of the ku70 gene to disable the NHEJ pathway resulted in a significant improvement in the number of HR events that occurred upon transformation but required

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the use of large target locus flanking sequences (1–2 kb) to direct recombinant DNA integration [38]. Strains with the ku70 deletion have a gene targeting efficiency of approximately 95%; however, transformation efficiency is much lower than that of random integration using NHEJ. Homologous integration reduces the burden of screening for targeting to high production loci, enabling replacement of major cellulases with copies of the recombinant gene which reduces the complexity of the secretome. Homologous integration also allows for more controlled side-by-side experiments where copy number and locus of integration are held constant and other variables can be tested. Another method for recombinant DNA integration that has been successful for T. reesei is the use of the recombinant bacterial recombinase-mediate DNA integration system loxP/CRE [39]. This system requires a pre-engineered host strain containing “landing pad” recognition sequences and counter-selectable markers placed at strategic locations in the genome like the cbhI and/or cbhII loci as target sites for recombinant DNA integration. These strains are also typically deficient for CBHI and CBHII which eliminate a majority of the secreted proteome. The use of ku70 deletion hosts has also been helpful in reducing the incidence of off-target or random integration when using heterologous recombinase-mediated gene integration systems like loxP/CRE [39]. Another enzyme-mediated DNA integration system is the Saccharomyces cerevisiae I-SceI mega-nuclease which is used for the generation of site-specific double-stranded breaks (DSB) that can be repaired using DNA cassettes flanked by regions of homology [40]. SceI recognizes an 18-basepair restriction site, so like the loxP/CRE systems, it requires a pre-engineered strain for targeting. One benefit of SceI and other recombinant recombinase-mediated systems over the use of NHEJ and HR is improved transformation efficiency and improved genetic stability of the transformed cells. Advancement of the tools used for targeted DNA integration has allowed for controlled side-by-side experiments where copy number and locus of integration are held constant and other variables can be tested. This has enabled researchers to explore other factors influencing recombinant protein production such as changes to the genome and modification to the recombinant gene expression cassette itself. This decade has seen a large suite of new recombinant genome editing tools including zinc-finger nucleases (ZFNs), transcription activator-like effector nucleases (TALENs), and the type II clustered regulatory interspaced short palindromic repeats (CRISPR) RNA-guided CRISPR-associated gene 9 (Cas9) (CRISPR/Cas9), all of which can be used as so-called molecular-scissors. These nucleases allow for targeted generation of DSB which can be repaired using DNA with homology flanks to the cut site or by non-homologous end joining. Both TALENS and CRISPR/Cas9 have been demonstrated in T. reesei, and both have very high

Industrial Relevance of T. reesei

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on-target activities [41, 42]. Cas9 catalyzes a DSB at the desired DNA target-site using an engineered guide RNA (gRNA) that contains a site-specific 20-bp “protospacer” sequence and an adjacent downstream 50 -NGG sequence (aka protospacer-adjacent motif, PAM). This system requires construction of a new gRNA for each target site of interest and is constrained by the need for the presence of a PAM sequence near the desired target site. TALENs are customizable restriction enzymes consisting of an array of repeat sequences which encode the target site-specific DNA-binding domains fused to a nonspecific nuclease domain. TALENs are not limited by target-site DNA requirements, and the DNA-binding domain can be modified to direct DSB to most locations in the genome. To target TALENs to different genome locations, the DNA-binding domain needs to be engineered with a target-site-specific sequence which requires complex molecular biology techniques due to the array of repetitive sequence in this domain. Target-specific gRNA construction for the CRISPR/Cas9 system can be carried out using standard molecular biology techniques making it a preferred approach for genome editing. The availability of different CRISPR-associated genes with different PAM sequence specificity has made the CRISPR toolbox more versatile. Over the last 32 years, significant advances in the tools used for recombinant DNA technology in T. reesei have led to a respectable repertoire of publications, patents, and recombinant DNA products made using T. reesei as a host (Figs. 1 and 2 and Table 1). Early work using the cbhI promoter and locus for recombinant gene expression allowed for high levels of transcription. Now, more challenging bottlenecks can be addressed using CRISPR-based genome editing to unlock the full secretion potential of T. reesei for recombinant production.

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Challenges for Heterologous Protein Secretion Using T. reesei Early reports on heterologous protein production using RUT-C30 were considerably lower than that of the native cellulases [24, 37]. One of the earliest reports of non-fungal heterologous protein production using T. reesei as a host includes the secretion of active bovine chymosin, which is used for milk-clotting in the cheese-making process [34]. Chymosin secretion reached yields of 40 mg/l, significantly better than other recombinant hosts tested at the time. Another early report of recombinant protein production using T. reesei was for the production of antibody FAB fragments [43–45]. This was the first example of the secretion of an active, assembled, multi-chain protein using T. reesei as a recombinant host. Yields from T. reesei were similar to those obtained from hybridoma cell lines and exceeded those of other microbial

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Fig. 2 Breakdown of the most commonly used microbial hosts for the production of commercial enzymes; adapted from the 2015 Association of Manufacturers and Formulators of Enzyme Products (AMFEP) list that contains 243 commercial enzymes manufactured by fermentation that have been approved for various food, feed, or technical applications [64]. Percent of total products is rounded down

production systems tested at the time. Successful secretion of a small peptide using T. reesei was also demonstrated using the hormone peptide obestatin (23 amino acids) [46]. In general, reports on heterologous secretion using more distant donors have only achieved low yields ranging from tens of milligrams to a gram per liter. There has been greater success reported for the heterologous production of fungal proteins in T. reesei, likely owing to the evolutionary proximity of the host and donor. Early examples include Hormoconis resinae glucoamylase and Aspergillus niger phytase, which achieved yields of 2–3 g/l in fermenters [47–49]. More recent examples of industrial scale production of recombinant, non-cellulases or hemicellulases, from T. reesei include amylase, protease, laccase, pectin degrading enzymes, phospholipase, and phytase, likely reaching yields significantly greater than 1 g/l (Table 1). Again, many of these recombinant genes are either from Trichoderma donors or other closely related Ascomycota like Aspergillus (Table 1). Studies comparing transcription and translation efficiencies of native cellulases, recombinant protein, and native-recombinant fusion proteins tell an interesting story about the different bottlenecks that exist for heterologous protein production in T. reesei. There are multiple examples where transcription was a limitation to heterologous production in T. reesei, even when using the strong

Industrial Relevance of T. reesei

35

cbhI promoter. The mRNA levels for bovine chymosin were only 1–5% of cbhI mRNA levels and only very low levels of the heavy chain Fb antibody fragment could be detected [34, 43–45]. Fusion of recombinant proteins of interest to a well-expressed native “carrier-protein” is a common approach to improve recombinant protein production. Several reports of fusion proteins resulting in improved secretion from T. reesei have also been published. One common variation is to use the T. reesei CBHI protein, which is composed of a catalytic domain and cellulose binding domain (CBD) separated by a flexible linker. The CBD can be replaced with the fusion protein of interest, and protease cleavage sites can be engineered into the linker, so that the fusion becomes separated during secretion. In many cases, cleavage in the linker spontaneously occurs by the action of native proteases during secretion, resulting in two separate protein products in the broth. Detailed studies on the steady-state mRNA levels in strains containing these fusions suggest that the CBHI fusion increases steady-state mRNA levels and may also help remove bottlenecks in the folding and secretion process for non-host proteins. This approach was successfully used to improve the secretion of bovine chymosin, Fb antibody fragments, and other recombinant proteins [22, 34, 43, 44]. Fusing the mature peptide of CBHI to the heavy chain Fb antibody fragment significantly increased mRNA levels by either improving transcription or stabilizing the mRNA. In this case, the CBHI-core-linker was secreted to very high levels, but the cleaved Fb antibody was still only secreted at low levels; this suggests that there is a disruption in the secretion process downstream of transcription and translation. Another interesting example of optimization of heterologous production in T. reesei was reported for the A. nidulans class I hydrophobin, DewA. When DewA was expressed using the cbhI promoter, only low levels of transcript, and no recombinant protein could be detected in the cell, associated with the cell wall or secreted into the media [50]. When DewA was expressed using the native T. reesei hydrophobin 2 (hfb2) promoter, higher levels of transcript and secreted protein were detected. This is a unique example of promoter-dependent expression and one could speculate that this effect is due to mRNA stabilization. An additional challenge for efficient production of recombinant proteins using T. reesei is the suite of native proteases that are secreted into the fermentation broth. Product stability can be achieved through multiple routes, including (1) selection and screening for recombinant enzymes that are stable when secreted into T. reesei broths, (2) use of enzyme protein engineering to improve stability in the protease environment of the broth, and/or (3) reducing the overall protease profile of the production host through strain engineering or classical mutagenesis and screening. Deletion of specific proteases was shown to be a

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successful approach for improving stability of biopharmaceutical molecules expressed in T. reesei [51, 52]. A systematic approach was initiated to identify the proteases responsible for instability of recombinantly expressed therapeutic antibodies, interferon alpha 2b (IFNa2b) or insulin-like growth factor (IGF1) and to generate a more suitable T. reesei host for their production. A proteomic study using the secretome of QM6a led to the identification of 39 proteases to target for further investigation. Purification and inhibitor studies led to a short list of 13 proteases related to degradation of these therapeutic proteins. Initial work to delete the seven most problematic proteases resulted in a 96% reduction in casein protease activity in the fungal broth. Antibody production in this host led to a significant improvement in product stability [51]. Subsequently, a strain with a total of nine protease deletions was engineered, further stabilizing the secreted IFNa2b [52]. Alternative secretion routes were also explored to avoid proteolysis and to simplify purification of biopharmaceutical molecules produced using T. reesei. One approach was to fuse the recombinant protein of interest to the native T. reesei hydrophobin I protein (HFB1) [53, 54]. Hydrophobins are small (~100 amino acids), surface-active proteins produced by filamentous fungi that contain four disulfide bridges and self-assemble at air–water interfaces. The HFB1 fusion is targeted to the ER for proper folding, disulfide bridge formation, and glycosylation. The fusion then accumulates intracellularly in, inert, soluble protein bodies (PB). PB encapsulation allows the fusion protein to avoid proteolysis while protecting the cell from the recombinant protein. This approach was demonstrated using the native T. reesei endoglucanase I (EGI) and resulted in yields of over 1 g/l. These fusions can then be purified by lysing the cell and applying a simple surfactant-based aqueous two phase purification system [53]. A similar approach used a fusion of the human alpha-galactosidase A (GLA) to the maize gamma zein storage protein (ZERA®). The gamma zein peptide naturally accumulates in the ER, and intermolecular ZERA–ZERA interaction recruits the fusion proteins to form PB structures around the ER membrane [55]. In this example, active human GLA was produced in T. reesei to yields of >0.5 g/l. In addition to extracellular proteases posing a challenge for stability, there are also intracellular quality control mechanisms that impact recombinant protein production in T. reesei. It is not uncommon for recombinant proteins to become mis-folded and accumulate in the ER. Protein mis-folding may be due to a mis-match between the protein translation rates, chaperone requirements for folding, and post-translation modification needs of the recombinant gene or protein and that of the host cell. When mis-folded proteins accumulate in the ER, there are several native cellular responses that help the cell overcome this secretion stress; these responses typically have a negative impact on recombinant

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protein production. Once an accumulation of unfolded proteins is detected in the ER, three pathways can be activated: (1) the repression under secretion stress (RESS) response causes the global downregulation of transcription when the cell senses the accumulation of unfolded protein in the ER, (2) the unfolded protein response (UPR) leads to the upregulation of ER foldases and chaperones to boost re-folding capacity, and (3) if sufficient re-folding is not achieved, ER-associated degradation (ERAD) results in the elimination of misfolded proteins. There are many examples where recombinantly expressed proteins become mis-folded in the ER and stimulate a combination of RESS, UPR, and ERAD. To achieve high yields of recombinant protein production using T. reesei, the reasons for recombinant protein mis-folding need to be understood and addressed. Overcoming this bottleneck will help bridge the gap between the productivity of native protein compared to that of recombinantly expressed protein in T. reesei.

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T. reesei for Broader Bio-Based Applications The bio-based economy is defined as all the consumer goods produced from biological feedstocks and biological processes (Fig. 3). The bio-based economy is driven by microbe and enzyme powered

Fig. 3 On an industrial scale, Trichoderma reesei is used to produce enzymes that enable a sustainable bio-based economy. The bio-based economy is built around the use of microbes and enzymes that serve as catalysts in the biorefineries which are fueled by renewable waste-based carbon feedstocks. These feedstocks consist of starch or sugar-based inputs and/or cellulosic-based inputs (municipal solid waste, MSW). The biorefinery is used to create value through the economical (and renewable) production of food and animal feed, for the generation of renewable fuels, and renewable chemicals and materials for their use in other technical industries. Initially, a majority of T. reesei-produced enzyme products were used for the generation of cellulosic ethanol; however, due to the development of T. reesei as a recombinant host, it is now used to produce enzymes for many of the applications

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biorefineries that utilize feed stocks including biomass from agriculture and forestry, natural gasses and waste from food manufacturing, construction, and municipal sources. It is estimated that this bio-based economy in the US was valued at $205 billion in 2017 [56–60]. Biorefinery products include fuels, chemicals, polymers, feed, and food ingredients that are generated from wastebased feedstocks by microbes and enzymes (Fig. 3). Industrial enzymes are key drivers of the bio-based economy and had an estimated value at 6 billion USD in 2017 [61]. To bring the biorefinery process to reality, production of commercial enzymes must be cost-competitive to allow the products from the biorefinery to compete with the production of cheap petrol-based chemicals. To reduce the cost of commercial enzyme production, enzyme yields must be high, recovery and formulation cheap, and fermentation costs must be kept low (energy input, water, time, feedstock). In the industrial setting, T. reesei is cultivated in liquid media containing cheap feedstocks including cellulose, xylan, or other plant biomass-based substrates that stimulate hypersecretion of a complex mixture of native or recombinant hydrolytic enzymes which can be harvested at the end of fermentation [62, 63]. Industrial scale yields are estimated to approach 100 g/l [10, 24]. The crude enzyme mixture can then be applied to different cellulosic substrates in the biorefinery setting to achieve hydrolysis, resulting in a rich sugar slurry which can then be used for downstream applications including: renewable energy, animal feed, food and beverage, and renewable chemicals and materials. Recombinant DNA tools are also necessary to increase yields of the target enzyme, improve product purity to make recovery and formulation easier, and engineer strains producing recombinant enzymes having improved specific activity and stability under the desired application conditions which vary with respect to the presence of inhibitors, pH, and temperature. T. reesei plays a keystone role in this biorefinery scheme for the industrial-scale enzymatic hydrolysis of biomass waste products into sugar, creating a cheap carbon source that can be transformed into other valuable chemicals using microbial fermentation. During the 1970s, at the time of the first global oil crisis, T. reesei was explored as part of the solution to the world’s growing dependence on fossil fuels (Fig. 1). In this biorefinery scenario, the hope was that T. reesei enzymes could be used for the cost-efficient hydrolysis of biomass to glucose which could be fermented by Saccharomyces cerevisiae to produce fuel ethanol. At the time, the infrastructure and technology were not mature enough to yield a cost-effective solution. Challenges related to the enzymatic hydrolysis included T. reesei enzyme yields, enzyme performance, and stability during hydrolysis. During the period of 2000–2015, around the time of the second global oil crisis, the U.S. Department of Energy made substantial investments in grants to support academic and industry

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labs aimed at jump-starting research into the production of ethanol from nontraditional sources, including cellulosic feedstocks (Fig. 1). The US government also instated aggressive mandates for cellulosic biofuel production to encourage industrial use of these emerging technologies. The infrastructure and technology had advanced significantly since the 1970s and the funding spurred the development of more uniform and accessible biomass substrates, improved efficiency of enzymatic hydrolysis, and more robust processes for the microbial fermentation to ethanol. Recombinant engineering of T. reesei was required to achieve the necessary titers and optimized enzyme formulation to deliver a costcompetitive solution to make enzymatic bioethanol production a reality. Enzyme optimization focused largely on creating a complex of enzymes tolerant to the pH, inhibitors, and the temperatures required for fast hydrolysis of crude, partially digested, highly varied substrates. In addition, enzyme complexes were tested for limiting enzyme factors, and recombinant techniques were used to adjust the overall enzyme ratios to achieve an improved balance for industrial scale hydrolysis. In 2017, the US production of cellulosic ethanol hit a record high of ten million gallons; however, this is only a small part of the total US ethanol production of 15.8 billion gallons, most of which is produced using corn-based feedstocks. Although cellulosic ethanol production through enzymatic hydrolysis is a reality, the market, policy, and technology must align for it to achieve its full potential. In addition to T. reesei’s role in bio-ethanol production, it is used to produce industrial enzymes with a broad range of applications supporting the bio-based economy. For bio-refinery applications, both prokaryotic and eukaryotic microorganisms are used for industrial enzyme production, although most products are made using filamentous fungi as production hosts (Fig. 2). In 2015, there were 243 commercially available enzyme products manufactured by fermentation of microorganisms [64]. Thirty of these products were made using Trichoderma as a production host, 21 of which are recombinant products produced using T. reesei (Table 1). These 30 Trichoderma-based products were approved for different combinations of use in all three biorefinery application areas: food, feed, and technical industries (Table 1). As highlighted previously, T. reesei was initially developed for its potential in enzymatic biomass hydrolysis for fuel ethanol production and is a major contributor to this area through the production of cellulases, catalase, beta-glucosidases, and xylanases. In textile processing, cellulases are sold for cotton softening and denim finishing, as well as for their use in detergent markets for color care, cleaning, and antiredeposition in washing powders. In the chemical area, T. reesei produced hemi-cellulases and laccases are used in the pulp and paper industry as partial replacements for chlorine in the bleaching process. There are five additional T. reesei-produced enzyme

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products approved by EFSA for food grade use which were published after the AMFEP list in 2015 (Table 1). Included in this group is a muramidase produced from T. reesei which was recently approved as a feed additive for chicken fattening [65]. In the feed area, T. reesei is also used to produce recombinant phytases to improve feed conversion for animals. Phytase hydrolyzes phytate in plant-based feeds releasing phosphorous in a form available to the animal, greatly reducing the need for supplemental inorganic phosphorus. In the food area, Trichoderma-produced amylase, trehalase, and glucanase are used for brewing, distilling, and baking applications; proteases for coagulation of milk in cheese making; phospholipases for cheese flavor development; and cellulases, xylanase, and pectinases for fruit juice clarification [66]. As development of recombinant tools for T. reesei advances, its reach into the industrial enzyme market will continue to grow, further enabling the bio-based economy. References 1. Mandels M, Eveleigh DE (2009) Reflections on the United States Military 1941-1987. Biotechnol Biofuels 2:20 2. Siu R, Reese E (1953) Decomposition of cellulose by microorganisms. Bot Rev 19(7):377 3. Montenecourt BS (1983) Trichoderma reesei cellulases. Trends Biotechnol 1(5):156–161 4. Simmons E (1977) Classification of some cellulase-producing Trichoderma species. Second International Mycological Congress, Abstracts 5. Eveleigh DE, Montenecourt BS (1979) Increasing yields of extracellular enzymes. Adv Appl Microbiol 25:57–74 6. Portnoy T, Margeot A, Linke R, Atanasova L, Fekete E, Sandor E, Hartl L, Karaffa L, Druzhinina IS, Seiboth B, Le Crom S, Kubicek CP (2011) The CRE1 carbon catabolite repressor of the fungus Trichoderma reesei: a master regulator of carbon assimilation. BMC Genomics 12:269 7. Peterson R, Nevalainen H (2012) Trichoderma reesei RUT-C30—thirty years of strain improvement. Microbiology 158(Pt 1):58–68 8. Montenecourt BS, Eveleigh DE (1977) Semiquantitative plate assay for determination of cellulase production by Trichoderma viride. Appl Environ Microbiol 33(1):178–183 9. Mandels M, Weber J, Parizek R (1971) Enhanced cellulase production by a mutant of Trichoderma viride. Appl Microbiol 21 (1):152–154

10. Schmoll M, Schuster A (2010) Biology and biotechnology of Trichoderma. Appl Microbiol Biotechnol 87(3):787–799 11. Bisaria VS, Ghose TK (1981) Biodegradation of cellulosic materials: Substrates, microorganisms, enzymes and products. Enzym Microb Technol 3(2):90–104 ˜ o12. Ghosh A, Al-Rabiai S, Ghosh BK, Trimin Vazquez H, Eveleigh DE, Montenecourt BS (1982) Increased endoplasmic reticulum content of a mutant of Trichoderma reesei (RUT-C30) in relation to cellulase synthesis. Enzym Microb Technol 4(2):110–113 13. Ilme´n M, Thrane C, Penttil€a M (1996) The glucose repressor gene cre1 of Trichoderma: isolation and expression of a full length and a truncated mutant form. Mol Gen Genet 251 (4):451–460 14. Geysens S, Dewerte I, Contreras R, Pakula T, Uusitalo J, Penttil€a M (2005) Cloning and characterization of the glucosidase II alpha subunit gene of Trichoderma reesei: a frameshift mutation results in the aberrant glycosylation profile of the hypercellulolytic strain Rut-C30. Appl Environ Microbiol 71(6):2910–2924 15. Martinez D, Berka RM, Henrissat B, Saloheimo M, Arvas M, Baker SE, Chapman J, Chertkov O, Coutinho PM, Cullen D, Danchin EGJ, Grigoriev IV, Harris P, Jackson M, Kubicek CP, Han CS, Ho I, Larrondo LF, de Leon AL, Magnuson JK, Merino S, Misra M, Nelson B, Putnam N, Robbertse B, Salamov AA, Schmoll M,

Industrial Relevance of T. reesei Terry A, Thayer N, Westerholm-Parvinen A, Schoch CL, Yao J, Barbote R, Nelson MA, Detter C, Bruce D, Kuske CR, Xie G, Richardson P, Rokhsar DS, Lucas SM, Rubin EM, Dunn-Coleman N, Ward M, Brettin TS (2008) Genome sequencing and analysis of the biomass-degrading fungus Trichoderma reesei (syn. Hypocrea jecorina). Nat Biotechnol 26 (5):553–560 16. Seidl V, Gamauf C, Druzhinina IS, Seiboth B, Hartl L, Kubicek CP (2008) The Hypocrea jecorina (Trichoderma reesei) hypercellulolytic mutant RUT-C30 lacks a 85 kb (29 geneencoding) region of the wild-type genome. BMC Genomics 9:327 17. Le Crom S, Schackwitz W, Pennacchio L, Martin J, Baker SE, Magnuson JK, Culley DE, Collett JR, Druzhinina IS, Seiboth B, Kubicek CP, Mathis H, Monot F, Margeot A, Cherry B, Rey M, Berka R (2009) Tracking the roots of cellulase hyperproduction by the fungus Trichoderma reesei using massively parallel DNA sequencing. Proc Natl Acad Sci U S A 106(38):16151–16156 18. Toyama N (1969) Application of Microbe to Cellulose Industry Cellulose and Cellulase. Textile Eng 22(8):549–555 19. Home S, Maunula H, Linko M (1983) Cellulases—a novel solution to some malting and brewing problems. Proceedings of the congress—European Brewery Convention:385 20. Linko M (1989) Enzymes in the forefront of food and feed industries. Food Biotechnol 3 (1):1 21. McCleary BV, Gibson TS, Allen H, Gams TC (1986) Enzymic hydrolysis and industrial importance of barley β-glucans and wheat flour pentosans. Starch/Staerke 38(12):433 22. Nevalainen H, Harkki A, Penttil€a M, Saloheimo M, Teeri T, Knowles J (1990) Trichoderma reesei as a production organism for enzymes for the pulp and paper industry. In: Kirk TK, Chang H-M (eds) Biotechnology in pulp and paper manufacture. Applications and fundamental investigations. ButterworthHeinemann, Boston, pp 593–599 23. Tyndall M (1990) Upgrading garment washing techniques. Am Dyestuff Rep 79(5):6p 24. Cherry JR, Fidantsef AL (2003) Directed evolution of industrial enzymes: an update. Curr Opin Biotechnol 14(4):438–443 25. Gryshyna A, Kautto L, Peterson R, Nevalainen H (2016) On the safety of filamentous fungi with special emphasis on Trichoderma reesei and products made by recombinant means. In: Schmoll M, Dattenbo¨ck C (eds) Gene expression systems in fungi: advancements

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and applications. Fungal biology. Springer, Cham 26. Frisvad JC, Moller LLH, Larsen TO, Kumar R, Arnau J (2018) Safety of the fungal workhorses of industrial biotechnology: update on the mycotoxin and secondary metabolite potential of Aspergillus niger, Aspergillus oryzae, and Trichoderma reesei. Appl Microbiol Biotechnol 102(22):9481–9515 27. Penttil€a M, R€atto¨ M, Knowles J, Nevalainen H, Salminen E (1987) A versatile transformation system for the cellulolytic filamentous fungus Trichoderma reesei. Gene 61(2):155–164 28. Zhong YH, Wang XL, Wang TH, Jiang Q (2007) Agrobacterium-mediated transformation (AMT) of Trichoderma reesei as an efficient tool for random insertional mutagenesis. Appl Microbiol Biotechnol 73(6):1348–1354 29. Hazell BW, Te’o VSJ, Bradner JR, Bergquist PL, Nevalainen KMH (2000) Rapid transformation of high cellulase-producing mutant strains of Trichoderma reesei by microprojectile bombardment. Lett Appl Microbiol 30 (4):282–286 30. Schuster A, Bruno KS, Collett JR, Baker SE, Seiboth B, Kubicek CP, Schmoll M (2012) A versatile toolkit for high throughput functional genomics with Trichoderma reesei. Biotechnol Biofuels 5(1):1 31. Smith JL, Ward M, Bayliss FT (1991) Sequence of the cloned pyr4 gene of Trichoderma reesei and its use as a homologous selectable marker for transformation. Curr Genet 19 (1):27–33 32. Deane EE, Whipps JM, Lynch JM, Peberdy JF (1999) Transformation of Trichoderma reesei with a constitutively expressed heterologous fungal chitinase gene. Enzym Microb Technol 24:419–424 33. Berge`s T, Barreau C (1991) Isolation of uridine auxotrophs from Trichoderma reesei and efficient transformation with the cloned ura3 and ura5 genes. Curr Genet 19(5):359–365 34. Harkki A, Uusitalo J, Bailey M, Penttil€a M, Knowles JKC (1989) A novel fungal expression system: secretion of active calf chymosin from the filamentous fungus Trichoderma Reesei. Nat Biotechnol 7(6):596–603 35. Shoemaker S, Schweickart V, Ladner M, Gelfand D, Kwok S, Myambo K, Innis M (1983) Molecular cloning of exo-cellobiohydrolase I derived from Trichoderma Reesei strain L27. Nat Biotechnol 1 (8):691 36. Teeri T, Salovuori I, Knowles J (1983) The molecular cloning of the major cellulase gene from Trichoderma Reesei. Nat Biotechnol 1 (8):696

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37. Prasun K, Mukherjee BAH, Singh US, Mukherjee M, Schmoll M (eds) (2013) Trichoderma: biology and applications, vol 28, issue 6. Ringgold, Beaverton 38. Guangtao Z, Hartl L, Schuster A, Polak S, Schmoll M, Wang T, Seidl V, Seiboth B (2009) Gene targeting in a nonhomologous end joining deficient Hypocrea jecorina. J Biotechnol 139(2):146–151 39. Steiger MG, Vitikainen M, Uskonen P, Brunner K, Adam G, Pakula T, Penttil€a M, Saloheimo M, Mach RL, Mach-Aigner AR (2011) Transformation system for Hypocrea jecorina (Trichoderma reesei) that favors homologous integration and employs reusable bidirectionally selectable markers. Appl Environ Microbiol 77(1):114–121 40. Ouedraogo J, Arentshorst M, Nikolaev I, Barends S, Ram A (2015) I- SceI-mediated double-strand DNA breaks stimulate efficient gene targeting in the industrial fungus Trichoderma reesei. Appl Microbiol Biotechnol 99 (23):10083–10095 41. Liu R, Chen L, Jiang Y, Zhou Z, Zou G (2015) Efficient genome editing in filamentous fungus Trichoderma reesei using the CRISPR/Cas9 system. Cell Discov 1:15007 42. Liu P, Wang W, Wei D (2017) Use of transcription activator-like effector for efficient gene modification and transcription in the filamentous fungus Trichoderma reesei. J Industrial Microbiol Biotechnol 44(9):1367–1373 43. Nyysso¨nen E, Penttil€a M, Harkki A, Saloheimo A, Knowles J, Ker€anen S (1993) Efficient production of antibody fragments by the filamentous fungus Trichoderma reesei. Bio/Technol 11(5):591 44. Ker€anen S, Penttil€a M (1995) Production of recombinant proteins in the filamentous fungus Trichoderma reesei. Curr Opin Biotechnol 6(5):534–537 45. Nyyssonen E, Keranen S (1995) Multiple roles of the cellulase CBHI in enhancing production of fusion antibodies by the filamentous fungus Trichoderma reesei. Curr Genet 28(1):71–79 46. Sun A, Peterson R, Te’o J, Nevalainen H (2016) Expression of the mammalian peptide hormone obestatin in Trichoderma reesei. New Biotechnol 33(1):99–106 47. Joutsjoki V, Torkkeli T, Helena Nevalainen K (1993) Transformation of Trichoderma reesei with the Hormoconis resinae glucoamylase P (gamP) gene: production of a heterologous glucoamylase by Trichoderma reesei. Curr Genet 24(3):223 48. Joutsjoki VV, Kuittinen M, Torkkeli TK, Suominen PL (1993) Secretion of the Hormoconis

resinae glucoamylase P enzyme from Trichoderma reesei directed by the natural and the cbh1 gene secretion signal. FEMS Microbiol Lett 112(3):281–286 49. Piddington CS, Houston CS, Paloheimo M, Cantrell M, Miettinen-Oinonen A, Nevalainen H, Rambosek J (1993) The cloning and sequencing of the genes encoding phytase (phy) and pH 2.5-optimum acid phosphatase (aph) from Aspergillus niger var. awamori. Gene 133(1):55–62 50. Schmoll M, Seibel C, Kotlowski C, Kubicek CP, Wo¨llert Genannt Vendt F, Liebmann B (2010) Recombinant production of an Aspergillus nidulans class i hydrophobin (DewA) in Hypocrea jecorina (Trichoderma reesei) is promoter-dependent. Appl Microbiol Biotechnol 88(1):95–103 51. Landowski CP, Huuskonen A, Wahl R, Westerholm-Parvinen A, Kanerva A, H€anninen A-L, Salovuori N, Penttil€a M, Natunen J, Ostermeier C, Helk B, Saarinen J, Saloheimo M (2015) Enabling low cost biopharmaceuticals: a systematic approach to delete proteases from a well-known protein production host Trichoderma reesei. PLoS One 10(8):1 52. Landowski CP, Mustalahti E, Sivasiddarthan D, Westerholm-Parvinen A, Saloheimo M, Wahl R, Croute L, Sommer B, Ostermeier C, Helk B, Saarinen J (2016) Enabling low cost biopharmaceuticals: high level interferon alpha-2b production in Trichoderma reesei. Microb Cell Factories 15(1):104 53. Linder MB, Qiao M, Laumen F, Selber K, Hyytia T, Nakari-Setala T, Penttil€a ME (2004) Efficient purification of recombinant proteins using hydrophobins as tags in surfactant-based two-phase systems. Biochemistry 43:11873–11882 54. Mustalahti E, Saloheimo M, Joensuu JJ (2013) Intracellular protein production in Trichoderma reesei (Hypocrea jecorina) with hydrophobin fusion technology. New Biotechnol 30 (2):262–268 55. Smith W, J€antti J, Oja M, Saloheimo M (2014) Comparison of intracellular and secretionbased strategies for production of human α-galactosidase A in the filamentous fungus Trichoderma reesei. BMC Biotechnol 14:91 56. The U.S. biobased economy. https://www.bio. org/toolkit/infographics/us-biobased-econ omy-economic-impact 57. Energy USDo (2017) U.S. Energy and Employment Report (USEER) 2017. BW Research Partnership. https://www.energy. gov/downloads/2017-us-energy-and-employ ment-report

Industrial Relevance of T. reesei 58. Golden JS, Handfield RB, Daystar J, Morrison B, McConnell TE (2019) An economic impact analysis of the US biobased product industry. USDA, A Joint Publication of the Duke Center for Sustainability & Commerce and the Supply Chain Resource Cooperative at North Carolina State University, SWWashington 59. Urbanchuck J (2017) Contribution of the ethanol industry to the economy of the United States in 2016. ABF Economics, Doylestown, PA 60. MarketLine (2017) Industry profile 2017. Global Biotechnology, London 61. Market Research F (2018) Global industrial enzymes markets, 7th edn Marketreasearch. com 62. Seiboth B, Pakdaman BS, Hartl L, Kubicek CP (2007) Lactose metabolism in filamentous fungi: how to deal with an unknown substrate. Fungal Biol Rev 21(1):42–48 63. Mach RL, Zeilinger S (2003) Regulation of gene expression in industrial fungi: Trichoderma. Appl Microbiol Biotechnol 60 (5):515–522 64. (AMFEP) Eiaomafoep (2015) List of commercial enzymes for food, feed and technical applications. https://amfep.org/_library/_files/ Amfep_List_of_Enzymes_update_May_2015. pdf 65. Rychen G, Aquilina G, Azimonti G, Bampidis V, Bastos ML, Bories G, Chesson A, Flachowsky G, Gropp J, Kolar B, Kouba M, Lo´pez-Alonso M, Lo´pez Puente S, Mantovani A, Mayo B, Ramos F, Saarela M, Villa RE, Wallace RJ, Wester P, Brantom P, Dierick NA, Herman L, Glandorf B, K€arenlampi S, Aguilera J, Anguita M, Cocconcelli PS (2018) Safety and efficacy of muramidase from Trichoderma reesei DSM 32338 as a feed additive for chickens for fattening and minor poultry species. EFSA J 16(7):5342 66. Raveendran S, Parameswaran B, Ummalyma SB, Abraham A, Mathew AK, Madhavan A, Rebello S, Pandey A (2018) Applications of microbial enzymes in food industry. Food Technol Biotechnol 56(1):16–30 67. Inflationdata.com (2019) Historical crude oil prices. https://inflationdata.com/articles/ inflation-adjusted-prices/historical-crude-oilprices-table/

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68. Allen F, Andreotti R, Eveleigh DE, Nystrom J (2009) Mary Elizabeth Hickox Mandels, 90, bioenergy leader. Biotechnol Biofuels 2:22 69. Gruber F, Kubicek CP, Visser J, de Graaff LH (1990) Cloning of the Trichoderma reesei pyrG gene and its use as a homologous marker for a high-frequency transformation system. Curr Genet 18(5):447–451 70. Efsa Panel on Food Contact Materials EaPA, Vittorio S, Jose´ Manuel Barat B, Claudia B, Beat Johannes B, Pier Sandro C, Riccardo C, David Michael G, Konrad G, Evgenia L, Alicja M, Gilles R, Inger-Lise S, Christina T, Henk Van L, Laurence V, Holger Z, Boet G, Francesca M, Andre´ P, Jaime A, Margarita AG, Magdalena A, Davide A, Ana G, Nata´lia K, Yi L, Andrew C (2018) Safety evaluation of the food enzyme endo-1,4-β-xylanase from a genetically modified Trichoderma reesei (strain DP-Nzd22). EFSA J 16(11):5479 71. Efsa Panel on Food Contact Materials EaPA, Vittorio S, Jose´ Manuel Barat B, Claudia B, Beat Johannes B, Pier Sandro C, Riccardo C, David Michael G, Konrad G, Evgenia L, Alicja M, Gilles R, Inger-Lise S, Christina T, Henk Van L, Laurence V, Holger Z, Boet G, Lieve H, Jaime A, Andrew C (2019) Safety evaluation of the food enzyme α,α-trehalase glucohydrolase from Trichoderma reesei (strain DP-Nzs51). EFSA J 17(5):5768 72. Silano V, Barat Baviera JM, Bolognesi C, Bru¨schweiler BJ, Cocconcelli PS, Crebelli R, Gott DM, Grob K, Lampi E, Mortensen A, Rivie`re G, Steffensen IL, Tlustos C, Van Loveren H, Vernis L, Holger Z, Jany KD, ˇ eljezˇic D (2019) Glandorf B, Penninks A, Z Safety evaluation of the food enzyme alphaamylase from a genetically modified Trichoderma reesei (strain DP-Nzb48). EFSA J 17 (1):5553 73. Silano V, Barat Baviera JM, Bolognesi C, Bru¨schweiler BJ, Cocconcelli PS, Crebelli R, Gott DM, Grob K, Lampi E, Mortensen A, Riviere G, Steffensen IL, Tlustos C, Van Loveren H, Vernis L, Zorn H, Glandorf B, Marcon F, Penninks A, Smith A (2019) Safety evaluation of the food enzyme lysophospholipase from Trichoderma reesei (strain RF7206). EFSA J 17(1):5548

Chapter 3 The Potential of Synthetic Biology for Trichoderma reesei Roland Martzy and Astrid R. Mach-Aigner Abstract Within the last 20 years, ground-breaking progress has been made in the field of synthetic biology, enabling the construction of novel pathways up to entire synthetic genomes in both prokaryotic and eukaryotic organisms. These innovations are primarily adapted for biotechnological applications, where filamentous fungi such as Trichoderma reesei are widely used to produce various enzymes of industrial interest. In the following chapter, we provide a broad overview on the current progress involving this particular organism, covering studies on synthetic promoters and transcription factors as well as synthetic expression platforms. Furthermore, this chapters aims to be a short introduction to the present book since many methods mentioned here are described in detail in the subsequent chapters. Key words Synthetic biology, Filamentous fungi, Tools for genetic engineering, Transcription factors, Gene expression, Whole-cell biocatalysis

1 Filamentous Fungi Are Comparatively Uncharted Territory in the Field of Synthetic Biology The first use of the term synthetic biology goes back to a publication by the French biologist Ste´phane Leduc [1]. Since then, important breakthroughs such as molecular cloning, genetic engineering, or DNA sequencing have been achieved [2], all of which have been assigned to the areas of molecular biology or biotechnology. It was only in the early 2000s that synthetic biology became popular when the first synthetic circuits and entire synthetic pathways were created in E. coli [3, 4]. Within the last 20 years, this field of research has witnessed not only ground-breaking progress in academic research, but also the rise of countless companies with an estimated global market net worth of 3.9 billion dollars in 2016 [5]. Furthermore, the International Genetically Engineered Machine (iGEM) competition annually attracts teams of students from all over the world to develop new ideas and projects based on a collection of genetic building blocks (BioBricks) [6]. For these

Astrid R. Mach-Aigner and Roland Martzy (eds.), Trichoderma reesei: Methods and Protocols, Methods in Molecular Biology, vol. 2234, https://doi.org/10.1007/978-1-0716-1048-0_3, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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reasons, authors have recently suggested that synthetic biology will be a major driving force in the next industrial revolution [7]. In fungal synthetic biology, the most extensively studied organisms are yeasts. Within this group, a large number of publications concentrate on the baker’s yeast Saccharomyces cerevisiae [8]. These studies comprise the construction of synthetic promoters and transcription factors (TFs) [9], or chimeric pathways [10] right up to the design of a synthetic yeast genome [11]. Although filamentous fungi play an essential role in the industry-scale production of primary and secondary metabolites, comparatively few studies directly relate to synthetic biology, a considerable part of which address Aspergillus species [12–15] next to Trichoderma reesei. In the following, we outline the advances that have been made for T. reesei in terms of molecular tools that can be used for synthetic biology applications. Some of the methods mentioned in this chapter can also be found as protocols in the present book. In addition, we provide some examples for engineering synthetic promoters, TFs, and expression platforms or for the use of T. reesei as a whole-cell biocatalyst that expresses a synthetic enzyme cascade.

2 A Variety of Genetic Engineering and Gene Editing Tools Have Been Developed and Adapted for Trichoderma reesei In order to introduce foreign DNA into T. reesei, different transformation methods have been established in recent decades, such as polyethylene glycol-mediated protoplast transformation [16], biolistic transformation [17], or electroporation [18]. Additionally, genes involved in the non-homologous end joining mechanism were deleted in the strains QM6a and TU6, which increased the site-directed integration rates based on homologous recombination up to 100% [18, 19]. Proceeding from these strains, a reusable bidirectionally selectable marker system [19] and several auxotrophy/prototrophy-based marker genes were used to improve and facilitate strain engineering [20]. However, these approaches are limited when targeting essential genes of which the deletion leads to pleiotropic effects. Besides, knocking out a variety of genes to study their function is still a tedious procedure. In order to overcome some of these problems, several studies have successfully demonstrated that the RNA interference (RNAi) system can be used to influence transcript levels in T. reesei [21, 22]. In 2018, Wang and coworkers reported a copper-controlled RNAimediated knockdown system that can be applied for studying genes that are subject to stringent regulation [23]. The advent of programmable nucleases eventually paved the way for sequence-specific cleavage of DNA, enabling a new way of gene editing. This was demonstrated for the first time with zinc

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finger proteins that contain the cleavage domain of an endonuclease, which formed the basis for zinc finger nucleases (ZFNs) [24]. Further progress in this area has been made through the development of transcription-activator-like effector nucleases (TALENs) [25]. However, all of these approaches were quickly surpassed by systems based on RNA-guided endonucleases, such as CRISPR/Cas9 [26]. In 2015, the CRISPR/Cas9 system was adapted for the first time for filamentous fungi using T. reesei as a model organism [27]. Since then, it has been widely applied to a large number of other filamentous fungi [28]. Although genome editing is nowadays dominated by the CRISPR/Cas9 system, the TALEN-based system was also recently established in T. reesei [29]. Furthermore, some applications using ZFNs have been shown, e.g., to increase cellulase production in strain Rut-C30 [30] or to identify new transcriptional repressors that are involved in the expression of cellulases [31]. Another essential component of synthetic biology is wellcharacterized promoters for the controlled expression of target genes. As regards this subject, in 2014, Wang and colleagues described the construction of a promoter collection to co-express two alkaline cellulases [32]. Furthermore, a recent review addressed promoters frequently used for recombinant gene expression in T. reesei and evaluated common structural features as well as the potential of a promoter toolbox to build up synthetic circuits and expression systems [33]. In two studies, promoters were described that are responsive to copper [34] or L-methionine [35], both of which act as a repressing agent and thus suppress the expression of the target gene. This can be used to create reversible gene knockouts for switching the system on and off in a controlled manner, e.g., in combination with the RNAi system mentioned above [23].

3 Substantial Effort Has Been Made Toward Engineering of Gene Expression in Trichoderma reesei A lot of research has been done in the past 25 years with respect to engineering regulatory proteins that are involved in the cellulase expression system of the fungus. One of the first proteins studied in more detail was Cre1, a transcriptional regulator that plays a crucial role in carbon catabolite repression (CCR). In 2002, Cziferszky and coworkers described the Ser241 residue in Cre1 as a target for phosphorylation, a modification that is essential for the protein to bind to the corresponding site on the DNA [36]. Unexpectedly, removing the phosphorylation site by mutating the serine to an alanine or glutamate resulted in Cre1 being permanently bound to its DNA-binding site, causing a steadily sustained CCR. To overcome this, the authors mutated the Glu244 residue within the casein

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kinase consensus sequence to a valine, resulting in a complete loss of Cre1 phosphorylation and thereby rendering the mutant strain carbon catabolite derepressed. Newer studies have also gained insight into how cellulase expression responds to a total lack of Cre1 and a truncated variant (Cre1–96), respectively [37, 38]. Mello-de-Sousa and coworkers found that Cre1–96 is still able to bind to its target site on the DNA. However, instead of acting as a repressor, it exerted an activating influence on the expression of target genes in strain Rut-C30, which is potentially attributed to a Cre1–96-mediated repositioning of nucleosomes [37]. Using a similar way of engineering, Derntl and colleagues introduced a point mutation in the transactivator Xyr1 by replacing an alanine with a valine on residue 824. This single amino acid exchange causes a glucose blind phenotype in industrial T. reesei strains, which leads to a greatly increased basal level of cellulase expression as well as a strongly deregulated expression of xylanases. Although Xyr1 is one of the main regulators of genes encoding for plant cell wall degrading enzymes (PCWDE), the mere overexpression of Xyr1 did not lead to an overall enhancement in PCWDE activity [39]. In 2017, Zhang and colleagues created a synthetic transcriptional activator by combining the DNA-binding domain of Cre1 with the DNA-binding domain and the putative effector domain of Xyr1. The Rut-C30 strain overexpressing this chimeric TF was no longer susceptible to CCR and showed an almost 13-fold increase in cellulase production on glucose compared to the Rut-C30 control strain [40]. In a recent study, the DNA-binding domain of Xyr1 was fused to the transactivating domain of two regulators of the sorbicillinoid gene cluster, Ypr1 and Ypr2, respectively, in order to learn more on the regulatory function of all involved Gal4-like TFs. In summary, it can be said that one of the fusions TF enabled an almost carbon sourceindependent induction of the main PCWDEs, which was demonstrated by the production of xylanases on glycerol [41]. As already stated, promoters are other crucial components for the regulation of gene expression. The cbh1 and xyn1 promoters are most commonly used for the expression of heterologous genes. Both the promoters harbor multiple binding sites for the main transactivator Xyr1 (XBS), i.e., the consensus sequence 50 -GGC (A/T)3-30 arranged in tandem or as inverted repeats, respectively [42, 43]. In a study from 2017, Kiesenhofer and colleagues investigated the impact of these cis elements on the strength of the promoters and their inducibility by different carbon sources. For this purpose, the authors rearranged the XBS and fused the artificial promoters to the goxA reporter gene from A. niger. The authors found that the configuration of the cis elements and their distance from the transcription start site had a greater influence on the promoter strength than the sheer number of XBS. Remarkably, when a certain cis element bearing an inverted repeat of the XBS

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was introduced into the cbh1 promoter, it did not only become stronger, but it was also inducible by xylan and wheat straw [43]. Similarly, Sun and coworkers replaced eight binding sites of the transcriptional repressor Ace1 in the cbh1 promoter with those of transactivators, such as Ace2, Xyr1, and Hap2/3/5. While the additional Hap2/3/5 binding sites completely abolished the transcription of the reporter gene DsRed, the Ace2 and Xyr1 DNA-binding sites increased the expression of DsRed and the secretion of a heterologously expressed mannanase from A. niger up to 5.9- and 5-fold, respectively [44]. Given these examples, TFs as well as promoters have been shown to be promising engineering targets to further develop industrial production strains. However, other interesting approaches to control gene expression in T. reesei are currently being investigated, for example, a system based on a synthetic transactivator that is induced by blue light [45]. Furthermore, the recent characterization of a long non-coding RNA that is involved in the regulation of cellulase-encoding genes in T. reesei might be a first step in the development of synthetic RNA molecules to improve gene expression [46].

4 Trichoderma reesei Can be Used as a Microbial Cell Factory to Produce Food Additives or Drug Precursors Owing to its saprophytic lifestyle, T. reesei is able to grow on renewable lignocellulosic biomass as a substrate [47]. For this reason, the fungus can be an attractive whole-cell biocatalyst to produce industrially relevant compounds in a more sustainable way. This was demonstrated by Jovanovic and colleagues, who characterized the gene coding for the erythrose reductase (Err1) in T. reesei, an enzyme that is responsible for the final step in the synthesis of the sugar alcohol erythritol [48]. This low-caloric substitute to common sugars in foods is currently produced using osmophilic yeasts; however, the glucose solution that is applied as a substrate makes this an (socio)economically irrational process. Therefore, the authors used pre-treated wheat straw as the sole carbon source and overexpressed the err1 gene under different native promoters, which led to an increased formation of erythritol on this non-food substrate [49]. In further consequence, employing T. reesei as a host cell factory might help render the production of this alternative sweetener more sustainable compared to previous approaches [49]. Due to its chitinolytic enzyme system, T. reesei not only is able to use cellulosic biomass as a substrate but can also utilize chitin as both a sole carbon and nitrogen source [50]. The wild-type strain thereby almost exclusively degrades the biopolymer to monomeric

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units of N-acetylglucosamine (NAG) [51]. Another example for T. reesei serving as a whole-cell biocatalyst is shown in a study by Steiger and coworkers, who introduced codon optimized variants of the NAG 2-epimerase gene from Anabaena sp. CH1 and the N-acetylneuraminic acid (NANA) synthase gene from Campylobacter jejuni into the genome of T. reesei. The resulting enzymatic cascade enables the fungus to convert the monomeric NAG to N-acetylmannosamine and subsequently, to NANA, which is an important precursor for antiviral drugs, such as Relenza [52]. As summarized in the study, traditional methods of preparing NANA are either too expensive or make use of unsustainable resources and processes. Therefore, the approach using T. reesei again highlights how widely unused biopolymers can be utilized efficiently as substrates for sustainable production of high value compounds.

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What Other Aces Does the Fungus Have Up Its Hyphae? For the most part, studies on T. reesei still are focused on optimizing strains in terms of improved cellulase production. In recent years, however, researchers have increasingly tried to exploit the high secretory capacity of this fungus for the expression of heterologous proteins [53]. The main components for this approach have so far involved the strong, inducible cbh1 promoter and a carrier peptide serving as secretory signal fused to the heterologous protein. However, this strategy most often resulted in unsatisfactory yields or a high background of co-secreted enzymes, which unnecessarily complicates the subsequent downstream processing [54]. For these reasons, a synthetic gene expression system (SES) for a universal use in fungal hosts was recently developed, which is based on a combination of two expression cassettes and synthetic TFs with corresponding promoters [55]. Its successful application in T. reesei was subsequently demonstrated by Rantasalo and colleagues, who applied the SES to produce highly enriched lipase B of Candida antarctica (CalB). The authors were able to eliminate unwanted background enzymes and at the same time obtained fully functional CalB protein at a concentration of 4 g/l [56]. Furthermore, Jorgensen and coworkers developed an expression platform that enables the easy combination of genetic tools in order to facilitate the high-throughput construction of T. reesei strains for large expression studies [57]. In light of these novel developments, this fungus might become an attractive alternative host for the production of a variety of proteins in the future, eventually replacing some of the widely used yeast expression systems.

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Chapter 4 Resistance Marker- and Gene Gun-Mediated Transformation of Trichoderma reesei Monika Schmoll and Susanne Zeilinger Abstract Transformation enables the transfer of DNA into fungal cells for subsequent integration into the genome. Due to its versatility in industrial application, transformation is of utmost importance in Trichoderma reesei and hence continuously optimized. As one of the most crucial obstacles in fungal transformation efforts, removal of the cell wall is required to efficiently target genome modification cassettes to the genome. Here we describe resistance marker-mediated gene gun (biolistic) transformation of fungal spores of T. reesei as an alternative to protoplast transformation. Key words Trichoderma reesei, Resistance marker, Hygromycin, Biolistic transformation, Gene gun, Particle bombardment

1

Introduction Understanding and hence exploiting the diverse biosynthetic capabilities of fungi—from secondary metabolites to enzymes and performance proteins—requires the possibility to manipulate their genomes. Traditionally, two factors challenged the success of genome manipulation efforts: crossing the fungal cell wall and overcoming efficient genome defense systems. Additionally, the selection of appropriate selection marker systems is of importance. Removal of the fungal cell wall for transformation is mostly done by protoplasting, which requires application of carbohydrate degrading enzymes with optimized functionalities and time spans for every individual species [1]. However, also alternative methods for transformation, which do not require protoplasting, have been explored and are still in use, like biolistic transformation (see also below), electroporation, or Agrobacterium-mediated transformation (for an overview see ref. 2). For T. reesei, transformation was first reported with the selection markers amdS and argB of Aspergillus nidulans [3]. Strains with uridine auxotrophy were later on complemented with the pyr4

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gene as selection marker [4]. Transformation using the hph (hygromycin B phosphotransferase) gene as selection marker followed soon [5]. To date, these selection systems are still most frequently used, although additional selection markers, such as phleomycin [6], geneticin [7], nourseothricin [8], or growth on mannitol [9] have also been applied in Trichoderma spp. [10]. On average, 600–800 transformants per microgram of DNA are reported for the most common methods [3–5]. Despite the efficiency of transformation, a major problem with strain construction is targeted integration of the construct of interest. For example, only about 2–5% of obtained transformants of a nonessential gene to be deleted indeed show homologous integration and hence successful deletion. As this is mainly due to nonhomologous end joining (NHEJ; [11]), also in T. reesei mutants were prepared lacking this mechanisms and showing increased efficiency of homologous integration by the deletion of tku70 [12], tmus53 [13], or tku80 [14]. Thereby, for some genes, efficiencies of more than 90% of homologous integration could be achieved. The downside of this improvement is that strains lacking NHEJ show decreased genetic stability and genome integrity and hence propagation of strains bearing such mutations has to be kept at a minimum to prevent unspecific mutations causing unrelated phenotypes [11]. Additionally, also some physiological alterations and modulations of gene regulation were reported. Increased ratios of homologous integration can also be achieved by the application of the split marker system in which selection is only successful upon appropriate integration of both parts of the marker cassette [15, 16]. The drawbacks connected with the NHEJ pathway and its elimination can in part be alleviated by using the CRISPR Cas9 system [17]. The CRISPR Cas9 system was also adapted for the application in T. reesei [18, 19]. This system allows for highefficiency gene targeting and genome editing at very specific sites, although selection of target sequences and integration motifs must be done carefully to avoid off-target effects. In recent years, large-scale efforts for the preparation of wholegenome knockout libraries were initiated for Neurospora crassa [20] and Aspergillus nidulans [21, 22]. For T. reesei, such an initiative was not started, but in preparation for high-throughput strain construction, an appropriate workflow including a yeast recombinant cloning procedure and primer design for all annotated gene models for three selection markers (hph, pyr4, and amdS) was established and tested [23]. In contrast to the first developments of fungal transformation systems, which mainly involved protoplast preparation, biolistic transformation relies on the penetration of the cell wall by particle bombardment. This approach, also known as “gene gun technique,” has been developed for the delivery of RNA or DNA into

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plant cells [24] and then was as well successfully adapted for the transformation of fungal spores or hyphae [25]. Successful biolistic transformation of intact conidia was reported for several Trichoderma species such as T. reesei, T. atroviride (former T. harzianum), and T. virens (former Gliocladium virens) [26–28] illustrating one of the main advantages of this approach: the use of conidia as target cells, thereby circumventing the preparation of protoplasts (which are practically always multi-nucleate necessitating several rounds of single spore isolation for getting mitotically stable transformants) and allowing the transformation of species from which protoplasts are hard to prepare. In addition, for T. atroviride and T. virens [26– 28], a direct comparison with the protoplast-mediated approach revealed higher transformation rates and genetic stability of the transformants [27]. Nevertheless, due to the expensive equipment needed—a special particle delivery device is required that under vacuum accelerates DNA-coated particles at high velocity onto the target cells—biolistic transformation is not very commonly used in fungi that are amenable to other methods. The same is true for T. reesei, for which gene gun transformation is of lower relevance for genome manipulation than protoplast-mediated transformation or other alternative methods like electroporation and Agrobacterium-mediated transformation.

2

Materials 1. The PDS-1000/He System (Bio-Rad) connected to a vacuum source and a tank of high-pressure (2400–2600), high-purity helium. 2. Microcarriers: Gold (0.6 μm) or Tungsten M-10 (0.7 μm) particles (see Note 1). Weigh 10 mg of the microparticles into a clean 1.5 mL reaction tube, wash them three times with 1 mL 70% ethanol, two times with 1 mL of sterile distilled water, resuspend the particles in 200 μL of sterile distilled water, and store them as 50 μL aliquots (see Note 2). 3. Macrocarriers and macrocarrier holders: Insert macrocarriers into macrocarrier holder and sterilize by autoclaving. 4. Rupture disks (650 psi). 5. Stopping screens: Sterilize by autoclaving or soaking in 70% (v/v) ethanol. 6. Conidia-containing agar plates. 7. NaCl–Tween solution: 0.8% (w/v) NaCl, 0.05% (v/v) Tween 80. For 5 mL, dissolve 0.4 g of NaCl and 25 μL of Tween 80 in distilled water, filter sterilize and store at room temperature. 8. Funnels (e.g., reaction tubes) filled with glass wool.

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9. Bottom medium for hygromycin as selection marker: 3% (w/v) malt extract and 2% (w/v) agar-agar or potato dextrose agar (PDA), selection reagent. For 400 mL (for 30–40 plates), dissolve 12 g of malt extract and 8 g agar-agar in water, adjust to 400 mL, and autoclave. Add 50–100 μg/mL of hygromycin (see Note 3). 10. Overlay medium for hygromycin as selection marker: 3% (w/v) malt extract and 2% (w/v) agarose or PDA. Prepare and add hygromycin as selection reagent as described above. 11. 0.1 M Spermidine (for molecular biology,
99.5%). 12. 2.5 M CaCl2. 13. Recombinant DNA: 1 μg/mL (circular or linear). 14. Absolute ethanol. 15. 70% (v/v) ethanol.

3

Methods Work under sterile conditions and perform all steps at room temperature unless otherwise specified.

3.1 Preparation of Bombardment Plates

1. Harvest fungal conidia from a freshly sporulated culture plate by adding NaCl–Tween solution onto the plate and removing conidia with a sterile spreader. Remove mycelial fragments by filtering the conidial solution through a funnel filled with glass wool. Spread 1  107 conidia in the center of agar plates containing an appropriate bottom selection medium. Use conidia-containing plates within 3 h for particle bombardment.

3.2 Coating Washed Microcarriers with DNA

1. Take a 50 μL aliquot of microcarrier particles and vortex rigorously. 2. Add 5 μL of DNA (1 μg/μL), 50 μL of 2.5 M CaCl2, and 20 μL of 0.1 M spermidine in the given order and vigorously vortex after each addition for at least 30 s. 3. Incubate on ice for at least 10 min. 4. Pellet the DNA-coated particles in a microfuge for 10 s and remove the supernatant. 5. Wash the particles with 500 μL of absolute ethanol by gently resuspending the particles. 6. After spinning down the particles, discard the supernatant. 7. Resuspend the DNA-coated particles in 50 μL absolute ethanol. 8. With a pipet tip, evenly load 10 μL of the DNA-coated particles onto the central 1 cm of each prepared macrocarrier and allow

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to dry (see Notes 4 and 5). Use loaded macrocarriers within 2 h. 9. Set up a negative control by processing the particles as described but with no DNA. 3.3 Preparation of the Gene Gun and Performing a Bombardment

1. Turn on the vacuum pump, the helium gas, and the PDS-1000/He System (see Notes 6 and 7). 2. Sterilize the chamber of the PDS-1000/He System by spraying or wiping with 70% ethanol. 3. Sterilize a rupture disk (650 psi) by dipping it in absolute ethanol, insert the disk into the rupture disc holder with sterile forceps, and fix it at the top of the bombardment chamber at the end of the gas acceleration tube (see Note 8). 4. Assemble macrocarrier and stopping screen and place the macrocarrier launch assembly DNA side down in the top slot inside the bombardment chamber (see Note 9). 5. Remove the lid of a conidia-containing agar plate and place the plate on the target shelf into the bombardment chamber at a target distance of either 3 or 6 cm (see Note 10). Close the bombardment chamber. 6. Evacuate the chamber to the desired level (about 2800 Hg) by pressing and holding the vacuum switch. 7. When ready, set the vacuum switch to the HOLD position. Then press and hold the fire switch (see Note 11). The helium pressure will build inside the gas acceleration tube until the rupture disk bursts, indicated by a gentle popping sound. Immediately after disk rupture, release the fire switch to avoid wasting helium. 8. Vent the chamber, open chamber door, and remove the bombarded agar plate.

3.4 Selection of Transformants

1. Incubate the plate with the bombarded conidia for 5–6 h at 28  C before overlaying with 4 mL overlay medium containing an appropriate concentration of the selection reagent (see Note 12). Continue incubation at 28  C for 3–4 days until colonies appear. 2. Pick transformants to small selection plates (30 mm) by excising them from the agar of the transformation plate and grow them until sporulation (see Note 13).

4

Notes 1. Gold and tungsten microparticles can be used with the PDS-1000/He System. Gold particles are highly uniform and

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biologically inert, but expensive. Tungsten particles are more irregular in shape, show higher size variation, and are prone to oxidation. 2. Microparticle aliquots may be stored at +4  C for up to 2 weeks. 3. Most selection reagents are temperature sensitive, so prepare stock solutions and add after autoclaving. The inhibitory concentration of the selection reagent should be determined for every strain and new batches of the chemical. For hygromycin, a concentration of 50–100 μg/mL is usually appropriate for T. reesei strains; however, inhibition may vary for different suppliers. 4. To avoid settling of the microparticles, it is important to rigorously vortex the tube before pipetting and to rapidly apply the particles onto the macrocarrier disk. 5. The dried particles should be visible on the center of the macrocarrier disk indicating that a sufficient amount has been loaded. 6. The higher the vacuum, the lower is the frictional deceleration of the particle. Hence, a vacuum of 25–2800 Hg is recommended. 7. The microparticles are accelerated by a helium shock wave generated by burst of the rupture disk. Hence, verify that the pressure regulator of the helium tank is set to 200 psi over the desired disk burst pressure. 8. The rupture disk has to be replaced after each single bombardment as it bursts when firing the gene gun. 9. The stopping screen and macrocarrier have to be replaced after each single bombardment. 10. Microparticle flight distance, in addition to vacuum, microparticle size, gas pressure, cell type, and density, is one of the most important parameters for optimization [28]. Four distances can be adjusted by positioning the target shelf at given levels within the bombardment chamber: level 1 ¼ 3 cm; level 2 ¼ 6 cm; level 3 ¼ 9 cm; level 4 ¼ 12 cm below the stopping screen. 11. The DNA-coated particles released from the macrocarrier by the helium shock wave after rupture disk burst pass through the grid of the stopping screen and penetrate into the conidia on the agar plate. 12. The appropriate concentration of the selection reagent (e.g., antibiotic) should be determined prior to transformation.

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13. Eventually, perform single-spore isolation. In any case, check for positive transformation by PCR of genomic DNA and/or Southern blot analysis. References 1. Rodriguez-Iglesias A, Schmoll M (2015) Protoplast transformation for genome manipulation in fungi. In: Van den Berg MA, Maruthachalam K (eds) Genetic transformation systems in fungi, vol 1. Springer, Cham, pp 21–40 2. van den Berg MA, Maruthachalam K (eds) (2015) Genetic transformation systems in fungi, vol I. Springer, Heidelberg, (ISBN 978-3-319-10142-2) 3. Penttil€a M, Nevalainen H, Ratto M, Salminen E, Knowles J (1987) A versatile transformation system for the cellulolytic filamentous fungus Trichoderma reesei. Gene 61 (2):155–164 4. Gruber F, Visser J, Kubicek CP, de Graaff LH (1990) The development of a heterologous transformation system for the cellulolytic fungus Trichoderma reesei based on a pyrG-negative mutant strain. Curr Genet 18(1):71–76 5. Mach RL, Schindler M, Kubicek CP (1994) Transformation of Trichoderma reesei based on hygromycin B resistance using homologous expression signals. Curr Genet 25(6):567–570 6. Malmierca MG, Cardoza RE, Alexander NJ, McCormick SP, Collado IG, Hermosa R, Monte E, Gutierrez S (2013) Relevance of trichothecenes in fungal physiology: disruption of tri5 in Trichoderma arundinaceum. Fungal Genet Biol 53:22–33 7. Gruber F, Bicker W, Oskolkova OV, Tschachler E, Bochkov VN (2012) A simplified procedure for semi-targeted lipidomic analysis of oxidized phosphatidylcholines induced by UVA irradiation. J Lipid Res 53(6):1232–1242 8. Atanasova L, Gruber S, Lichius A, Radebner T, Abendstein L, Munsterkotter M, StralisPavese N, Labaj PP, Kreil DP, Zeilinger S (2018) The Gpr1-regulated Sur7 family protein Sfp2 is required for hyphal growth and cell wall stability in the mycoparasite Trichoderma atroviride. Sci Rep 8(1):12064 9. Guangtao Z, Seiboth B, Wen C, Yaohua Z, Xian L, Wang T (2010) A novel carbon source-dependent genetic transformation system for the versatile cell factory Hypocrea jecorina (anamorph Trichoderma reesei). FEMS Microbiol Lett 303(1):26–32 10. Malmierca MG, Cardoza RE, Gutierrez S (2015) Trichoderma transformation methods.

In: Van den Berg MA, Maruthachalam K (eds) Genetic transformation systems in fungi, Fungal biology, vol I. Springer, Cham, pp 41–48 11. Krappmann S (2007) Gene targeting in filamentous fungi: the benefits of impaired repair. Fungal Biol Rev 21(1):25–29 12. Guangtao Z, Hartl L, Schuster A, Polak S, Schmoll M, Wang T, Seidl V, Seiboth B (2009) Gene targeting in a nonhomologous end joining deficient Hypocrea jecorina. J Biotechnol 139(2):146–151 13. Steiger MG, Vitikainen M, Uskonen P, Brunner K, Adam G, Pakula T, Penttila M, Saloheimo M, Mach RL, Mach-Aigner AR (2011) Transformation system for Hypocrea jecorina (Trichoderma reesei) that favors homologous integration and employs reusable bidirectionally selectable markers. Appl Environ Microbiol 77(1):114–121 14. Stappler E, Dattenbo¨ck C, Tisch D, Schmoll M (2017) Analysis of light- and carbon-specific transcriptomes implicates a class of G-proteincoupled receptors in cellulose sensing. mSphere 2(3):e00089-00017 15. Derntl C, Kiesenhofer DP, Mach RL, MachAigner AR (2015) Novel strategies for genomic manipulation of Trichoderma reesei with the purpose of strain engineering. Appl Environ Microbiol 81(18):6314–6323 16. Gravelat FN, Askew DS, Sheppard DC (2012) Targeted gene deletion in Aspergillus fumigatus using the hygromycin-resistance splitmarker approach. Methods Mol Biol 845:119–130 17. Donohoue PD, Barrangou R, May AP (2018) Advances in industrial biotechnology using CRISPR-Cas systems. Trends Biotechnol 36 (2):134–146 18. Liu R, Chen L, Jiang Y, Zhou Z, Zou G (2015) Efficient genome editing in filamentous fungus Trichoderma reesei using the CRISPR/Cas9 system. Cell Discov 1:15007 19. Rantasalo A, Vitikainen M, Paasikallio T, Jantti J, Landowski CP, Mojzita D (2019) Novel genetic tools that enable highly pure protein production in Trichoderma reesei. Sci Rep 9(1):5032 20. Colot HV, Park G, Turner GE, Ringelberg C, Crew CM, Litvinkova L, Weiss RL, Borkovich KA, Dunlap JC (2006) A high-throughput

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gene knockout procedure for Neurospora reveals functions for multiple transcription factors. Proc Natl Acad Sci U S A 103 (27):10352–10357 21. De Souza CP, Hashmi SB, Osmani AH, Andrews P, Ringelberg CS, Dunlap JC, Osmani SA (2013) Functional analysis of the Aspergillus nidulans kinome. PLoS One 8(3): e58008 22. Son S, Osmani SA (2009) Analysis of all protein phosphatase genes in Aspergillus nidulans identifies a new mitotic regulator, fcp1. Eukaryot Cell 8(4):573–585. https://doi.org/10. 1128/EC.00346-08 23. Schuster A, Bruno KS, Collett JR, Baker SE, Seiboth B, Kubicek CP, Schmoll M (2012) A versatile toolkit for high throughput functional genomics with Trichoderma reesei. Biotechnol Biofuels 5(1):1 24. Klein RM, Wolf ED, Wu R, Sanford JC (1987) High-velocity microprojectiles for delivering

nucleic acids into living cells. Nature 327:70–73 25. Li D, Tang Y, Lin J, Cai W (2017) Methods for genetic transformation of filamentous fungi. Microb Cell Factories 16(1):168 26. Hazell BW, Te’o VS, Bradner JR, Bergquist PL, Nevalainen KM (2000) Rapid transformation of high cellulase-producing mutant strains of Trichoderma reesei by microprojectile bombardment. Lett Appl Microbiol 30 (4):282–286 27. Lorito M, Hayes CK, Di Pietro A, Harman GE (1993) Biolistic transformation of Trichoderma harzianum and Gliocladium virens using plasmid and genomic DNA. Curr Genet 24 (4):349–356 28. Te’o VS, Bergquist PL, Nevalainen KM (2002) Biolistic transformation of Trichoderma reesei using the Bio-Rad seven barrels hepta adaptor system. J Microbiol Methods 51(3):393–399

Chapter 5 Use of Auxotrophic Markers for Targeted Gene Insertions in Trichoderma reesei Irene Tomico-Cuenca and Christian Derntl Abstract In this chapter, we describe a routinely used strategy for targeted gene insertions in Trichoderma reesei using auxotrophic markers. Generally, targeted gene integrations are advantageous over random, ectopic integration, because the copy number and locus of integration are controlled, abolishing the risk of pleiotropic effects. The use of auxotrophic markers allows a direct, cheap, and easy method for selection. The first step is the construction of recipient strains in a NHEJ-deficient strain. We routinely use deletion strains of pyr4, encoding for the orotidine 50 -phosphate decarboxylase (EC 4.1.1.23) and/or asl1, encoding for the argininosuccinate lyase (EC 4.3.2.1). In the second step, the gene of interest is inserted together with the marker gene. Here we describe the necessary strategy for the construction of the recipient strains and insertion constructs, a PEG-mediated transformation protocol, and a protocol for genetic confirmation of the gene insertion. Key words Transformation, Gene integration, Heterologous expression, Auxotrophic marker, Protoplast, Trichoderma reesei

1

Introduction Trichoderma reesei has been widely used in basic and applied research for its interest in regulatory mechanisms of cellulase and hemicellulase gene expression [1–6] and secretion pathways [7]. For the genetic manipulation of the wild-type strain, several transformation methods have been described, such as PEG-mediated transformation of protoplasts [8], electroporation [9], particle bombardment [10], and Agrobacterium tumefaciensmediated transformation [11]. Regardless the chosen transformation method, markers for the selection of transformants are required. Some examples of most used marker genes are hph (hygromycin B resistance) [12], pyr4 (orotidine 5-phosphate decarboxylase) [13], and amdS (utilization of acetamide) [8]. For gene insertions in T. reesei, a commonly used strategy is co-transformation of two plasmids, one bearing a marker gene

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and the other containing the gene of interest (GOI). This method results in the random integration of the gene of interest in 80% of the cases and in variable copy numbers [8]. This causes different problems. First, integration of the marker gene does not ensure integration of the gene of interest. Second, the random integration can trigger unpredictable locus effects, and third, the copy number is incontrollable. These problems can be solved by site-specific integration, which needs a high efficiency of homologous integration as prerequisite. This is achieved by the deletion of genes participating in the nonhomologous end-joining mechanism, such as tku70 and tmus53 [9, 12]. A strategy for targeted insertions was described using I-SceI-mediated double-strand breaks in T. reesei [14]. The S. cerevisiae I-SceI endonuclease recognizes a specific sequence that can be inserted in the desired place in the genome of T. reesei. Then, T. reesei is transformed with a plasmid encoding for the endonuclease. This method increases the efficiency of transformation and decreases the number of random integrations. However, it does not offer a high versatility regarding the use of auxotrophy and prototrophy regain. In this chapter, we describe a laborsaving transformation strategy, using strains with a complete deletion of pyr4 or a partial deletion of asl1 as recipient strains for the transformation and further prototrophic selection (see Fig. 1). For this purpose, first, the cassettes containing the truncated or null versions of the markers must be constructed. There are plasmids for the here-described gene deletions available [15]. However, the readers can choose to construct new plasmids using a cloning method of their preference. These deletion plasmids are then transformed into T. reesei, resulting in auxotrophic strains (see Fig. 1). There are corresponding deletion strains in a QM6a Δtmus53 background already available [15]. Next, the targeting construct (containing the gene of interest as well as the marker gene to regain prototrophy) is assembled. Plasmids containing multiple cloning sites are available [15]. Alternatively, the readers can choose to construct new plasmids following the scheme in Fig. 1 by a method of their choice. Finally, GOI is inserted in front of the auxotrophic marker genes, which are reconstituted as a result of a successful transformation (see Fig. 1). In case a split marker approach (see Fig. 1a) is followed, a 100% transformation rate can be expected. We recommend verifying the correct and exclusive integration of the GOI at the auxotrophic marker loci using PCR and Southern blot assays.

2 2.1

Materials Transformation

1. Spore-harvesting solution: 0.8% (w/v) NaCl and 0.05% (v/v) Tween-80 dissolved in distilled water. Autoclave.

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Fig. 1 Schematic view of the strategy followed for the design of the split marker system (a) and the complete deletion marker system (b). In the parental strain, the gene conferring prototrophy (green arrow) is partially or completely deleted using a marker gene of choice (blue arrow). The auxotrophic strain (recipient) is used in a further transformation with a cassette containing the gene of interest (yellow arrow) and parts of the gene or the complete gene conferring prototrophy. Gray boxes represent the homologous recombination areas, and black arrows represent the binding sites for the primers used in the PCR analyses for genomic testing

2. MEX medium: Weigh 1 g of peptone and 30 g of malt extract and dissolve in 1 L of tap water. Add required supplements (5 mM uridine and/or 2.5 mM arginine). Add 15 g of agar. Autoclave. 3. Cellophane discs (approximately diameter of 7 cm, six pieces per strain): put between wet pieces of Whatman Filter Paper (approximately 10 cm  10 cm). Autoclave. 4. Lysing enzymes from Trichoderma harzianum.

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5. Solution A: 100 mM KH2PO4 and 1.2 M sorbitol dissolved in distilled water. Adjust pH to 5.6 using 3 M KOH. Autoclave and store at 4  C. 6. Glass wool funnel: Place a piece of glass wool in the neck of a funnel and autoclave it. 7. STC solution: 10 mM Tris–HCl, pH 7.5, 50 mM CaCl2, and 1 M sorbitol dissolved in distilled water. Filter sterile and store at 4  C. 8. 20 μg of linearized plasmid DNA dissolved in 15 μL of sterile water (see Note 1). 9. PEG solution: 25% (w/v) PEG 6000, 10 mM Tris–HCl, pH 7.5, and 50 mM CaCl2 dissolved in distilled water. Some heat might be needed to completely dissolve the PEG 6000. Filter sterile and do not autoclave. 10. 0.1 M phosphate–citrate buffer: Weigh 17.8 g of Na2HPO4 and dissolve it in 500 mL of distilled water. Adjust to a pH of 5.0 using 1 M citric acid. Fill up to 1 L with distilled water. 11. Mineral salt solution: Weigh 5.6 g of (NH4)2SO4, 8.0 g of KH2PO4, 1.2 g of MgSO4, and 1.6 g of CaCl2·2H2O and dissolve in 1 L of distilled water. 12. Trace elements solution: Weigh 250 mg of FeSO4·7H2O, 85 mg of MnSO4·H2O, 70 mg of ZnSO4·7H2O, and 100 mg of CoCl2·2H2O. Dissolve in 800 mL of distilled water. Adjust to a pH of 2.0 using 0.1 M HCl. Fill up to 1 L with distilled water. 13. Selection medium (100 mL per transformation): Weigh 10 g of glucose and 182.17 g of sorbitol. Add 500 mL of 0.1 M phosphate–citrate buffer, 250 mL of mineral trace solution, 20 mL of trace elements solution, and 1 mL of 5 M urea solution. Dissolve and add to 1 L of distilled water. Add 15 g of agar and autoclave. 14. Selection plates without sorbitol: See item 13, but without sorbitol. 15. Spore isolation plates: See item 13, but without sorbitol. Add 0.1% IGEPAL or Triton X-100. 2.2 Isolation of Chromosomal DNA and PCR Screening

1. Glass beads: Diameters of 0.1 mm, 1 mm, and 5 mm. 2. CTAB buffer: 1.4 M NaCl, 100 mM Tris–HCl, pH 8.0, 10 mM EDTA, 2% (v/v) CTAB, 1% (w/v) polyvinylpyrrolidone. 3. FastPrep-24 tissue and cell homogenizer. 4. Phenol. 5. Chloroform.

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6. RNase A: 10 mg/mL. 7. 70% ethanol. 8. TE buffer: 10 mM Tris–HCl, pH 8.0, 1 mM EDTA. 9. NanoDrop spectrophotometer.

3

Methods

3.1 Design of Plasmids

3.2

Transformation

The schematic view of the strategy for the design of the deletion constructions and the constructions with the gene of interest is presented in Fig. 1. The complete sequence of the plasmids carrying these constructs can be found in [15] (see Note 2). For the construction of the Δpyr4 cassette, the 50 -flank and the terminator sequence of pyr4 are amplified and inserted into the plasmid of preference (see Fig. 2). For the construction of the Δasl1 cassette, the coding sequence of hph (hygromycin B resistance) is inserted between the 50 - and 30 -flanking regions of asl1 into the chosen plasmid (see Fig. 3). The plasmids are then transformed into T. reesei. The deletion of pyr4 can be directly selected for using 5-FOA. The deletion of asl1 is selected for using hygromycin B. The resulted strains are auxotrophic for uridine or arginine, respectively (see Figs. 2 and 3). For the construction of the cassette containing the gene of interest using pyr4 as marker, first the complete pyr4 locus is inserted into a plasmid. Afterwards, the coding sequence of the gene of interest in inserted into the previous plasmid downstream of the 50 -flank of pyr4 (see Fig. 2). In the case of using asl1 as marker, the same procedure is followed. The insertion of the complete asl1 locus into the desired plasmid is followed by the insertion of the gene of interest downstream of the 50 -flank of asl1 (see Fig. 3). The resulted strains will have regained the prototrophy for uridine and arginine, respectively (see Figs. 2 and 3). 1. Prepare a light-green spore suspension by harvesting spores from a fresh plate and resuspend them in 1 mL of the sporeharvesting solution using sterile cotton swabs. First, transfer 1 mL of the spore-harvesting solution into a sterile 1.5 mL reaction tube, then wet the cotton swab with the sporeharvesting solution and roll it on the sporulated petri dish. Resuspend spores in the spore-harvesting solution by pressing the head of the cotton swab against the inside of the reaction tube. Harvest approximately an area of 5 cm2. The final spore suspension should have a light moss-green color. 2. Spread freshly autoclaved cellophane discs on MEX plates (containing uridine or arginine, if applicable). Spread 100 μL of the

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Fig. 2 The pyr4 locus during gene deletion and targeted integration. Schematic view of the deletion of pyr4 in the parental strain QM6aΔtmus53 by homologous recombination and the integration of the gene of interest upstream of pyr4 by homologous recombination resulting in a re-established strain additionally bearing the gene of interest. Green arrow represents pyr4, gray boxes represent homologous recombination areas, yellow arrow represents the gene of interest

spore suspension on each plate using a Digralski inoculation loop and incubate them at 30  C for 20 h. 3. Dissolve 200 mg of lysing enzymes in 30 mL of solution A, let stir on a magnetic stirrer for approximately 10 min, and then filter sterile using a 0.45 μm syringe filter (15 mL each in two sterile petri dishes). 4. Transfer mycelium from 2–3 cellophane plates (depending on how well they grew) to a sterile plate containing 15 mL of solution A with 150 mg of lysing enzymes. 5. Tear the mycelium into little pieces using sterile tweezers. 6. Incubate at 30  C for 3–4 h until protoplasts are formed (see Note 3). 7. Filter protoplast suspension through a glass wool funnel and collect in a 50 mL centrifugation tube placed on ice. From here on, keep protoplasts always chilled.

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Fig. 3 The asl1 locus during gene deletion and targeted integration. Schematic view of the partial deletion of asl1 in parental strain QM6aΔtmus53 by homologous recombination and the integration of the gene of interest by homologous recombination resulting in a re-established strain bearing the gene of interest. Green arrow represents asl1, gray boxes represent homologous recombination areas, blue arrow represents the marker gene hph, yellow arrow represents the gene of interest

8. Pellet protoplasts by centrifugation at 3000  g for 10 min at 4  C. 9. Carefully resuspend protoplasts in 4 mL of ice-cold STC solution. 10. Repeat centrifugation in step 8 (see Note 4). 11. Carefully resuspend protoplasts in cold STC solution (200 μL per transformation, although the number of protoplasts should not be lower than 106 per final 200 μL). 12. Prepare as many sterile, pre-chilled 50 mL tubes as different transformations plus one for the negative control. Transfer 200 μL of protoplast suspension to each tube. 13. Add the linearized plasmid DNA. For the negative control, add only 15 μL of sterile water. 14. Add 50 μL of PEG solution. 15. Mix carefully and incubate on ice for 20 min.

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16. Prepare four plates by pouring 10 mL of selection medium in each one and let them chill individually (do not stack them). 17. Add 2 mL of PEG solution to the transformation mixture, mix carefully, and incubate for 5 min on room temperature. 18. Carefully add 4 mL of STC solution. 19. Add molten 50  C warm selection medium to a final volume of 40 mL and mix carefully by inverting twice. 20. Pour 10 mL on each of the previously prepared plates and let chill again individually. 21. Incubate at 30  C until colonies are visible (maximum 1 week). 22. Pick individual colonies by transferring small pieces of overgrown agar to fresh selection plates without sorbitol and let them grow at 30  C until conidiation (see Note 5). 23. Perform a streak out of spores on spore isolation plates. Let the plates incubate at 30  C until single colonies are visible (see Note 6). Pick single colonies by transferring small pieces of overgrown agar to fresh selection plates medium without sorbitol and let them grow at 30  C. 3.3 Isolation of Chromosomal DNA and PCR Screening

1. Weigh 0.37 g of small glass beads (0.1 mm diameter), 0.25 g of medium glass beads (1 mm diameter), and one large glass bead (5 mm diameter) to a 2 mL screw cap tube. 2. Add 1 mL of CTAB buffer and a small amount of mycelium (approximately 50 mg) scraped off a fresh plate or filtered from a liquid culture. 3. Disrupt the mycelium using the FastPrep-24 cell homogenizer (level 4 for 30 s). Alternatively, a piece of mycelium can be frozen in liquid nitrogen and ground using mortar and pestle and then added to a 2 mL reaction tube containing 1 mL of CTAB. 4. Incubate at 65  C for 20 min. 5. Transfer the supernatant (approximately 800 μL) to a fresh 2 mL reaction tube. 6. Add 400 μL of phenol, mix well by vigorous shaking until the foam disappears, add 400 μL of chloroform, and mix again. 7. Incubate at room temperature for 5–10 min. 8. Centrifuge at 4  C and 12,000  g for 10 min. 9. Transfer 650 μL of the top aqueous phase to a fresh 1.5 mL reaction tube. Add 650 μL of chloroform and mix well by shaking for 30 s. 10. Repeat centrifugation.

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11. Transfer 500 μL of the top aqueous phase to a fresh 1.5 mL tube and add 1.5 μL of RNase A. Mix by inverting and incubate at 37  C for 30 min. 12. Add 350 μL of isopropanol, mix well by inverting and incubate for 10 min at room temperature. 13. Precipitate DNA by centrifugation at 4  C and 20,000  g for 30 min. 14. Remove supernatant and wash DNA pellet with 1 mL of 70% ethanol. 15. Centrifuge at 4  C and 20,000  g for 10 min. 16. Remove supernatant and dry at 50 evaporated.



C until ethanol is

17. Dissolve in 200 μL of distilled water and measure concentration and purity using a NanoDrop spectrophotometer. 18. Perform PCR assays using suitable primers to verify the integration at the correct locus (see Fig. 1). We recommend performing a Southern blot analysis to verify the absence of additionally integrated constructs. The forward primer in the 50 -flank and the reverse primer in the 30 -flank must be upstream and downstream, respectively, of the sequence included in the construct (see Fig. 1).

4

Notes 1. We digest 20 μg of plasmid DNA overnight with a suitable restriction enzyme and then precipitate the linearized DNA by adding 2.5 volumes of 96% ethanol. The DNA can be pelleted easily by centrifugation at 4  C and 20,000  g for 30 min. The pellet is then washed with 1 mL of 70% ethanol and, after letting it dry, dissolved in 15 μL of sterile water. 2. The plasmid construction we described in this chapter is described in detail in [15]. The reader can follow their own procedure. Nevertheless, we strongly recommend the use of at least 900 bp for homologous recombination areas and the approximate coordinates described in Figs. 2 and 3. 3. The release of protoplasts can be supported by pipetting the solution every 30 min or 1 h to help the activity of the cell wall degrading enzymes by mechanically breaking down the cell wall. 4. By the end of the centrifugation in this step, the pellet should be a yellowish smear all over the tube wall.

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5. We progress working only with colonies that have formed a uniformly, round colony with well-defined edges after 2 days of incubation. 6. Pick the colonies as soon as possible (normally after 2 days) to minimize the risk of picking a fused colony. References 1. Strauss J, Mach RL, Zeilinger S et al (1995) Crel, the carbon catabolite repressor protein from Trichoderma reesei. FEBS Lett 376:103–107. https://doi.org/10.1016/ 0014-5793(95)01255-5 2. Saloheimo A, Aro N, Ilme´ M, Penttil€a M (2000) Isolation of the ace1 gene encoding a Cys2-His2 transcription factor involved in regulation of activity of the cellulase promoter cbh1of Trichoderma reesei. J Biol Chem 275:5817–5825. https://doi.org/10.1074/ jbc.275.8.5817 3. Aro N, Saloheimo A, Ilme´n M, Penttil€a M (2001) ACEII, a novel transcriptional activator involved in regulation of cellulase and xylanase genes of Trichoderma reesei. J Biol Chem 276:24309–24314. https://doi.org/10. 1074/jbc.M003624200 4. Zeilinger S, Ebner A, Marosits T et al (2001) The Hypocrea jecorina HAP 2/3/5 protein complex binds to the inverted CCAAT-box (ATTGG) within the cbh2 (cellobiohydrolase II-gene) activating element. Mol Gen Genomics 266:56–63. https://doi.org/10.1007/ s004380100518 5. Seiboth B, Karimi RA, Phatale PA et al (2012) The putative protein methyltransferase LAE1 controls cellulase gene expression in Trichoderma reesei. Mol Microbiol 84:1150–1164. https://doi.org/10.1111/j.1365-2958.2012. 08083.x 6. H€akkinen M, Valkonen MJ, WesterholmParvinen A et al (2014) Screening of candidate regulators for cellulase and hemicellulase production in Trichoderma reesei and identification of a factor essential for cellulase production. Biotechnol Biofuels 7:14. https://doi.org/ 10.1186/1754-6834-7-14 7. Saloheimo M, Pakula TM (2012) The cargo and the transport system: secreted proteins and protein secretion in Trichoderma reesei (Hypocrea jecorina). Microbiology 158:46–57. https://doi.org/10.1099/mic.0. 053132-0 8. Penttil€a M, Nevalalnen H, R€atto M et al (1987) A versatile transformation system for

the cellulytic filamentous sungus Trichoderma reesei. Gene 61:155–164. https://doi.org/10. 1016/0378-1119(87)90110-7 9. Schuster A, Bruno KS, Collett JR et al (2012) A versatile toolkit for high throughput functional genomics with Trichoderma reesei. Biotechnol Biofuels 5:1–10. https://doi.org/10. 1186/1754-6834-5-1 10. Lorito M, Hayes C, Di Pietro A, Harman G (1993) Biolistic transformation of Trichoderma harzianum and Gliocladium virens using plasmid and genomic DNA. Curr Genet 24:349–356. https://doi.org/10.1007/ BF00336788 11. De Groot MJA, Bundock P, Hooykaas PJJ, Beijersbergen AGM (1998) Agrobacterium tumefaciens-mediated transformation of filamentous fungi. Nat Biotechnol 16:839–842 12. Steiger MG, Vitikainen M, Uskonen P et al (2011) Transformation system for Hypocrea jecorina (Trichoderma reesei) that favors homologous integration and employs reusable bidirectionally selectable markers. Appl Environ Microbiol 77:114–121. https://doi.org/ 10.1128/AEM.02100-10 13. Gruber F, Visser J, Kubicek CP, de Graaff L (1990) The development of a heterologous transformation system for the cellulolytic fungus Trichoderma reesei based on a pyrG-negative mutant strain. Curr Genet 18:71–76. https://doi.org/10.1007/BF00321118 14. Ouedraogo JP, Arentshorst M, Nikolaev I et al (2015) I-SceI-mediated double-strand DNA breaks stimulate efficient gene targeting in the industrial fungus Trichoderma reesei. Appl Microbiol Biotechnol 99:10083–10095. https://doi.org/10.1007/s00253-015-68291 15. Derntl C, Kiesenhofer DP, Mach RL, MachAigner AR (2015) Novel strategies for genomic manipulation of Trichoderma reesei with the purpose of strain engineering. Appl Environ Microbiol 81:6314–6324. https://doi. org/10.1128/AEM.01545-15

Chapter 6 Open the Pores: Electroporation for the Transformation of Trichoderma reesei Franziska Wanka Abstract During the electroporation of T. reesei, linearized exogenous DNA is absorbed into swollen conidia by an electrical impulse. The advantage of this method is that it is less time-consuming, less expensive, and easier to perform than the classical protoplast transformation while at the same time having a comparable efficiency. Key words Electroporation, Transformation, Strain engineering, Genetic modification, T. reesei, Swollen conidia

1

Introduction Electroporation as a method of transformation describes the induction of uptake of external substances (e.g., DNA) from the surrounding medium into a microorganism by an electrical impulse [1]. The transmembrane electric field pulse induces the generation of short-term microscopic pores in a plasma membrane, which allow the DNA to enter the cell [2]. The reversibility of permeabilization of the cell membrane depends on the constitution of the conidia and the proper amplitude and duration of the pulse. The pulse needs to exceed a certain minimum level, which seems to depend on the cell size. However, if the pulse exceeds a certain threshold, pores become too large for repair and the cells will die. For filamentous fungi, the electrical parameters are assumed with 200–800 Ω resistance, 25 mF capacitance, and 2–15 kV/cm field strength [3]. The first electroporations for filamentous fungi were performed in combination with the protoplast/PEG method [2], but a protocol for swollen conidia was also developed and improved the applicability and the cost-effectiveness of the method [4]. The presented protocol for electroporation of T. reesei was adapted from the patent application US2010/0304468 [5]. For

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this purpose, freshly harvested conidia are allowed to swell for 4–6 h (depending on the strain) in a complex liquid medium. Then the swollen conidia are washed and osmotically stabilized, DNA is added, and the electric shock is applied at 1.8 kV. After the cells have recovered, they are plated on selection plates and incubated for 3–4 days at 28  C. The method described here is less time-consuming and easier to handle, and with re-usage of electroporation cuvettes, it can also be performed at lower costs than protoplast transformation with a similar integration efficiency.

2

Materials Prepare all solutions using ultrapure water (prepared by purifying deionized water, to attain a sensitivity of 18 MΩ-cm at 25  C) and analytical grade reagents. Prepare and store all reagents at room temperature (unless indicated otherwise). Diligently follow all waste disposal regulations when disposing waste materials. 1. Potato dextrose agar plates (PDA): Dissolve 39 g of potato dextrose agar in 1 L of ultrapure water. After autoclaving, pour approximately 20 mL of lukewarm agar into Petri dishes (9 cm diameter). Use PDA plates with respective selection markers for the selection, e.g., with 100 μg/mL hygromycin B, geneticin (G418), or 200 μg/mL nourseothricin. 2. Sodium chloride Tween solution: Dissolve 8.5 g of sodium chloride and 0.5 g of Tween 80 in 1 L of ultrapure water and autoclave. 3. Glass wool tubes (see Note 1). 4. Yeast extract peptone dextrose medium (YPD): Add 10 g of yeast extract and 20 g of peptone to 900 mL of ultrapure water and autoclave. Separately autoclave 20 g of D-glucose monohydrate in 100 mL of ultrapure water and then combine. 5. 1.1 M D-Sorbitol: Weigh 200.42 g of D-sorbitol and add ultrapure water to a volume of 1 L. Autoclave and store at 4  C. 6. Gene Pulser®/MicroPulser™ Electroporation Cuvettes, 0.2 cm gap (Bio-Rad Laboratories, Hercules, CA, USA). 7. MicroPulser (Bio-Rad Laboratories, Hercules, CA, USA).

3

Methods Carry out all procedures at room temperature unless otherwise specified.

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Fig. 1 QM6a at different steps during the electroporation protocol. (a) Fully sporulated PDA plate of QM6a. (b) Self-prepared glass wool tube on top of a 15 mL tube during filtration of the conidia suspension. (c) Erlenmeyer flask containing YPD medium before (left) and after inoculation (right). (d) Pelleted swollen conidia after the first centrifugation. (e) Electroporation set up with cuvette inside the MicroPulser. (f) Transformants start to grow white mycelium on the PDA plate with hygromycin 4 days after electroporation

Fig. 2 Conidia vs. swollen conidia from QM6a. Liquid samples were spread on a microscope slide and were examined with a Leica Microscope DMi8 with a 63 objective. (a) Microscopic picture of a freshly filtered conidia solution before and (b) 4 h after incubation in YPD medium at 30  C and 300 rpm

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Preparations

1. The DNA for the transformation can be prepared by either PCR amplification (see Note 2) or digestion from a plasmid of interest (midi-preparation). To obtain the desired highly concentrated DNA (6–10 μg in 10 μL of ultrapure water per reaction), use sodium acetate precipitation or lyophilization. 2. Grow the desired T. reesei strain on PDA plates until the plate is covered with fresh conidia (10–12 days, see Note 3). One fully sporulated PDA plate is sufficient for four transformation reactions. 3. Pour a dot of 5 mL of sodium chloride Tween solution on the plate and use a cotton stick to detach the conidia. Take a cut blue pipette tip to transfer the suspension through a glass wool tube (see Fig. 1b) into a 15 mL tube. Vortex the tube for 20 s. 4. Subsequently, transfer this conidia solution into an autoclaved 250 mL Erlenmeyer flask containing 100 mL of YPD medium (see Fig. 1c). The incubation time depends on the strain (QM strains: 4 h, Rut-C30 strains: 6 h) and is carried out at 30  C with shaking at 300 rpm. 5. Examine the swelling of the conidia under the microscope (see Fig. 2b) when you apply this method for your strain for the first time. 6. The cell suspension is divided into two 50 mL tubes and pelleted for 5 min at 2200  g and 4  C (see Fig. 1d). 7. After discarding the supernatant, wash the cells in each tube in 25 mL of precooled 1.1 M D-sorbitol and centrifuge for 5 min at 2200  g and 4  C. 8. Resuspend both cell pellets in 1 mL of 1.1 M D-sorbitol and transfer into two 2 mL tubes. Centrifuge at 1500  g and 4  C for 5 min and repeat the last washing step with D-sorbitol. Finally, resuspend the cells in 150 μL of precooled 1.1 M Dsorbitol.

3.2

Electroporation

1. Aliquot 75 μL of the competent conidia into 1.5 mL tubes (four tubes in total) and add 10 μL of DNA or 10 μL of ultrapure water serving as negative control. 2. After incubation on ice for 30 min, transfer the mixture into a sterile electroporation cuvette (see Note 4) and apply an electrical pulse of 1.8 kV with the program Ec1 of the MicroPulser (see Fig. 1e).

3.3 Recovery and Selection

1. For recovery, immediately transfer the cells back into the tube. Add 400 μL of precooled 1.1 M D-sorbitol and 125 μL of YPD (pre-mixed) and then incubate the mixture at 28  C and 800 rpm on a heating block for at least 1 h.

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2. Subsequently, spread out approximately 150 μL of the suspension per PDA plate with the respective selection marker (four plates per reaction). Defined media such as acetamide medium can also be applied. 3. Incubate the plates at 28  C for 3–6 days, depending on the strain and the influence of the genome modification (see Fig. 1f). 4. Streak-isolate the transformants in two rounds. 5. After the second round of isolation, perform screening PCRs. 6. Deposit positive strains as cryostocks with 50% (w/w) glycerol on 80  C.

4

Notes 1. The glass wool tubes are self-prepared (see Fig. 1b). A hole at the bottom of the 1.5 mL tubes is made with a hot needle. Then glass wool is put into the tube with tweezers before autoclaving several of them in a beaker glass. The glass wool retains mycelium and agar residues and allows only conidia to pass through the filter. 2. For the preparation of DNA via PCR, we use the Phusion High-Fidelity DNA Polymerase (Thermo Fisher Scientific, Waltham, USA). In electroporations where the DNA template was produced with the Phire Hot Start II DNA Polymerase (Thermo Fisher Scientific, Waltham, USA), it occurred that the micropulser could not run the Ec1 program. 3. For the preparation of plates with enough conidia for the transformation, QM strains are inoculated in the middle of the plate with a piece of agar or with conidia of a cryovial (see Fig. 1a). Rut-C30, on the other hand, is spread with a cotton stick in 5  5 lines (squares), so that it quickly generates many conidia. 4. The electroporation cuvettes can be reused many times. The used cuvette is rinsed three times with 70% ethanol and then thoroughly washed with deionized water. After drying, it is sterilized under UV light for approximately 5 min and then stored with the lid on at room temperature.

References 1. Neumann E, Schaefer-Ridder M, Wang Y, Hofschneider PH (1982) Gene transfer into mouse lyoma cells by electroporation in high electric fields. EMBO J 1(7):841–845

2. Bowyer P (2001) DNA-mediated transformation of fungi. Oxford University Press, Oxford, pp 33–46 3. Sa`nchez O, Aguirre J (1996) Efficient transformation of Aspergillus nidulans by

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electroporation of germinated conidia. Fungal Genet Rep 43(21):48–51. https://doi.org/10. 4148/1941-4765.1317 4. Schuster A, Bruno KS, Collett JR, Baker SE, Seiboth B, Kubicek CP, Schmoll M (2012) A versatile toolkit for high throughput functional genomics with Trichoderma reesei. Biotechnol

Biofuels 5(1):1. https://doi.org/10.1186/ 1754-6834-5-1 5. Kim S, Miasnikov A (2013) Method for introducing nucleic acids into fungal cells. US Patent application US 2010/0304468 A1, 23 Aug 2010

Chapter 7 Sexual Crossing of Trichoderma reesei Rita B. Linke Abstract This chapter describes how mating assays in Trichoderma reesei can successfully be performed and which specific prerequisites of industrial strains originating from strain QM6a have to be met for successful mating experiments. Key words Trichoderma reesei, Mating assay, Fruiting body, Ascospores, Sexual crossing

1

Introduction Filamentous fungi represent key organisms in many industrial and biotechnological applications, e.g., for the production of enzymes, vitamins, polysaccharides, polyhydric alcohols, pigments, lipids, organic acids, and glycolipids [1, 2]. In contrast to plants and yeasts for which strain improvement is for a long time accomplished by crossing of different strains with desirable traits, these techniques are not applicable for many filamentous fungi due to the lack of the knowledge of a sexual reproduction cycle. Only a minority of fungal species (approximately 20%) is known to reproduce by sexual means [3]. However, this does not necessarily mean that a sexual state of a fungal species does not exist. Several species which were believed to reproduce solely asexually were identified to be able to undergo sexual reproduction during the past years [4, 5], e.g., Candida albicans [6], Aspergillus flavus [7], Aspergillus fumigatus [8], Penicillium roqueforti [9], and Trichoderma reesei [10]. Once the sexual reproduction cycle of a fungal species has been identified, it can provide a valuable laboratory tool. By crossing of strains exhibiting different genetic traits and subsequent analysis of the progeny, it is possible to determine the function of genes and whether a trait is mono- or polygenic in basis [11, 12]. In combination with next-generation sequencing methodologies, it can be used to locate and identify individual genes of interest [11, 13]. For biotechnological applications, it offers the ability to apply classical

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breeding strategies for strain improvement as used in plant breeding programs, to restore strain fitness of highly mutated industrial strains by outcrossing with wild-type strains or to outcross undesired genes like secondary metabolite clusters or antibiotic resistance markers whose presence may interfere with regulatory requirements [14]. For more comprehensive details and examples of laboratory applications of the sexual cycle in fungal species, the reader is referred to [11, 15]. The species Trichoderma reesei was also long believed to reproduce solely asexually. In 2009, Seidl and coworkers [10] first described the ability of T. reesei to undergo sexual reproduction. They performed a screening of the T. reesei genome database v2.0 [16], which lead to the identification of the mat1-2-1 gene. The Neurospora crassa MAT1-2 idiomorph Mat1-2-1 encodes a protein with a high mobility group domain, as the main regulator of sexual development in MAT1-2 strains. Tblastn searches of the T. reesei genome database with the mat1-2-1 homologs of other fungal species then revealed that T. reesei strain QM6a carries the MAT12 mating type locus and was thereby identified to be heterothallic, meaning that the two mating type loci, MAT1-1 and MAT1-2, are located on different strains [17]. Consequently, for successful sexual reproduction, two strains of opposite mating type are needed necessitating the availability of a strain of opposite MAT1-1 mating type. To identify a MAT1-1 strain of T. reesei, Seidl et al. [10] made use of a single ascospore culture of H. jecorina CBS999.97, a strain formerly reported to be able to form mature fruiting bodies on agar plates under laboratory conditions [18]. Fruiting bodies of Hypocrea spp., so-called stromata (see Fig. 1), consist of a pigmented hyphal mass in which the actual fruiting bodies, the perithecia (see Fig. 1), are embedded [19]. These contain the asci with each 16 ascospores (see Fig. 1) that are ejected from the fruiting body upon maturation. For sexual crossings to be applied in T. reesei, the challenge that all strains nowadays used in industry and most strains used in academic research originate from the same isolate, QM6a [20], which was identified to carry the MAT1-2 locus, had to be taken. A first attempt to simply switch the mating type in one of the QM6a lineages to induce sexual reproduction failed, and the T. reesei QM6a strain line was identified to be female sterile [10], i.e., the strain is unable to produce the specialized hyphal tissues needed for fruiting body development [21]. Recently, however, a single point mutation in the ham5 gene of T. reesei was identified as the cause of female sterility [14] paving the way for the application of sexual crossings as laboratory tool. For successful crossing of two QM6a derived lineages, therefore, two technical requirements have to be met—the mating type of one partner has to be switched to MAT1-1 and female fertility has to be restored by the introduction of a functional version of ham5 from a T. reesei wild-type strain.

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Fig. 1 Left photo: Stromatum with embedded perithecia. Right photo: Cross-section through a single perithecium containing asci with 16 bipart ascospores

2

Materials 1. T. reesei strains to be crossed (see Note 1). 2. Medium for strain cultivation: Either potato dextrose agar (PDA) or malt extract (MEX) agar (3% malt extract and 1.5% agar) can be used depending on which medium the used strains grow better. 3. Plates for single spore isolation: Either PDA or MEX agar supplemented with 0.1% Triton X-100. 4. Physiological salt solution: 0.8% NaCl with 0.1% Tween-80. 5. Sterile glass wool tubes: 1.5 mL reaction tubes stuffed with glass wool and a small hole in the bottom of the tubes. 6. Sterile cotton swabs.

3

Methods

3.1 Preparatory Work

1. Make sure to use purified cultures of the strains to be crossed. Therefore, cultivate strains either on PDA or MEX under conditions inducing conidiation (at 28  C in darkness). 2. Harvest conidiospores in sterile physiological salt solution and filter suspension over glass wool tubes to remove rests of mycelium. 3. Perform a single-cell streak out on plates for single-spore isolation.

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Fig. 2 Mating assay of T. reesei QM6a and H. jecorina CBS999.97. Fruiting bodies are formed in the contact zone of the two strains as indicated by black arrows

4. Grow purified strains on PDA or MEX plates (at 28  C in darkness). 5. Optional: Perform a PCR to test for the presence of the appropriate mating type (MAT1-1 or MAT1-2). 3.2

Mating Assay

1. Cut a piece of agar from each plate containing one of the mating partners (see Note 2). 2. Place the agar pieces on opposite sides of a PDA or MEX plate (see Notes 3 and 4). 3. Place the plates on the benchtop at temperature between 18 and 22  C under day/night rhythm (see Note 5) until formation of fruiting bodies (Fig. 2), and ascospore discharge can be detected (see Note 6) as a white haze on the cover of the Petri dish (see Notes 7–9).

3.3 Harvest of Ascospores

1. Check plates regularly for the appearance of ascospores ejected from fruiting bodies. 2. Use a sterile cotton swab soaked with physiological salt to harvest ascospores from the cover of the Petri dish (see Notes 10–12). 3. Plate ascospores on plates for single spore isolation to obtain single colonies. 4. Test colonies for the desired traits.

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Notes 1. If two QM6a derivatives are intended to be crossed, the mating type of one of the mating partners has to be switched from MAT1-2 to MAT1-1. Additionally, one of the strains has to be complemented with a functional version of ham5 to restore female fertility (compare [14]). 2. Alternatively, spore suspensions of the two mating partners can be mixed and plated on either PDA or MEX plates. Fruiting bodies will occur spread on the entire plate. However, most often fruiting bodies formed will be smaller than when strains are opposed on the plate, and fruiting bodies only form in the contact zone, which makes handling more difficult. 3. Fruiting bodies are not formed immediately after the strains get in contact with each other. Up to 2 weeks might pass after contact until fruiting bodies are formed. For this reason, it is worth to pour thicker plates, so that the medium does not dry out until fruiting bodies are formed. 4. Do not close plates with parafilm. 5. Most strain combinations form fruiting bodies only when plates are subjected to day/night rhythm. Some strain combinations, e.g., when using strains CBS999.97 MAT1-1 and CBS999.97 MAT1-2 combination, however, have been found to form fruiting bodies also when kept in the incubator at 28  C in darkness (cf. [10]). 6. Time needed for fruiting body formation varies greatly. If strains CBS999.97 MAT1-1 or CBS999.97 MAT1-2 are used as one of the mating partners, fruiting bodies will be obtained after 7–14 days. For other strain combinations, much more time might elapse before fruiting bodies are formed. 7. It has to be noted that strains of opposite mating type not necessarily produce fruiting bodies if opposed in a mating assay. It seems several strains have lost capability to form fruiting bodies to undergo sexual reproduction, either generally or with specific partners or under specific conditions. This was frequently observed when crossing of different wild-type strains was attempted. 8. Fruiting bodies are formed after contact of the two mating partners. If the aim is to produce ascospores only, mating assays can be pre-grown at 28  C until the partners get in contact with each other and be then transferred to the benchtop with day/night rhythm. 9. Another possibility to speed up the process of fruiting body formation is to use smaller Petri dishes, so that mating partners get in contact quicker.

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10. Fruiting bodies vary in size. Some of them are very small, others bigger. Unfortunately, ascospores are not always ejected from the fruiting bodies, which makes it harder to harvest them. If ascospores are not ejected from the fruiting bodies, harvest the fruiting body. Rinse the fruiting body with sterile physiological salt solution and wipe the surface with a wetted sterile cotton swab to rub off conidiospores from the surface. Then cut the fruiting body and squeeze out ascospores. The resultant spore suspension is then filtered over sterile reaction tubes containing glass wool. 11. Alternatively, picked fruiting bodies can be rolled across the surface of a MEX plate to remove adhering conidiospores. Thereafter proceed as described in Note 10. Unfortunately, the risk of co-harvesting some conidiospores still remains. 12. Alternatively, ascospores might also be harvested using a micromanipulator to sequentially isolate each ascospore from a hexadecade. References 1. Adrio JL, Demain AL (2003) Fungal biotechnology. Int Microbiol 6:191–199 2. Rokem JS (2010) Industrial Mycology. In: Doelle HW, Rokem S, Berovic M (eds) Biotechnology, vol 6. Encyclopedia of Life Support Systems Publishers, Developed under the Auspices of the UNESCO. Eolss Publishers, Paris, pp 75–97 3. Dyer PS, Paoletti M (2005) Reproduction in Aspergillus fumigatus: sexuality in a supposedly asexual species? Med Mycol 43(Suppl 1):7–14 4. Dyer PS, O’Gorman CM (2011) A fungal sexual revolution: Aspergillus and Penicillium show the way. Curr Opin Microbiol 14 (6):649–654 5. Martin SH, Steenkamp ET, Wingfield MJ, Wingfield BD (2013) Mate-recognition and species boundaries in the ascomycetes. Fungal Divers 58(1):1–12 6. Hull CM, Raisner RM, Johnson AD (2000) Evidence for mating of the “asexual” yeast Candida albicans in a mammalian host. Science 289:307–310 7. Horn BW, Moore GG, Carbone I (2009) Sexual reproduction in Aspergillus flavus. Mycologia 101:423–429 8. O’Gorman CM, Fuller H, Dyer PS (2009) Discovery of a sexual cycle in the opportunistic fungal pathogen Aspergillus fumigatus. Nature 457:471–474 9. Ropars J, Lo´pez-Villavicencio M, Dupont J, Snirc A, Gillot G, Coton M, Jany JL,

Coton E, Giraud T (2014) Induction of sexual reproduction and genetic diversity in the cheese fungus Penicillium roqueforti. Evol Appl 7:433–441 10. Seidl V, Seibel C, Kubicek CP, Schmoll M (2009) Sexual development in the industrial workhorse Trichoderma reesei. Proc Natl Acad Sci USA 106(33):13909–13914 11. Ashton GD, Dyer PS (2016) Sexual development in fungi and its uses in gene expression systems. In: Schmoll M, Dattenbo¨ck C (eds) Gene expression systems in fungi: advancements and applications. Springer, Cham, pp 335–350 12. Chuang YC, Li WC, Chen CL, Hsu PW, Tung SY, Kuo HC, Schmoll M, Wang TF (2015) Trichoderma reesei meiosis generates segmentally aneuploid progeny with higher xylanaseproducing capability. Biotechnol Biofuels 8:30 13. Moore D, Novak Frazer LA (2002) Essential fungal genetics. Springer-Verlag, New York 14. Linke RB, Thallinger GG, Haarmann T, Eidner J, Schreiter M, Lorenz P, Seiboth B, Kubicek CP (2015) Restoration of female fertility in Trichoderma reesei QM6a provides the basis for inbreeding in this industrial cellulase producing fungus. Biotechnol Biofuels 8:155 15. Houbraken J, Dyer PS (2015) Induction of the sexual cycle in filamentous ascomycetes. In: van den Berg MA, Maruthachalam K (eds) Genetic transformation systems in fungi, Fungal biology, vol 2. Springer, Cham

Sexual Crossing 16. Martinez D, Berka RM, Henrissat B, Saloheimo M, Arvas M, Baker SE, Chapman J, Chertkov O, Coutinho PM, Cullen D, Danchin EGJ, Grigoriev IV, Harris P, Jackson M, Kubicek CP, Han CS, Ho I, Larrondo LF, de Leon AL, Magnuson JK, Merino S, Misra M, Nelson B, Putnam N, Robbertse B, Salamov AA, Schmoll M, Terry A, Thayer N, Westerholm-Parvinen A, Schoch CL, Yao J, Barabote R, Barbote R, Nelson MA, Detter C, Bruce D, Kuske CR, Xie G, Richardson P, Rokhsar DS, Lucas SM, Rubin EM, Dunn-Coleman N, Ward M, Brettin TS (2008) Genome sequencing and analysis of the biomass-degrading fungus Trichoderma reesei (syn. Hypocrea jecorina). Nat Biotechnol 26:553–560 17. Debuchy R, Berteaux-Lecellier V, Silar P (2010) Mating systems and sexual

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morphogenesis in ascomycetes. In: Borkovich KA, Ebbole DJ (eds) Cellular and molecular biology of filamentous fungi. ASM, Washington, pp 501–535 18. Lieckfeldt E, Kullnig CM, Samuels GJ, Kubicek CP (2000) Sexually competent, sucroseand nitrate-assimilating strains of Hypocrea jecorina (Trichoderma reesei) from South American soils. Mycologia 92:374–380 19. Samuels GJ, Petrini O, Manguin S (1994) Morphological and macromolecular characterization of Hypocrea schweinitzii and its Trichoderma anamorph. Mycologia 86:421–435 20. Reese ET (1976) History of the cellulase program at the U.S. army Natick Development Center. Biotechnol Bioeng Symp 6:9–20 21. Dyer PS, Kueck U (2017) Sex and the imperfect fungi. Microbiol Spectr 5(3)

Chapter 8 CRISPR/Cas9-Mediated Genome Editing of Trichoderma reesei Gen Zou and Zhihua Zhou Abstract In this protocol, we describe the establishment of a CRISPR/Cas9 system in Trichoderma reesei by generating a specific, codon-optimized Cas9-expressing strain and by in vitro transcription of a gRNA. This system induces mutagenesis or introduces a gene in a targeted way based on PEG-mediated protoplast transformation. Up to three targets, multiplexed genome editing can be obtained in one transformation. Key words CRISPR/Cas9, Protoplast transformation, Specific codon optimization, gRNA transcription, Multiplexed genome editing

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Introduction The type II clustered regularly interspaced short palindromic repeats (CRISPR)/CRISPR-associated gene (Cas) system is the most popular genome editing tool at this time [1–4]. A single multidomain Cas9 catalyzes a double-strand break in the target DNA composed of a 20-bp sequence matching the protospacer of the guide RNA (gRNA) and an adjacent downstream 50 -NGG-30 nucleotide sequence (termed as the protospacer-adjacent motif (PAM)) [5]. Many studies have demonstrated that the CRISPR/ Cas9 system is a powerful genome editing method that facilitates genetic alterations in genomes in a variety of organisms [1, 6– 9]. The filamentous fungus Trichoderma reesei, which is recognized by its Generally Recognized as Safe status by the US Food and Drug Administration, is the most widely used producer of commercial lignocellulolytic enzyme preparations [10] and is expected to be a potential cell factory for different heterologous proteins due to its powerful ability to synthesize and secrete proteins in high quantities [11]. The establishment of a genome editing system can be used to develop T. reesei as a super cell factory for lignocellulolytic enzyme preparations and other heterologous proteins as well

Astrid R. Mach-Aigner and Roland Martzy (eds.), Trichoderma reesei: Methods and Protocols, Methods in Molecular Biology, vol. 2234, https://doi.org/10.1007/978-1-0716-1048-0_8, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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as to characterize the regulatory mechanisms for induction, synthesis, and secretion of proteins [12–14]. In most cases, fungal codon optimization is necessary for Cas9 expression [15, 16]. Constitutive promoters that are not induced by external factors are most commonly used to drive the Cas9 expression [15, 17, 18]. However, many studies have shown that the expression of Cas9 may have negative effects in recipient organisms [19, 20]. Therefore, transient or inducible expression of Cas9 is a much more reliable strategy [16, 21]. RNA polymerase type III U6 promoters are often used for gRNA transcription [22]. Although the mature U6 snRNAs are highly conserved from yeast to mammals, the U6 snRNA gene sequences are interrupted by introns in some fungi. And the RNA polymerase III-associated B box promoter element is translocated into the intron sequence [23] (see Note 1). Due to the complex structure of U6 snRNA, other RNA polymerase III promoters such as U3 promoter [24], different tRNA promoters [25], and 5S rRNA promoter [26] are used for transcription of gRNA. RNA polymerase II promoters facilitated by ribozymes are also effective to generate gRNA [26] (see Note 2). Although there are many strategies to express Cas9 and generate gRNA in the fungal cell, the transient transformation of in vitro synthesized Cas9 protein and single gRNA is a safe option to reduce side effects [16, 27]. In this protocol, we demonstrate how to express the Cas9 endonuclease in T. reesei after codon optimization, and how to employ the CRISPR/Cas9 system to induce mutagenesis or introduce a new gene by homologous recombination into a target site of the T. reesei genome. In addition, it can be used to simultaneously generate multiple genome modifications by co-transforming gRNAs and donor DNAs for different targets. With small modifications and after the introduction of a specific, codon-optimized Cas9 endonuclease, this method can also be applied for other filamentous fungi that are compatible with protoplast transformation. CRISPR/Cas9-stimulated genome editing is based on a sophisticated genetic manipulation system. In this protocol, we will employ the Agrobacterium-mediated transformation [28] and PEG-mediated protoplast transformation [16] to establish the CRISPR/Cas9 system in T. reesei.

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Materials Prepare all solutions using double-distilled water and analytical grade reagents. Prepare and store all reagents at room temperature (unless indicated otherwise). Diligently follow all waste disposal regulations when disposing waste materials.

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2.1 Construction of a Cas9-Expressing T. reesei Strain

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1. Trichoderma reesei strain, e.g., ATCC56765. 2. Agrobacterium strain that harbors the gene encoding a codonoptimized Cas9 with a nuclear localization signal (NLS) and a selection marker gene. E.g., strain LBA1100 that carries a binary vector harboring the gene encoding the codonoptimized Cas9 with a SV40 NLS (toCas9) and a gene conferring hygromycin resistance (see Notes 3–7). 3. Physiological salt solution: 0.02% (v/v) Tween 80. Sterilize by autoclaving at 68 kPa pressure (115  C) for 30 min. 4. Potato dextrose agar (PDA) plates for Trichoderma sporulation: Add 20 g of dextrose and 15 g of agar into infusion from 200 g of potatoes. Add distilled water to a total volume of 1 L. Sterilize by autoclaving at 68 kPa pressure (115  C) for 30 min. Mix well before dispensing. 5. PDA screening plates: Add 20 g dextrose and 15 g of agar into infusion from 200 g of potatoes. Add distilled water to a total volume of 1 L. Sterilize by autoclaving at 68 kPa pressure (115  C) for 30 min. Add 1 mL of 0.2 M cefotaxim (final concentration 200 μM; kills Agrobacterium cells) and 600 μL of 50 mg/mL hygromycin (final concentration 30 μg/mL; for the selection of T. reesei transformants). Mix well before dispensing. 6. Liquid low-sodium culture (LC) medium: Dissolve 8 g of NaCl, 10 g of tryptone and 5 g of yeast extract in distilled water to a total volume of 1 L. Sterilize by autoclaving at 98 kPa pressure (121  C) for 20 min. Add 200 mL of 50 mg/mL kanamycin solution (final concentration 100 mg/ mL) and 200 mL of 10 mg/mL rifampicin solution (final concentration 20 mg/mL) to 100 mL of medium. 7. LC medium plates: Dissolve 8 g of NaCl, 10 g of tryptone, 5 g of yeast extract, and 15 g of agar in water to a total volume of 1 L. Sterilize by autoclaving at 98 kPa pressure (121  C) for 20 min. Add 200 mL of 50 mg/ml kanamycin solution (final concentration 100 mg/mL) and 200 mL of 10 mg/mL rifampicin solution (final concentration 20 mg/mL) to 100 mL of medium and pour into Petri dishes. 8. K-buffer: Add 1.25 M KH2PO4 solution to 1.25 M K2HPO4 solution until pH 4.8 is reached. Sterilize by autoclaving at 98 kPa pressure (121  C) for 20 min. 9. 1 M MES: Dissolve 195.24 g of MES in water to a total volume of 1 L. Adjust to pH 5.5 by adding NaOH and filter-sterilize. Solution can be stored for a month in the dark or aliquoted and frozen at 20  C.

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10. MN buffer: Dissolve 30 g of MgSO4∙7H2O and 15 g of NaCl in water to a total volume of 1 L. Sterilize by autoclaving at 98 kPa pressure (121  C) for 20 min. 11. Induction medium (IM) (liquid): Add 0.8 mL of K-buffer, 20 mL of MN buffer, 1 mL of 1% (w/v) CaCl2∙2H2O, 10 mL of 0.01% (w/v) FeSO4, 5 mL of trace elements for IM, 2.5 mL of 20% (w/v) NH4NO3, 10 mL of 50% (v/v) glycerol and 10 mL of 20% (w/v) glucose to 900.7 mL of sterilized water to obtain 969 mL of liquid IM. Sterilize by autoclaving at 68 kPa pressure (115  C) for 30 min. Add 40 mL of 1 M MES, pH 5.5. Mix well before dispensing. 12. IM (solid): Dissolve 15 g of agar in water to a total volume of 905.7 mL. Add 0.8 mL of K-buffer, 20 mL of MN buffer, 1 mL of 1% (w/v) CaCl2∙2H2O, 10 mL of 0.01% (w/v) FeSO4, 5 mL of trace elements for IM, 2.5 mL of 20% (w/v) NH4NO3, 10 mL of 50% (v/v) glycerol, and 5 mL of 20% (w/v) glucose to obtain a total volume of 960 mL. Sterilize by autoclaving at 68 kPa pressure (115  C) for 30 min. Add 40 mL of 1 M MES, pH 5.5. Mix well before dispensing. 13. Trace elements for IM: Dissolve 100 mg of ZnSO4∙7H2O, 100 mg of CuSO4∙5H2O, 100 mg of H3BO3, 100 mg of MnSO4∙H2O, and 100 mg of Na2MoO4∙2H2O in water to a total volume of 1 L. Sterilize by autoclaving at 98 kPa pressure (121  C) for 20 min. 14. 0.2 M AS (30 ,50 -dimethoxy-40 -hydroxyacetophenone, HOC6H2(OCH3)2COCH3): Dissolve 785 mg of AS in DMSO to a total volume of 20 mL. Aliquot and store in the dark at 20  C (see Note 8). 15. Glass spreader. 16. Microcentrifuge. 2.2 Preparation of gRNA In Vitro

1. Polymerase for the amplification of the gRNA template DNA, e.g., Q5 polymerase. 2. Primer (e.g., Forward: 50 -TAATACGACTCACTATAGGGCG AGGGCGGCAACATCGT TTTTAGAGCTAGAAATAGCAA GTTAAAA-30 . Reverse: 50 -AAAAGCACCGACTCGGTGCC ACTTTTTCAAGTTGATAACGGACTAGCCTTATTTTAAC TTGCTATTTCTAGCTCTAAA-30 ) (see Note 9). 3. Phenol-chloroform-isoamyl alcohol: 25:24:1 (v/v/v), saturated with 10 mM Tris–HCl, pH 8.0, 1 mM EDTA. 4. Chloroform. 5. Absolute ethanol, ice cold. 6. 3 M sodium acetate, pH 5.2. 7. 70% (v/v) ethanol.

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8. Kit for in vitro transcription, e.g., MEGAscript® Kit. 9. Stop solution: 5 M ammonium acetate, 100 mM EDTA, pH 8.0. 10. RNA precipitation solution: 7.5 M lithium chloride, 50 mM EDTA, pH 8.0. 11. Deoxyribonuclease I (DNaseI): e.g., TURBO™ DNase. 12. 1.5 mL reaction tubes. 13. Diethyl pyrocarbonate (DEPC). 14. Centrifuge and microcentrifuge. 2.3 Protoplast Transformation

1. Cellophane cut in discs fitting PDA plates. 2. Lysing enzymes for protoplast preparation: E.g., lysing enzymes from Trichoderma harzianum. 3. Filter for protoplasts: E.g., folded miracloth in two layers in a glass hopper. 4. Solution I: 1.2 M sorbitol, 0.1 M KH2PO4, pH 5.5. Sterilize by autoclaving at 98 kPa pressure (121  C) for 20 min. 5. Solution II: 1 M sorbitol, 50 mM CaCl2, 10 mM Tris–HCl, pH 7.5. Sterilize by autoclaving at 98 kPa pressure (121  C) for 20 min. 6. Solution III: 25% (w/v) PEG6000, 50 mM CaCl2, 10 mM Tris–HCl, pH 7.5. Sterilize by autoclaving at 98 kPa pressure (121  C) for 20 min. 7. Regeneration medium (RM) for protoplasts: Dissolve 182.17 g of sorbitol, 1 g of MgSO4∙7H2O, 10 g of KH2PO4, 6 g of (NH4)2SO4, 10 g of tri-sodium citrate∙2H2O, 10 g of glucose∙H2O, 1 mL of trace elements for RM, and 12 g of low melting point agarose in water to a total volume of 1 L. Sterilize by autoclaving at 68 kPa pressure (115  C) for 30 min. Mix well before dispensing. 8. Trace elements for RM: Dissolve 5 g of FeSO4∙7H2O, 1.6 g of MnSO4∙H2O, 1.4 g of ZnSO4∙7 H2O, and 1.2 g of CoCl2 in water to a total volume of 1 L. Sterilize by autoclaving at 98 kPa pressure (121  C) for 20 min. 9. Transformant screening reagent: E.g., 100 mg/mL of 5-fluoroorotic acid monohydrate (5-FOA) dissolved in DMSO, filter-sterilized. 10. Supplement for auxotrophic transformants: 1.25 M uridine, filter-sterilized. 11. Petri dishes (9 cm diameter). 12. 50-mL centrifugation tubes.

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Methods

3.1 Expression of toCas9 in T. reesei

1. Transfer a single colony of an Agrobacterium strain, which carries the gene encoding a codon-optimized Cas9 with an NLS and a selection marker gene, from an LC medium plate into 4 mL of liquid LC medium. Cultivate at 28  C with shaking at 200 rpm for 24 h. 2. Spin down 1 mL of the Agrobacterium culture in a microcentrifuge for 10 min at 5000  g at room temperature. Remove the supernatant and wash the cells by gently resuspending the pellet in 10 mL of liquid IM with 0.2 μM AS for 7 h at 28  C with shaking at 200 rpm. 3. To obtain conidiospores of T. reesei, which are used as recipients of toCas9 with a SV40 NLS, grow T. reesei for 7 days on a PDA plate at 28  C (see Note 10). 4. Harvest the spores from the plate by adding 3 mL of physiological salt solution and gently rubbing the surface with a glass spreader. 5. Mix 100 μL of the induced Agrobacterium cells (OD600 0.6–0.8) and 100 μL of the Trichoderma conidiospores (concentration of 1  107 spores per mL). 6. Pipette 200 μL of the cell-conidiospore mixture onto an IM plate with 0.2 μM AS and spread the mixture evenly using a glass spreader. Incubate the plate for 2 days at 26  C in the dark. 7. Collect Agrobacterium–T. reesei cell mixture from the plate by adding 2 mL of physiological salt solution and rubbing the surface with a glass spreader. 8. Transfer the mixture onto PDA screening plates using sterile tweezers. Incubate the plates for 4 days at 28  C in the dark until colonies appear. 9. Transfer the transformants to PDA screening plates for purification and single conidiospore isolation and further verification. A positive transformant is used as a recipient strain for gRNA in the following procedure.

3.2 Transcription of gRNA

1. The template DNA, including the T7 promoter (see Note 11), the 20-bp target sequence (see Note 12), the crRNA, the tracrRNA, and a polyT terminator, is amplified as a dimer using a pair of primers (forward and reverse) with 28 bp overlapping sequence (see Fig. 1). The PCR (40 cycles) essentially follows the instruction of the used polymerase, with modified primer concentration (fivefold) and without addition of any template (see Note 13).

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Fig. 1 Scheme for the design of the gRNA template DNA and its mature product. (a) Schematic drawing of the components of the template DNA: sequence of the T7 promoter is shown in underlined letters. The promoter sequence has a 3-bp overlap with the protospacer. The first base incorporated into the RNA during transcription is shown in bold letter. The preferred base of the first and second base of the protospacer is a guanine in order to ensure efficient transcription. The sequence of the terminator is shown in italic letters. (b) Scheme of the mature gRNA (chimeric RNA). crRNA, CRISPR-derived RNA; tracrRNA, trans-activating crRNA

2. Transfer the reaction to a 1.5 mL reaction tube and dilute to 500 μL with water. Add 500 μL of phenol-chloroform-isoamyl alcohol and mix thoroughly. 3. Spin down the mixture in a microcentrifuge for 10 min at 10,000  g, 4  C. Transfer the supernatant to a new reaction tube and mix thoroughly with an equal volume of chloroform. 4. Spin down the mixture in a microcentrifuge for 10 min at 10,000  g, 4  C. Transfer the supernatant to a new reaction tube. Add three volumes of ice-cold ethanol and 0.1 volumes of 3 M sodium acetate. Mix thoroughly and keep at 20  C for at least 30 min. 5. Then pellet the DNA for 15 min in a microcentrifuge at 10,000  g, 4  C. Remove the supernatant and wash the pellet with 1 mL 70% ethanol. 6. Centrifuge again for 5 min at 10,000  g, 4  C. Remove the supernatant most completely with a very fine-tipped pipet and dry the tube at air for 5 min. 7. Resuspend the template DNA in DEPC-treated water at a concentration of 0.1–0.5 μg/μL. 8. Mix 1 μL of template DNA, ATP solution, CTP solution, GTP solution, UTP solution, 10 reaction buffer, and enzyme mix

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from an in vitro transcription kit. Add DEPC-treated water to a total volume of 10 μL (see Note 14). 9. Gently flick the tube or gently pipette the mixture up and down and then microfuge briefly to collect the reaction mixture at the bottom of the tube. Incubate overnight at 37  C (see Note 15). 10. Add 0.5 μL of DNaseI, mix thoroughly and incubate 15 min at 37  C (see Note 16). 11. Stop the reaction using 1 μL of stop solution and precipitate the RNA by adding 20 μL of DEPC-treated water and 30 μL of RNA precipitation solution. Mix thoroughly. Keep at 20  C for at least 30 min. 12. Centrifuge at 10,000  g for 15 min to pellet the gRNA. Remove the supernatant and wash the pellet with 1 mL of 70% ethanol. Centrifuge again at 10,000  g (4  C) for 10 min to maximize the removal of unincorporated nucleotides. 13. Remove the supernatant most completely with a very finetipped pipet and dry the tube at air for 5 min. 14. Resuspend the gRNA in 10 μL of DEPC-treated water. 15. Determine the concentration of the gRNA and store at 70  C. 3.3 Genome Editing Based on Protoplast Transformation

1. To obtain conidiospores of a toCas9 expressing T. reesei, grow the strain (obtained at step 8, see Subheading 3.1) for 7 days on a PDA plate at 28  C. 2. Harvest the spores from the fungal plate by adding 3 mL of physiological salt solution and gently rubbing the surface with a glass spreader. 3. Pipette 100 μL of spore suspension (~105 spores) onto a cellophane disc placed on a PDA plate and spread evenly using a glass spreader. Prepare six to eight plates and incubate for 12 h at 28  C. 4. Weigh 0.1 g of lysing enzyme to 20 mL of solution I. Mix thoroughly. 5. Wash and collect the mycelium from the cellophane discs using solution I with lysing enzyme. Transfer the mixture into a new Petri dish and incubate for 2 h at 28  C with shaking at 10–15 rpm. 6. Filter protoplast suspension in a 50-mL centrifugation tube by using miracloth. 7. Centrifuge at 1500  g for 10 min and remove the supernatant. Wash the protoplast pellet with 4 mL solution II. 8. Centrifuge at 1500  g for 10 min and remove the supernatant. Suspend the protoplast pellet with 250 μL solution II.

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9. Dilute the protoplast suspension to approximately 107 protoplasts per mL with solution II. 10. Add 50 μL of solution III and 10–50 μg of gRNA (obtained at step 14, see Subheading 3.2) (see Notes 17–19) to 200 μL of diluted protoplast suspension and mix very gently (see Note 20). Keep on ice for 20 min. 11. Add 2 mL of solution III and mix very gently. Incubate at 25  C for at least 5 min. 12. Melt 50 mL of RM and cool down to about 50  C. Add 0.2 mL of 1.25 M uridine (final concentration 5 mM) and 0.75 mL of 100 mg/mL 5-FOA (final concentration 1.5 mg/mL). 13. Transfer the protoplast suspension to 50.95 mL of RM. Mix gently and pour evenly into three Petri dishes. Incubate at 28  C for 7 days in the dark until colonies appear (see Note 21). 14. Transfer the transformants to selection medium (see Note 22) for purification and single conidiospore isolation and subsequent genomic analysis.

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Notes 1. The U6 promoter is the most commonly used type III promoter for in vivo synthesis of gRNA in most organisms. However, the U6 promoter has a more intricate structure (the B box promoter element is translocated into the downstream of transcription start site) in T. reesei [23] than those in many other organisms such as Triticum aestivum or Arabidopsis thaliana [29, 30]. 2. The gRNA can also be synthesized in vivo by type II promoter with the help of a ribozyme. The hepatitis delta virus (HDV) RNA and hammerhead (HH) ribozyme are most often used to synthesize gRNA by the HH-gRNA-HDV construct. Ribozyme processes the RNA transcripts to unit lengths in a selfcleavage reaction [31]. 3. The toCas9 with a SV40 NLS has been constructed into a binary vector. It can be easily inquired from addgene. Addgene no. 92125 (pDHt/sk-PC) is a construct with a strong constitutive promoter of T. reesei (namely ppdc) (https://www. addgene.org/92125/). Addgene no. 92127 (pDHt/sk-CC) is a construct with an inducible promoter of T. reesei (namely pcbh1) (https://www.addgene.org/92127/). 4. The plasmids pDHt/sk-PC and pDHt/sk-C bear the NLS fused at the C-terminus. An additional NLS at the N-terminus may increase editing efficiency.

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5. The plasmids pDHt/sk-PC and pDHt/sk-CC bear the coding sequence of the hph gene conferring hygromycin resistance. 6. Alternatively, the binary vectors pDHt/sk-PC and pDHt/skCC can be introduced into Agrobacterium following the protocol published by Michielse et al. [28]. 7. Alternatively, pDHt/sk-PC and pDHt/sk-CC can be introduced into T. reesei directly by protoplast-based transformation. Compared with protoplast transformation, Agrobacterium-mediated transformant is much easier to manipulate. However, protoplast transformation allows introducing more genetic materials (including RNAs and proteins) into cells than just DNA. 8. Do not thaw and use an aliquot more than twice, as activity of AS decreases during thawing and freezing. AS is degraded by light. 9. The 20-bp protospacer targeting T. reesei ura5 (50 -GGCGAGGGCGGCAACATCGT-30 ) is included in the forward primer. It can be substituted by other protospacer sequences when targeting other genes. 10. Conidiospores can be replaced by protoplasts or mycelium. However, conidiospores are very easy to obtain and have high efficiency in Agrobacterium-mediated transformation. 11. Besides the T7 promoter, the SP6 and the T3 promoter are available for transcription of gRNA in vitro. 12. If the T7 promoter is used, the 20-bp target sequence should start from at least two guanines at the 50 -end (see Fig. 1). 13. The template DNA of gRNA including the T7 promoter, the 20-bp target sequence, the crRNA, the tracrRNA, and a polyT terminator can be inserted into a vector to obtain enough template by progeny in E. coli. 14. The total reaction volume is extended to 10 μL. This guarantees that enough gRNA required for protoplast transformation will be obtained. 15. Incubation time is extended to overnight due to the large reaction volume. 16. While removing DNA by DNaseI, 0.5 μL of product can be diluted to a series of concentrations (50 to 250) and detected by gel electrophoresis. 17. Cas9-induced double-strand break repair mainly involves non-homologous end joining (NHEJ) and homology directed repair (HDR). For stimulating NHEJ, transformation of gRNA without donor DNA will cause mutation in protospacer and PAM sequence. For stimulating HDR, transformation of gRNA together with donor DNA (homologous flanks longer

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than 200 bp) will replace the targeted endogenous gene. 1–5 μg of donor DNA is required for protoplast transformation to stimulate HDR with gRNA. 18. For simultaneous transformation of multiple gRNAs, the volumes of solution II and solution III can be increased accordingly. 19. For the stimulation of HDR, the selection marker gene can be included into donor DNA. 20. Protoplasts are very fragile. Pipe the protoplast suspension gently along the tube wall to reduce the impact. 21. Incubation time can be adjusted according to selection markers used. 22. PDA can replace RM to shorten incubation time. References 1. Haft DH, Selengut J, Mongodin EF, Nelson KE (2005) A guild of 45 CRISPR-associated (Cas) protein families and multiple CRISPR/ Cas subtypes exist in prokaryotic genomes. PLoS Comput Biol 1:e600 2. Bru¨ggemann H, Lomholt HB, Tettelin H, Kilian M (2012) CRISPR/cas loci of type II Propionibacterium acnes confer immunity against acquisition of mobile elements present in type I P. acnes. PLoS One 7:e34171 3. Nakayama T, Fish MB, Fisher M, OomenHajagos J, Thomsen GH, Grainger RM (2013) Simple and efficient CRISPR/Cas9mediated targeted mutagenesis in Xenopus tropicalis. Genesis 51:835–843 4. Cho SW, Kim S, Kim JM, Kim JS (2013) Targeted genome engineering in human cells with the Cas9 RNA-guided endonuclease. Nat Biotechnol 31:230–232 5. DiCarlo JE, Norville JE, Mali P, Rios X, Aach J, Church GM (2013) Genome engineering in Saccharomyces cerevisiae using CRISPR-Cas systems. Nucleic Acids Res 41:4336–4343 6. Wijshake T, Baker DJ, van de Sluis B (2014) Endonucleases: new tools to edit the mouse genome. Biochim Biophys Acta 1842:1942–1950 7. Wang T, Wei JJ, Sabatini DM, Lander ES (2014) Genetic screens in human cells using the CRISPR-Cas9 system. Science 343:80–84 8. Ota S, Hisano Y, Ikawa Y, Kawahara A (2014) Multiple genome modifications by the CRISPR/Cas9 system in zebrafish. Genes Cells 19:555–564 9. Nødvig CS, Hoof JB, Kogle ME, Jarczynska ZD, Lehmbeck J, Klitgaard DK, Mortensen

UH (2018) Efficient oligo nucleotide mediated CRISPR-Cas9 gene editing in Aspergilli. Fungal Genet Biol 115:78–89 10. Punt PJ, van Biezen N, Conesa A, Albers A, Mangnus J, van den Hondel C (2002) Filamentous fungi as cell factories for heterologous protein production. Trends Biotechnol 20:200–206 11. Zhong Y, Liu X, Xiao P, Wei S, Wang T (2011) Expression and secretion of the human erythropoietin using an optimized cbh1 promoter and the native CBH I signal sequence in the industrial fungus Trichoderma reesei. Appl Biochem Biotechnol 165:1169–1177 12. Stricker AR, Steiger MG, Mach RL (2007) Xyr1 receives the lactose induction signal and regulates lactose metabolism in Hypocrea jecorina. FEBS Lett 581:3915–3920 13. Weninger A, Hatzl AM, Schmid C, Vogl T, Glieder A (2016) Combinatorial optimization of CRISPR/Cas9 expression enables precision genome engineering in the methylotrophic yeast Pichia pastoris. J Biotechnol 235:139–149 14. Derntl C, Rassinger A, Srebotnik E, Mach RL, Mach-Aigner AR (2016) Identification of the main regulator responsible for synthesis of the typical yellow pigment produced by Trichoderma reesei. Appl Environ Microbiol 82:6247–6257 15. Nødvig CS, Nielsen JB, Kogle ME, Mortensen UH (2015) A CRISPR-Cas9 system for genetic engineering of filamentous fungi. PLoS One 10:e0133085 16. Liu R, Chen L, Jiang YP, Zhou ZH, Zou G (2015) Efficient genome editing in filamentous

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fungus Trichoderma reesei using the CRISPR/ Cas9 system. Cell Discov 1:15007 17. Sarkari P, Marx H, Blumhoff ML, Mattanovich D, Sauer M, Steiger MG (2017) An efficient tool for metabolic pathway construction and gene integration for Aspergillus niger. Bioresour Technol 245:1327–1333 18. Matsu-ura T, Baek M, Kwon J, Hong C (2015) Efficient gene editing in Neurospora crassa with CRISPR technology. Fungal Biol Biotechnol 2:4 19. Jacobs JZ, Ciccaglione KM, Tournier V, Zaratiegui M (2014) Implementation of the CRISPR-Cas9 system in fission yeast. Nat Commun 5:5344 20. Enkler L, Richer D, Marchand AL, Ferrandon D, Jossinet F (2016) Genome engineering in the yeast pathogen Candida glabrata using the CRISPR-Cas9 system. Sci Rep 6:35766 21. Pohl C, Kiel JA, Driessen AJ, Bovenberg RA, Nyga˚rd Y (2016) CRISPR/Cas9 based genome editing of Penicillium chrysogenum. ACS Synth Biol 5:754–764 22. Arazoe T, Miyoshi K, Yamato T, Ogawa T, Ohsato S, Arie T, Kuwata S (2015) Tailormade CRISPR/Cas system for highly efficient targeted gene replacement in the rice blast fungus. Biotechnol Bioeng 112:2543–2549 23. Canzler S, Stadler PF, Hertel J (2016) U6 snRNA intron insertion occurred multiple times during fungi evolution. RNA Biol 13:119–127 24. Nissim L, Perli SD, Fridkin A, Perez-Pinera P, Lu TK (2014) Multiplexed and programmable regulation of gene networks with an integrated RNA and CRISPR/Cas toolkit in human cells. Mol Cell 54:698–710

25. Schwartz CM, Hussain MS, Blenner M, Wheeldon I (2016) Synthetic RNA polymerase III promoters facilitate high-efficiency CRISPR-Cas9-mediated menome editing in Yarrowia lipolytica. ACS Synth Biol 5:356–359 26. Zheng X, Zheng P, Zhang K, Cairns TC, Meyer V, Sun J, Ma Y (2019) 5S rRNA promoter for guide RNA expression enabled highly efficient CRISPR/Cas9 genome editing in Aspergillus niger. ACS Synth Biol 8:1568–1574 27. Al Abdallah Q, Ge W, Fortwendel JR (2017) A simple and universal system for gene manipulation in Aspergillus fumigatus: in vitro-assembled Cas9-guide RNA ribonucleoproteins coupled with microhomology repair templates. mSphere 2:e00446–e00417 28. Ram AFJ, Michielse CB, Hooykaas PJJ, van den Hondel CAMJJ (2008) Agrobacteriummediated transformation of the filamentous fungus Aspergillus awamori. Nat Protoc 3:1671–1678 29. Schiml S, Fauser F, Puchta H (2014) The CRISPR/Cas system can be used as nuclease for in planta gene targeting and as paired nickases for directed mutagenesis in Arabidopsis resulting in heritable progeny. Plant J 80:1139–1150 30. Shan Q, Wang Y, Li J, Zhang Y, Chen K, Liang Z, Zhang K, Liu J, Xi JJ, Qiu JL, Gao C (2013) Targeted genome modification of crop plants using a CRISPR-Cas system. Nat Biotechnol 31:686–688 31. Avis JM, Conn GL, Walker SC (2012) Cis-acting ribozymes for the production of RNA in vitro transcripts with defined 5’ and 3’ ends. Methods Mol Biol 941:83–98

Chapter 9 The Copper-Controlled RNA Interference System in Trichoderma reesei Lei Wang, Weixin Zhang, Xiangfeng Meng, and Weifeng Liu Abstract Trichoderma reesei is capable of secreting large amounts of lignocellulose-degrading enzymes. Although the genome sequence of T. reesei has been available, the molecular mechanisms of the hyper-production of cellulases, including the transcriptional regulation and the protein secretion, have not been completely elucidated yet. This is partially due to the lack of genetic manipulation approaches. RNA interference (RNAi) is a powerful tool for functional genomic studies in eukaryotes. Some successful examples of RNAi have already been reported; however, these systems were either uncontrolled or relied on a nutrient source inducible promoter. Here, we present a copper-controlled RNAi system in T. reesei for reversible silencing of different target genes. As the proof of concept, T.reesei xyr1, the key transcriptional activator of cellulase genes, has been knocked down using this method. Key words RNA interference, tcu1, Copper, Knockdown, Reversible, T. reesei

1

Introduction Although the genomic sequence of T. reesei is now available, the function of a large proportion of genes in the genome remain uncharacterized due to limited and intricate genetic manipulation approaches [1]. The function of particular genes in T. reesei is often studied by homologous recombination (HR)-based gene knockout [2, 3]. However, the HR frequency in T. reesei is inherently very low. In order to ensure the occurrence of HR in T. reesei, laborintensive constructions of plasmids with long homologous arms (usually >1 kb) on both sides of the target gene are required [4]. Even with these efforts, large amounts of screening are still needed to obtain the appropriate knockout strain. Although deleting particular genes in the nonhomologous end joining (NHEJ) pathway has been shown to increase HR in T. reesei [4, 5], side effects may occur from elimination of these important gene guarding genome integrity. Furthermore, charactering the function of essential genes is difficult considering that a large proportion of

Astrid R. Mach-Aigner and Roland Martzy (eds.), Trichoderma reesei: Methods and Protocols, Methods in Molecular Biology, vol. 2234, https://doi.org/10.1007/978-1-0716-1048-0_9, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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them are proven to be intractably knocked out or their deletion causes severe growth defects or even lethal, thus preventing further phenotypic characterization [6]. The post-transcriptional gene silencing by small interfering RNA (siRNA)-mediated RNAi has been shown to be a powerful tool for functional genomic studies in eukaryotes. On the other hand, tunable promoters controlled by external conditions have been shown to be useful for investigating the function of target genes in T. reesei [6, 7]. Here, we present the experimental approaches of a developed copper-controlled RNAi-mediated knockdown system in T. reesei by combining the previously identified copper-responsive Ptcu1 promoter with the routine RNAi vector for reversible silencing of different target genes. Importantly, the characterization of target gene function is facilitated by comparing the phenotype in the medium with copper (mimicking the wild-type) and without copper (gene knocking down) [8].

2

Materials

2.1

Strains

2.2

Plasmids

Escherichia coli DH5α was used for plasmid construction and was routinely cultured in Luria broth (LB) with a rotary shaker at 37
C and 200 rpm. T. reesei QM9414 ATCC 26921 was used in this work as parental strain. 1. pRLMex30 [9]. 2. T-Vector pMD™ 19 (Simple) (Takara, Tokyo, Japan).

2.3 Sequences of Main DNA Elements Used in the Construction of pKD-hph 2.3.1 Promoter of tcu1

CTGTGTGGCATCACTCATGTTCTGGATGTGCCGAGCACG CACTATAGAGTGAGCGCCAGCCATGGCGTTGCGCATTTG GTCTCTTGATAGAAAAGGCAGGACGGGTGACTCTGTGCC AGATGGTTGCGGCGCCGAGTGTTGGTGATTGGCAGCGT TATGTTGCAAAGGAGCTTTATCGGGCAGCACAGAGAACT TAATATAAGCATGACGAGCAGCCAGATAAGTTCAATACCC AAGAAATTCTAGGGCTCTAGTTATGTTGTCCACTTGGGA GGCTTGGAGGGATGGTCCCCCCCCCCCCCCTTAAAGTA ATTCGCGGCTGTTGTATCGGTGCGCTTTATCCGCCAAC GGTTTTGATAGTTGCACTTGTATCTGCTGGAATGCTTCC ACAATGCTGTAGTCAGTGATACAAGGTTAATGATGTCGC TTGATGCGGATTCTTGAAGGTGGGGAACTACTGCAGGT GAGGACATATGAGCAAGTTTGCGGGAGATTATGAAGATC GCAGGTTGGCTGACTTGACCACCCTTATCAGGCACATTT GATTCGGCCTATATATGTGTGAGATTTATCTCAGCAATGC TCAGACAACTCCTCTGAATGGTTTGGCGATTTTGCTTTGG CTTGATCATCGCTTGTCTTTGCAGTCACCCTACTTCACA GGACTCCCAAGGAGTGCATCAATCCACAAGAGCCTACT GCCAAATCCAGGAGTTCAATCTATACGACCTGGTTGATAC GACA.

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2.3.2 Intron of cel5a

GTGAGTACCCTTGTTTCCTGGTGTTGCTGGCTGAAAAGT TGGGCGGGTATACAGCGATGCGGACTGCAAGAACACCG CCGGTCCGCCACCATCAAGATGTGGGTGGTAAGCGGCG GTGTTTTGTACAACTACCTGACAGCTCACTCAGGAACTG AGAATTAATGGAAGTCTTGTTACAG.

2.3.3 Terminator of cel6a

GGCTTTCGTGACCGGGCTTCAAACAATGATGTGCGATGG TGTGGTTCCCGGTTGGCGGAGTCTTTGTCTACTTTGGTT GTCTGTCGCAGGTCGGTAGACCGCAAATGAGCAACTGAT GGATTGTTGCCAGCGATACTATAATTCACATGGATGGTCT TTGTCGATCAGTAGCTAGTGAGAGAGAGAGAACATCTATC CACAATGTCGAGTGTCTATTAGACATACTCCGAGAATAAA GTCAACTGTGTCTGTGATCTAAAGATCGATTCGGCAGTCG AGTAGCGTATAACAACTCCGAGTACCAGCAAAAGCACGTC GTGACAGGAGCAGGGCTTTGCCAACTGCGCAACCTTGCT TGAATGAGGATACACGGGGTGCAACATGGCTGTACTGAT CCATCGCAACCAAAATTTCTGTTTATAGATCAAGCTGGTA GATTCCAATTACTCCACCTCTTGCGCTTCTCCATGACATG TAAGTGCACGTGGAAACCATACCCAAATTGCCTACAGCTG CGGAGCATGAGCCTATGGCGATCAGTCTGGTCATGTTAA CCAGCCTGTGCTCTGACGTTAATGCAGAATAGAAAGCCG CGGTTGCAATGCAAATGATGATGCCTTTGCAGAAATGGCT TGCTCGCTGACTGATACCAGTAACAACTTTGCTTGGCCGT CTAGCGCTGTTGATTGTATTCATCACAACCTCGTCTCCCT CCTTTGGGTTGAGCTCTTTGGATGGCTTTCCAAACGTTAA TAGCGCGTTTTTCTCCACAAAGTATTCGTATGGACGCGC TTTTGCGTGTATTGCGTGAGCTACCAGCAGCCCAATTGG CGAAGTCTTGAGCCGCATCGCATAGAATAATTGATTGCGC ATTTGATGCGATTTTTGAGCGGCTGTTTCAGGCGACATTT CGCCCGCCCTTATTTGCTCCATTATATCATCGACGGCATG TCCAATAGCCCGGTGATAGTCTTGTCGAATATGGCTGTCG TGGATAACCCATCGGCAGCAGATGATAATGATTCCGCAGC AC AAGCTCGT.

2.4 T. reesei DNA Transformation

1. Restriction enzyme SspI. 2. Minimal medium (MM): Add about 500 mL of deionized water to a 1 L glass beaker. Weigh 20 g of glucose, 5 g of (NH4)2SO4, 15 g of KH2PO4, and 10 g of peptone to the glass beaker. Mix and adjust the pH with NaOH to 5.5. Make up to 1 L with deionized water. Autoclave and store at room temperature (see Note 1). 3. Regeneration medium: Add about 500 mL of deionized water to a 1 L glass beaker. Weigh 182.17 g of sorbitol, 20 g of glucose, 5 g of (NH4)2SO4, and 15 g of KH2PO4 to the glass beaker. Mix and adjust the pH with NaOH to 5.5. Make up to 1 L with deionized water. Aliquot 250 mL per Erlenmeyer flask and add 3.75 g of agar A. Autoclave and store at room temperature (see Notes 1 and 2).

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4. Screening medium: Add about 500 mL of deionized water to a 1 L glass beaker. Weigh 20 g of glucose, 5 g of (NH4)2SO4, and 15 g of KH2PO4 to the glass beaker. Mix and adjust the pH with NaOH to 5.5. Make up to 1 L with deionized water. Aliquot 250 mL per Erlenmeyer flask and add 3.75 g of agar A. Autoclave and store at room temperature (see Notes 1–3). 5. MEA (malt extract agar) medium: Add about 100 mL of deionized water to a 500 mL Erlenmeyer flask. Weigh 9 g of malt extract, 1.5 g of peptone, and 4.5 g of agar A to the Erlenmeyer flask. Make up to 300 mL with deionized water. Mix and heat to dissolve agar A. Aliquot 4 mL of MEA medium per glass tube and autoclave to make agar slant. Store at room temperature. 6. SK buffer: Add about 500 mL of deionized water to a 1 L glass beaker. Weigh 218.6 g of sorbitol, 1.251 g of KH2PO4, and 0.182 g of K2HPO4 to the glass beaker. Make up to 1 L with deionized water and mix. Autoclave and store at room temperature. 7. STC buffer: Add about 500 mL of 10 mM Tris–HCl, pH 7.5 to a 1 L glass beaker. Weigh 218.6 g of sorbitol and 5.54 g of CaCl2 to the glass beaker. Make up to 1 L with 10 mM Tris– HCl, pH 7.5 and mix. Autoclave and store at room temperature. 8. PTC buffer: Add about 30 mL of 10 mM Tris–HCl, pH 7.5 to a 100 mL glass beaker. Weigh 60 g of PEG 4000 and 0.554 g of CaCl2 to the glass beaker. Make up to 100 mL with 10 mM Tris–HCl, pH 7.5 and mix. Autoclave and store at room temperature (see Note 4). 9. Snailase: Store at 4
C. 10. Lysing enzymes from Trichoderma harzianum: store at 4
C. 11. 200 MgSO4: Add about 100 mL of deionized water to a 500 mL glass beaker. Weigh 30 g of MgSO4∙7H2O to the glass beaker. Make up to 250 mL with deionized water and mix. Autoclave and store at room temperature. 12. 200 CaCl2: Add about 100 mL of deionized water to a 500 mL glass beaker. Weigh 30 g of CaCl2 to the glass beaker and make up to 250 mL with deionized water. Add a few drops of HCl and mix (see Note 5). Autoclave and store at room temperature. 13. 1000 trace elements: Add about 50 mL of deionized water to a 100 mL glass beaker. Weigh 0.5 g of FeSO4∙7H2O, 0.16 g of MnSO4∙H2O, 0.14 g of ZnSO4∙7H2O, and 0.2 g of CoCl2 to the glass beaker and make up to 100 mL with deionized water. Mix and make it sterile by filtering through 0.22 μm filter membrane. Store at room temperature.

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14. 1000 CuSO4: Add about 50 mL of deionized water to a 100 mL glass beaker. Weigh 0.5 g of CuSO4∙5H2O to the glass beaker and make up to 100 mL with deionized water. Mix and make it sterile by filtering through 0.22 μm filter membrane. Store at room temperature. 15. 100 Triton X-100: Add about 50 mL of deionized water to a 100 mL glass beaker. Transfer 10 mL of Triton X-100 to the glass beaker. Make up to 100 mL with deionized water. Autoclave and store at room temperature. 16. Sterile sand core funnels G1/G2. 17. 40% (v/v) glycerol: Add 80 mL of glycerol to a 250 mL glass beaker and make up to 200 mL with deionized water. Autoclave and store at room temperature. 2.5 Fermentation of T. reesei

1. Mandels–Andreotti (MA) medium: Add about 800 mL of deionized water to a 1 L glass beaker. Weigh 17.907 g of Na2HPO4∙12 H2O, 1.4 g of (NH4)2SO4, 2.0 g of KH2PO4, and 0.3 g of urea to the glass beaker. Add 0.5 mL of Tween-80 to the glass beaker. Mix and adjust the pH to 5.0 with anhydrous citric acid. Make up to 1 L with deionized water. Use 1% (v/v) glycerol as carbon source and 2 g/L peptone for pre-culture. Use 1% (w/v) Avicel as carbon source for the induction of cellulase expression (see Notes 1 and 6).

2.6 Dodecyl Sulfate, Sodium Salt (SDS)– Polyacrylamide Gel Electrophoresis (SDS– PAGE)

1. Mini-PROTEAN Tetra Cell (Bio-Rad Laboratories, Hercules, CA, USA). 2. 10% SDS: Add about 700 mL of deionized water to a 1 L glass beaker. Weigh 100 g of SDS to the beaker. Add deionized water to a volume of 1 L. Mix and store at room temperature (see Note 7). 3. 10 Tris-glycine buffer: Add about 500 mL of deionized water to a 1 L glass beaker. Weigh 30.3 g of Tris and 144 g of glycine to the glass beaker, mix, and make it up to 1 L with deionized water. Dilute 100 mL of 10 Tris–glycine buffer to 990 mL with deionized water and add 10 mL of 10% SDS as running buffer. 4. 1.5 M Tris–HCl buffer, pH 8.8: Add about 700 mL of water to a 1 L glass beaker. Weigh 181.7 g of Tris and transfer to the beaker. Mix and adjust pH with HCl to 8.8. Add deionized water to a volume of 1 L. 5. 1 M Tris–HCl buffer, pH 6.8: Add about 500 mL of water to a 1 L glass beaker. Weigh 121.2 g of Tris and transfer to the beaker. Mix and adjust pH with HCl to 6.8. Add deionized water to a volume of 1 L.

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6. 10% ammonium persulfate (APS): Add about 5 mL of water to a 15 mL centrifuge tube. Weigh 1 g of ammonium persulfate and transfer to the tube. Add water to a volume of 10 mL. Mix and store at 4
C (see Note 8). 7. 30% acryl/bis solution. 8. 5 SDS loading buffer: Add 3 mL of 1 M Tris–HCl buffer, pH 6.8 to a 15 mL centrifuge tube. Add 4.5 mL of glycerol, 2 mL of 10% SDS, 0.5 g of β-mercaptoethanol, and 10 mg of bromophenol blue to the tube orderly. Add water to a volume of 10 mL. Mix and store at room temperature. 9. Coomassie blue staining solution: Add 450 mL of ethanol and 100 mL of acetic acid to a 1 L glass beaker. Weigh 1 g of Coomassie blue R-250 and transfer to the beaker. Add water to a volume of 1 L. Mix and filter. Store at room temperature. 10. Destaining solution: Add 100 mL of ethanol, 100 mL of acetic acid, and 800 mL of water to a 1 L glass beaker. Mix and store at room temperature. 2.7

Commercial Kits

1. Fungal DNA Kit D3390 (Omega, Doraville, GA, USA). 2. TRIzol™ Plus RNA Purification Kit (Invitrogen, Carlsbad, CA, USA). 3. PrimeScript RT Reagent Kit (Takara, Tokyo, Japan). 4. SYBR Green Supermix (Takara, Tokyo, Japan). 5. TURBO DNA-free Kit (Invitrogen, Carlsbad, CA, USA).

3

Methods

3.1 Design of the Copper-Responsive RNAi System

We previously identified the tcu1 gene encoding a copper transporter in T. reesei and showed that its expression is tightly controlled by the external copper levels. Specifically, the tcu1 gene or specific genes driven by the Ptcu1 promoter are highly transcribed in native conditions while their transcription is repressed if the external copper levels are above 500 nM [7]. The RNAi-mediated silencing system was proven to be an effective strategy for target gene knockdown in eukaryotes. We combined the controllability of the Ptcu1 promoter with the RNAi-mediated silencing system in T. reesei. As shown in Fig. 1, the gene-specific sequence (see Note 9) is cloned as a tandem inverted repeat with the insertion of an intron from the cel5a gene in between to build the synthetic hairpin RNA (hpRNA) expression cassette, which is then inserted downstream of the Ptcu1 promoter. The transcription of the hpRNA fragments for target genes will result in the formation of double-stranded RNA that is digested by the Dicer nuclease with the formation of siRNA to induce degradation of target mRNAs. In the present strategy, the target genes are expected to be downregulated when no copper

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Fig. 1 The main features and restriction sites used for the pKD-hph vector construction. Ptcu1 and Tcel6a were employed as promoter and terminator, respectively, for the expression of RNAi fragments. Between Ptcu1 and Tcel6a, a cel5a intron was inserted (a). The forward DNA fragment of a particular gene and its reverse complemented fragments were inserted into the pKD-hph vector using EcoRV-KpnI and SpeI-NotI, respectively. Therefore, the Ptcu1 promoter drives the transcription of hpRNA fragments, which can be processed by Dicer into siRNA and then directs the degradation of the target gene mRNA by the RNAi machinery. The transcription of the RNAi fragment can be turned off when copper is added to the medium (b) (reproduced from ref. 8 with permission from BioMed Central)

ions are included in the medium. Silencing can be reversed by adding copper ions into the medium, thus mimicking gene complementation (see Fig. 1a, b). 3.2 Construction of the pKD-hph-xyr1 Plasmid

1. Amplify the expression cassette of the hygromycin resistance gene from pRLMex30 and insert it into the commercial T-Vector pMD™ 19 (Simple) plasmid, resulting in pMD19Thph. 2. Amplify the 744-bp Ptcu1 promoter, 1041-bp Tcel6a terminator, and 179-bp Icel5a intron by PCR from T. reesei genomic DNA and orderly insert into pMD19T-hph to generate pKD-hph (see Fig. 1a). The sequences of these DNA elements are supplied in Subheading 2.3. 3. Amplify the 788-bp xyr1 cDNA sequence and its reverse complemented fragment by PCR from the total cDNA of Avicelinduced T. reesei and insert them into the pKD-hph plasmid at EcoRV-KpnI and SpeI-NotI sites, respectively, to construct pKD-hph-xyr1 (see Fig. 1b).

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3.3 T. reesei Transformation of the pKD-hph–xyr1 Plasmid

1. Linearize the pKD-hph–xyr1 plasmid with SspI. 2. Inoculate 5  107 spores of T. reesei QM9414 in 100 mL of MM medium and cultivate at 30
C for 24 h. 3. Collect hyphae using a sterile sand core funnel G1 and wash twice with sterile deionized water. 4. Put 5–10 g of hyphae (wet weight) into a 100 mL Erlenmeyer flask and add 8 mL of SK buffer. 5. Add 0.08 g of snailase and lysing enzymes, respectively. Digest for 2–3 h at 30
C with shaking at 200 rpm (see Note 10). 6. Filter the lysate with a sterile sand core funnel G2 and collect the filtrate to a 10 mL sterile tube. 7. Collect protoplasts by centrifugation at 4
C and 4000  g for 10 min. 8. Discard the supernatant, add 4 mL of pre-cooled STC buffer to wash the protoplasts and centrifuge at 4
C with 4000  g for 10 min. Repeat the washing step once. 9. Discard the supernatant, resuspend protoplasts with 100 μL of STC buffer. 10. Add 10 μg of linearized pKD-hph-xyr1 plasmid and 25 μL of PTC buffer to protoplast solution. Mix gently and place on ice for 20 min. 11. Add 1 mL of PTC buffer, mix gently, and incubate at room temperature for 5 min. 12. Add 2 mL of STC buffer and mix gently. 13. Spread 300 μL of the transformed protoplast solution on regeneration medium agar plates and incubate at 30
C for several days (see Notes 1 and 11). 14. Pick the transformants and transfer them to screening medium agar plates (see Note 1). Incubate at 30
C for 1–2 days. 15. Transfer the well-grown hyphae onto MEA slants and incubate at 30
C for 3–5 days for sporulation (see Note 12). 16. Collect conidia using 40% glycerol. 17. Dilute conidia of step 16 using sterile water and spread 100 μL of an appropriate dilution onto screening medium agar plates and cultivate at 30
C for 3 days. 18. Put a single colony onto an MEA slant and incubate at 30
C for 3–5 days (see Note 12). 19. Repeat steps 16–18 at least once. 20. Collect conidia using 40% glycerol and store at 80
C (see Note 13). 21. Inoculate about 1  107 conidia into 5 mL of MM medium (see Note 1), cultivate at 30
C and 200 rpm for 2 days.

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Fig. 2 The correct integration event of the hpRNA expression cassette was verified by genomic PCR using four primers (a). DNA gel electrophoresis of PCR products showed the amplification of the DNA fragments with the expected size for the correct RNAi transformants (b) (reproduced from ref. 8 with permission from BioMed Central)

22. Extract T. reesei genomic DNA of candidate RNAi transformants using Fungal DNA Kit D3390 (see Note 14). 23. Confirm the correct integration of the hpRNA expression cassette by amplifying it using indicated primers with genomic DNA as template (see Fig. 2a, b) (see Note 15). 3.4 Analyze the Phenotype of RNAi Transformants

1. Inoculate the conidia of two to three correct RNAi transformants into MA medium (see Notes 1 and 6) for preculture at 30
C and 200 rpm for 48 h. The parental strain QM9414 is used for control. 2. Transfer an equal amount of hyphae (about 3–5 g of wet weight) into induction medium containing 1% Avicel (see Notes 1 and 6) and cultivate at 30
C with shaking at 200 rpm. 3. Collect hyphae at 6 h of induction and extract the RNA using the TRIzol™ Plus RNA Purification Kit (see Note 14). 4. Remove the genomic DNA from the isolated total RNA using the TURBO DNA-free Kit (see Note 14). 5. Perform reverse transcription using the PrimeScript RT Reagent Kit (see Note 14). 6. Perform quantitative PCR using the SYBR Green Supermix (see Note 14). 7. Analyze data using the 2ΔΔCT method (see Note 16). Use the endogenous actin gene as the control for normalization (see Fig. 3). The value of the Y-axis represents the calculation result of 2ΔΔCT. The computation of ΔΔCT was performed as follows: ΔΔCT ¼ (CTxyr1  CTactin)6h  Q(CTxyr1  CTactin)0h, where Q represents the sample of parental strain QM9414 cultured on glycerol without copper.

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Fig. 3 qRT-PCR analysis of intracellular mRNA levels of the endogenous xyr1 in parental and Ptcu1–xyr1KD strains performed at 6 h with or without CuSO4. A significant difference (T-test *P < 0.05) is detected for the expression of xyr1 in parental and Ptcu1–xyr1KD strains without CuSO4 (reproduced from ref. 8 with permission from BioMed Central)

8. Collect the fermentation broth every 12 h under Avicelinduced conditions, transfer 80 μL to a centrifugation tube and add 20 μL of 5 SDS loading buffer to the tube. Mix and boil for about 10 min. 9. Prepare separating gel solution by mixing 4 mL of water, 2.5 mL of 1.5 M Tris–HCl buffer, pH 8.8, 3.3 mL of 30% acryl/bis solution, 100 μL of 10% SDS, 100 μL of 10% ammonium persulfate, and 10 μL of TEMED in a 50 mL glass beaker. Cast the mixture into the gel plate and gently overlay with deionized water. 10. Prepare stacking gel solution by mixing 3.4 mL of water, 0.63 mL of 1 M Tris–HCl buffer, pH 6.8, 0.83 mL of 30% acryl/bis solution, 50 μL of 10% SDS, 50 μL of 10% ammonium persulfate, and 5 μL of TEMED in a 50 mL glass beaker. Remove the water on top of the separating gel and cast the mixture into the gel plate. Insert a 10-well gel comb immediately without introducing air bubbles. 11. When the stacking gel is solidified, remove the comb and load 20 μL of prepared sample per well. Run the electrophoresis at 120 V till the bromophenol blue dye reaches the bottom of the gel.

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Fig. 4 SDS–PAGE analysis of the total extracellular protein produced by the Ptcu1–xyr1KD strain and the parental strain QM9414, which were cultured on 1% (w/v) Avicel supplied with or without CuSO4, respectively, at the indicated time points (Reproduced from ref. 8 with permission from BioMed Central)

12. Open the gel plate, transfer the gel to glass culture dish and add 50 mL of Coomassie blue staining solution. Shake with a rotary incubator for about 1 h (see Note 17). 13. Remove the Coomassie blue staining solution, add 50 mL of destaining solution and shake overnight with a rotary incubator. 14. Take gel pictures and analyze the results (see Fig. 4).

4

Notes 1. Add 1.25 mL of 200 MgSO4, 1.25 mL of 200 CaCl2, and 250 μL of 1000 trace element solution per 250 mL of medium before use. 2. Add 600 μL of hygromycin B (50 mg/mL) into 250 mL of medium before use. 3. Add 2.5 mL of 100 Triton X-100 into 250 mL of screening medium for isolation of positive spores. 4. The PTC buffer is prone to precipitation under low temperatures. Heat to dissolve it.

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5. Add a few drops of HCl to avoid the precipitation of Ca(OH)2. 6. Add 250 μL of 1000 CuSO4 into the medium to rescue the phenotype of RNAi transformants. 7. Wear a face mask to avoid the inhalation of SDS powder through the respiratory tract. 8. The shelf life of 10% ammonium persulfate should be less than 2 weeks. 9. The minimal length of cDNA sequence used to knockdown xyr1 in our assay is 300 bp. However, we have not tried lower lengths than 300 bp, which might also work. 10. If the hyphae are not sufficiently digested, prolong the digestion appropriately. 11. We find that the time of hyphae growth in regeneration medium ranges from 3 to 6 days. 12. We find that humid medium is disadvantageous for conidia formation. Prepare the MEA slant several days before use. 13. The conidia is recommended to be stored at 80
C for longterm reservation. 14. Use the kit according to the manufacturer’s instructions. 15. We find the hpRNA expression cassette is not efficiently amplified using primer pairs F1/R2. 16. The quantitative transcript analysis of xyr1 using primers designed in different regions of the xyr1 cDNA is inconsistent although the defective phenotype for degradation of Avicel is present in RNAi transformants. We suspect that the overexpressed exogenous dsRNA (double-strand RNA) will disturb the transcript analysis of native xyr1. We recommend using a Western blot to verify the expression of target genes with unknown function. 17. The Coomassie blue staining solution can be recycled.

Acknowledgments This work is supported by Grants from the National Key R&D Program of China (2019YFA0905700), National Natural Science Foundation of China (31670040, 31970029, 31770047, 31800047, 31970071 and 31800024), Major Basic Research of Shandong Provincial Natural Science Foundation (ZR2019ZD19), Shandong Provincial Natural Science Foundation (ZR2018BC006), China Postdoctoral Science Foundation (2017M622186), Young Scholars Program of Shandong University, and the Fundamental Research Funds of Shandong University (2017TB0010).

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References 1. Martinez D, Berka RM, Henrissat B, Saloheimo M, Arvas M, Baker SE, Chapman J, Chertkov O, Coutinho PM, Cullen D, Danchin EGJ, Grigoriev IV, Harris P, Jackson M, Kubicek CP, Han CS, Ho I, Larrondo LF, de Leon AL, Magnuson JK, Merino S, Misra M, Nelson B, Putnam N, Robbertse B, Salamov AA, Schmoll M, Terry A, Thayer N, WesterholmParvinen A, Schoch CL, Yao J, Barabote R, Nelson MA, Detter C, Bruce D, Kuske CR, Xie G, Richardson P, Rokhsar DS, Lucas SM, Rubin EM, Dunn-Coleman N, Ward M, Brettin TS (2008) Genome sequencing and analysis of the biomass-degrading fungus Trichoderma reesei (syn. Hypocrea jecorina). Nat Biotechnol 26 (5):553–560. https://doi.org/10.1038/ nbt1403 2. Penttil€a M, Nevalainen H, R€atto¨ M, Salminen E, Knowles J (1987) A versatile transformation system for the cellulolytic filamentous fungus Trichoderma reesei. Gene 61(2):155–164. https:// doi.org/10.1016/0378-1119(87)90110-7 3. Hartl L, Seiboth B (2005) Sequential gene deletions in Hypocrea jecorina using a single blaster cassette. Curr Genet 48(3):204–211. https:// doi.org/10.1007/s00294-005-0011-8 4. Guangtao Z, Hartl L, Schuster A, Polak S, Schmoll M, Wang T, Seidl V, Seiboth B (2009) Gene targeting in a nonhomologous end joining deficient Hypocrea jecorina. J Biotechnol 139 (2):146–151. https://doi.org/10.1016/j. jbiotec.2008.10.007

5. Steiger MG, Vitikainen M, Uskonen P, Brunner K, Adam G, Pakula T, Penttil€a M, Saloheimo M, Mach RL, Mach-Aigner AR (2011) Transformation system for Hypocrea jecorina (Trichoderma reesei) that favors homologous integration and employs reusable bidirectionally selectable markers. Appl Environ Microbiol 77(1):114. https://doi.org/10. 1128/AEM.02100-10 6. Bischof RH, Horejs J, Metz B, Gamauf C, Kubicek CP, Seiboth B (2015) l-Methionine repressible promoters for tuneable gene expression in Trichoderma reesei. Microb Cell Factories 14 (1):120. https://doi.org/10.1186/s12934015-0308-3 7. Lv X, Zheng F, Li C, Zhang W, Chen G, Liu W (2015) Characterization of a copper responsive promoter and its mediated overexpression of the xylanase regulator 1 results in an inductionindependent production of cellulases in Trichoderma reesei. Biotechnol Biofuels 8(1):67. https://doi.org/10.1186/s13068-015-0249-4 8. Wang L, Zheng F, Zhang W, Zhong Y, Chen G, Meng X, Liu W (2018) A copper-controlled RNA interference system for reversible silencing of target genes in Trichoderma reesei. Biotechnol Biofuels 11(1):33. https://doi.org/10.1186/ s13068-018-1038-7 9. Mach RL, Schindler M, Kubicek CP (1994) Transformation of Trichoderma reesei based on hygromycin B resistance using homologous expression signals. Curr Genet 25(6):567–570. https://doi.org/10.1007/BF00351679

Chapter 10 Batch Cultivation of Trichoderma reesei Birgit Jovanovic´ Abstract This chapter explains how to perform a batch cultivation of Trichoderma reesei in bench top bioreactors, exemplarily using wheat straw as sole carbon source, and a selection of recommended, frequently used analyses to monitor the cultivation (intra- and extracellular as well), which are microscopic analysis, sodium hydroxide soluble protein, Bradford assay, and GC analysis. Key words Batch cultivation, Bioreactor, Microscopic analysis, Erythritol, Sodium hydroxide soluble protein, Bradford assay, Trichoderma reesei

1

Introduction Cultivation in shake flasks, which is frequently performed in the lab, is quite different from industrial scale production. To close this gap, one has to perform cultivation experiments in bioreactors as the first step for up-scaling. The main aspects to consider are oxygen transfer and shearing forces, cultivation duration, substrate depletion, product and byproduct aggregation. Oxygen transfer in shake flasks can be sufficiently provided by moderate shaking, which might result in an aggregation of the mycelia, but does not cause strong shearing forces. In bioreactors, on the other hand, oxygen has to be actively introduced and distributed by stirring, which can easily cause shearing forces that are capable of tearing the mycelia apart. Since sufficient oxygen supply is a very crucial factor, one will start with stirring rates as high as necessary to keep a proper oxygen level. Microscopic analysis of the mycelia during the cultivation is used to ensure proper mycelia growth is still granted. Cultivation in shake flasks can be performed as batch or fed-batch (normally manually added) and is generally quite limited in its duration. Bioreactors, in contrast, enable continuous feeding, which allows studying if high substrate concentrations, as present at the beginning of a batch, can be repressing and should be therefore

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avoided. Due to the possibility of longer cultivation times in bioreactors, one can also examine the effect of product and byproduct aggregation. Whereas product removal normally cannot be achieved in a lab-scale bioreactor, harming byproducts often can be treated, like neutralization of acids to keep proper cultivation conditions or precipitation or complexation of repressors. So there are many aspects that need to be considered before up-scaling a cultivation method. Exemplarily, we demonstrated the possibility of batch cultivation of recombinant strains of Trichoderma reesei QM6aΔtmus53 with wheat straw as sole carbon source for production of erythritol. Experiments were done in 2-L bench top bioreactors for a period of 4 days [1]. The analysis of the taken cultivation samples presented here have their focus on methods suitable to examine the overall growth and productivity of the fungus.

2

Materials Prepare all solutions using distilled water and analytical grade reagents. Prepare and store all reagents at room temperature unless indicated otherwise.

2.1

Batch Cultivation

1. 2-L Bench top bioreactors. 2. Trace elements solution: 250 mg/L iron(II) sulfate heptahydrate, 85 mg/L manganese(II) sulfate monohydrate, 70 mg/L zinc sulfate heptahydrate, and 100 mg/L calcium chloride dihydrate. 3. Pretreated straw: Wheat straw, pretreated by an alkaline organosolv process for lignin removal [2] (see Notes 1 and 2). 4. Cultivation medium: 3.5 g/L ammonium sulfate, 5 g/L potassium dihydrogen phosphate, 1.25 g/L magnesium sulfate heptahydrate, 0.625 g/L sodium chloride, 1.25 g/L peptone from casein, and 1.5 mL/l trace elements solution. Always prepare fresh medium prior to use. 5. Tween® 80. 6. Antifoam Y-30 Emulsion. 7. Syringe (sterile), 30 mL. 8. Air filter. 9. Potter-Elvehjem tissue grinder. 10. Bright field microscope.

2.2 Sodium Hydroxide Soluble Protein

1. 0.1 M Sodium hydroxide solution. 2. Sonifier® 250 Cell Disruptor (e.g., Branson, Danbury, CT, USA). 3. Centrifuge.

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Bradford Assay

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1. Bradford reagent, diluted 1:5. Prepare fresh before use. 2. Bovine serum albumin (BSA) standards: 10 μg/mL, 20 μg/ mL, 50 μg/mL, 100 μg/mL (store at 4  C). 3. UV-Vis spectrophotometer.

2.4 Gas Chromatography

1. Sonifier® 250 Cell Disruptor (e.g., Branson, Danbury, CT, USA). 2. Centrifuge. 3. Myo-inositol. 4. 96% ethanol. 5. Pyridine. 6. Hexamethyldisilazane. 7. Trimethylsilyl chloride. 8. GC equipment (e.g., Agilent Technologies, Santa Clara, CA, USA). 9. HP-5-column (30 m, inner diameter 0.32 mm, film 0.26 μm) (Agilent).

3 3.1

Methods Batch Cultivation

1. Prepare 1.3 L of medium per bioreactor, aliquot in bioreactor vessels, and add pretreated straw (3 g/L), Tween® 80 (0.5 mL/L) and Antifoam Y-30 Emulsion (1 mL/ bioreactor). 2. Equip bioreactor with probes for temperature and oxygen measurement, filter for outgoing air, sample taking pipe, and autoclave (see Note 3). 3. Connect bioreactor to station and adjust stirring rate to 500 rpm, temperature to 28  C, and aeration rate to 0.5 vvm. 4. Inoculate with 109 conidia per liter. 5. Draw a full syringe of cultivation broth (30–35 mL) every 12 h (see Note 4). One drop of the sample is used for microscopic analysis, 20 mL is thoroughly decomposed with PotterElvehjem tissue grinder, 10 mL is used for the analysis of mycelia and medium. 6. At the end of the batch cultivation, measure the amount of the remaining cultivation broth and siphon it to retrieve the insoluble compounds for dry mass determination.

3.2 Microscopic Analysis

Examination of cultivation broth under a microscope is useful for monitoring the state of the cultivation. The order of observations is as follows:

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1. Conidia They should be only found in reasonable number shortly after incubation. If there are still a lot of conidia aside fully grown mycelia, then too much conidia were taken for inoculation. 2. Germinating mycelium Few mycelium with short, unbranched filaments, maybe still attached to conidia. 3. Branching Some mycelium with longer filaments that starts to branch. 4. Fully developed mycelium A bunch of mycelium with long filaments and lots of branches. 5. Decaying mycelium Broken filaments, amount of intact mycelium and long, branched filaments are decreasing. Heavy mycelium degradation indicates the end of cultivation. If a lot of broken filaments are already found shortly after germination/branching, then sheering forces caused by stirring are too strong in the bioreactor. 3.3 Sodium Hydroxide Soluble Protein

1. To determine protein production in the whole broth (intraand extracellular) centrifuge 2 mL of the decomposed broth (20,000  g, 10 min, 4  C) and discard the supernatant. 2. Resuspend the pellet in 3 mL 0.1 M NaOH and sonicate with a Sonifier® (power 70%, duty cycle 40%, power 20 s, pause 40 s, 10 cycles, on ice). 3. Incubate 3 h at room temperature. 4. Centrifuge (20,000  g, 10 min, 4  C) and use the supernatant for the determination of protein concentration by Bradford assay.

3.4

Bradford Assay

1. Dilute sample appropriately to be within the range of BSA standards (normally 1:10 to 1:100). 2. Prepare measurement samples as follows: Add 20 μL of sample or standard to 1 mL of Bradford reagent and incubate for exactly 10 min at room temperature (see Note 5). 3. Measure absorption spectrophotometer.

3.5 Gas Chromatography

at

595

nm

on

a

UV-Vis

1. For quantitative erythritol determination in the whole cultivation broth (intra- and extracellular), sonicate 20 mL of the decomposed broth with a Sonifier® (power 70%, duty cycle 40%, power 20 s, pause 40 s, 10 cycles, on ice) (see Note 6).

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2. Centrifuge (20,000  g, 10 min, 4  C) to separate insoluble compounds. 3. Take 300 μL of the clear supernatant and supplement with 10 ng myo-inositol as internal standard. Gently mix with 1.2 mL ethanol (96%) and incubate for 30 min at room temperature for protein precipitation (see Note 7). 4. Centrifuge (20,000  g, 10 min, 4  C) and discard the precipitate. 5. Vacuum the supernatant until dry. 6. Silylate the dry remains with 50 μL of pyridine, 250 μL of hexamethyldisilazane, and 120 μL of trimethylsilyl chloride. 7. Measure by GC with a HP-5-column with mobile phase helium at a flow of 1.4 L/min. For the column, use a temperature profile as follows: 50  C for 1 min, ramping 150–220 (ΔT 4  C/min), ramping 220–320  C (ΔT 20  C/min), 320  C for 6.5 min. Detection is performed with FID at 300  C. To determine retention times, use standard substances (see Note 8).

4

Notes 1. If wet pretreated straw is used, it is recommended to shred the straw first in a little amount of cultivation medium with a blender and then transfer to the bioreactor with the rest of the medium. If dried pretreated straw is used, it can be shred dry in a chopper and then be added to the whole amount of medium. 2. Instead of pretreated straw, any other carbon source utilizable by T. reesei can be used in proper amount. 3. The bioreactor needs a while to autoclave (about 3–5 h, depending on autoclave used) and to adjust to the desired setting afterwards. Best is to let it at least a whole night stabilize its temperature before inoculation. 4. Always wear protective glasses when working with the bioreactor. The vessel can build up pressure if the outgoing air filter is layered, which will be released when taking samples. Additionally the straw can easily block the sampling pipe and make the sample burst out when the under-pressure applied by the sampling syringe suddenly breaks the clogging. 5. When doing the Bradford assay, it is really important that every sample is incubated for exactly 10 min. Use a stopwatch when adding the sample or standard to the Bradford reagent and spare the time before pipetting the next sample that it will take to make a measurement (approximately 20–30 s). Be sure to measure the samples in the same order they were prepared.

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6. If you are interested in any other substances besides erythritol occurring in the cultivation broth, just follow the sample preparation for GC analysis up to point 4 (with suitable internal standard) and use the GC or HPLC method of your choice. 7. If the amount of the substance to be analyzed by GC or HPLC is low, a considerably higher amount of sample can be used at step 3. Do not reduce the whole supernatant received in step 2 to dry since it will not re-dissolve well due to the high protein content. 8. A lot of further analyses are possible. The un-decomposed part of the sample taken from the bioreactor can be separated in mycelium and broth by vacuuming. From the mycelium, RNA may be isolated for transcript analyses by, e.g., RT-qPCR. The broth can be directly analyzed with GC or HPLC to determine any extracellular product formation. References 1. Jovanovic´ B, Mach RL, Mach-Aigner AR (2014) Erythritol production on wheat straw using Trichoderma reesei. AMB Express 4:34

2. Fackler K, Ters T, Ertl O, Messner K (2012) Method for lignin recovery. Patent WO/2012/ 027767. http://www.sumobrain.com/patents/ WO2012027767.html

Chapter 11 Image Analysis Method for the Characterization of Trichoderma reesei During Fermentations Nicolas Hardy, Maxime Moreaud, and Fadhel Ben Chaabane Abstract This chapter describes the use of a specific image analysis method with the plug im! software for the characterization of filamentous fungus morphology. It details an application of this method with samples obtained from a fermentation process of a Trichoderma reesei strain. This fully automated and accurate image analysis method provides quantitative and representative data for morphological and topological analyses. Key words Filamentous fungus, Image processing, Automating, Correlative microscopy, Trichoderma reesei

1

Introduction Performing fermentations with filamentous fungi is really complicated as the fungal morphology impacts the process conditions which in turn affect the morphology and possibly the productivity of the fungus [1]. In fact, the filamentous morphology induces an increase of the media viscosity that strongly reduces the rate of oxygen mass transfer [2]. To ensure a sufficient oxygen concentration in the broth, the power input has to be increased, which results in an increase of the shear stresses on the mycelial structures that are prone to size reduction. To better understand the complex link that exists between the process conditions and the fungus morphology, a new image analysis method is now available [3]. One of the major bottlenecks that are addressed with this method concerns data acquisition of macro- and micro-morphology characteristics of the fungi at the same time. The other major barrier that has been overcome concerns the capacity of acquiring the morphological characteristics of three-dimensional objects from two-dimensional images. Indeed, data acquisition is more complicated when a fungus with “spreadable morphology” is used as it is mainly composed of long and thin hyphae spread out over the three dimensions.

Astrid R. Mach-Aigner and Roland Martzy (eds.), Trichoderma reesei: Methods and Protocols, Methods in Molecular Biology, vol. 2234, https://doi.org/10.1007/978-1-0716-1048-0_11, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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Fig. 1 Overall description of the image analysis method used to measure morphological and typological criteria of fungi. Each slice is divided into 60 spots. In each spot, a stack of 60 big images composed of mosaics of 16 subimages is made. The 60 stacks of big images are transformed into a single big sharp image, on which segmentation and extraction of morphological and topological criteria are performed

This chapter describes the method and protocol for the characterization of the morphology of filamentous fungi. It provides quantitative and representative data based on a fully automated image analysis method using microscopy images and a new algorithm called FACE that allows sharp images to be created at all positions, segmentation of fungi, and morphological analysis using skeletons and topological approaches. This method is applied to a fungus strain of industrial interest, Trichoderma reesei (summed up in Fig. 1).

2

Materials Prepare all solutions using ultrapure water (prepared by purifying deionized water, to attain a sensitivity of 18 MΩ cm at 25
C) and analytical grade reagents. Prepare and store all reagents at room temperature (unless otherwise indicated). Diligently follow all waste disposal regulations when disposing waste materials. For sample handling, micropipettes were used with cutting tips to avoid size selection, and all solutions were mixed with a vortex to ensure homogeneity. 1. 50 mM phosphate-citrate buffer, pH 4.8. 2. Commercial lactophenol blue solution. 3. Nail varnish or paraffin. 4. Slide and cover slip. 5. Bright field microscope: In our works, we used an AxioImager M2p Carl Zeiss AG bearing an NAchroplan 20X objective (Carl Zeiss, Oberkochen, Germany). It is equipped with a motorized stage in X-Y-Z axis (steps: 0.1 μm, 0.1 μm, and 25 nm), with a color camera 5 megapixels Axicam 105 Color through a video

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adapter 60 N-C 2/300 0.5 (Carl Zeiss, Oberkochen, Germany). 6. The Axiovision software: This software can perform at least two complex tasks that are mandatory for our method: the Z-Stack (a series of images acquired at different focus positions) and the MosaiX (a series of overlapping tiled images acquired over a defined area). 7. Internet connection to download plug im!: All images are processed using plug im! software from IFPEN. plug im! is an open access platform for signal, image, and threedimensional volume processing. Its objective is to provide state-of-the-art and advanced algorithms from both industry and academia. plug im! can be downloaded and used freely for research purposes from www.plugim.fr.

3

Methods

3.1 Preparation of Samples for Image Analysis

Sampling preparation for image analysis has two goals. The first one is to isolate fungi to analyze one object per one object. The second one is to stain samples to improve the contrast between fungi and background, which facilitates the automatic image analysis process, particularly the segmentation part. For sample handling, micropipettes were used, and all solutions were mixed with a vortex to assure homogeneity and to separate distinct objects (fungi). 1. Dilute the sample taken from a flask or a bioreactor in 50 mM phosphate-citrate buffer, pH 4.8 to a final biomass concentration between 2 and 5 g/L [4] (see Note 1). 2. Mix 200 μL of the diluted samples with 800 μL of a commercial lactophenol blue solution (see Note 2). 3. Place a drop of the sample in the middle of the slide. 4. Place one side of a coverslip at an angle, so that its edge touches the slide and the outer edge of the drop, and lower the coverslip slowly avoiding the creation of air bubbles (see Note 3). 5. Seal with nail varnish or paraffin between the slide and the cover slip before microscopic observations (see Note 4).

3.2 Image Acquisition

1. Switch on the camera and the microscope and start the software Axiovision. 2. Set up the slide under the microscope and ensure that the light is directed to the camera. 3. Activate the camera controls by clicking on the relevant camera button in the acquire menu or by selecting it from the work area (see Note 5).

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4. Fix the parameters (see Note 6) by using Axiovision’s 6D-Acquisition Icon which opens a floating window: l

Open the “C00 tab: Click on “measure” and choose an exposure time of 300 μs and light intensity (190 on 255 gray levels) and validate by clicking on OK.

l

Open the “MosaiX” tab and click “Center,” choose number of columns: 4 and rows: 4, choose overlap of adjacent images: 0%.

l

Open the “XY” tab and click on “create,” then choose 10*6.

l

Open “Z-Stack”: Choose 60 for “slices” and 2 μm for slice distance.

5. Start the acquisition by giving a name to the experiment. With these parameters, the acquisition duration is around 8 h. 6. Export the file using the TIFF format. 3.3 Image Processing and Analysis

plug im! is used to process and analyze all the images. The first processing steps create filtered, sharp, and segmented images of fungi. Then, the following steps allow skeleton-based morphological analysis, as well as topological analysis. All necessary plugins, listed below, can also be downloaded from the plugins section of the website.

3.3.1 Installation

The installation steps are as follows: 1. Connect to the plug im! website https://www.plugim.fr/ and download the installer (setup.exe). 2. Install the main program through the execution file. 3. Download also the following plugin archives from the website (https://www.plugim.fr/plugin/list): l

3D volume from tiff images list [3D from list].

l

Flowing bilateral filter 3D [Flowing Bilateral 3D].

l

Fast mean Absolute difference with Confidence propagation for Extended depth of field [FACE].

l

Top Hat segmentation [TopHat segmentation].

l

Mathematical morphology basic operations [Morpho Maths].

l

Closing ends [Closing ends].

l

Border kill [Border kill].

l

Opening by criteria [Criteria opening].

l

Fungi Topological Analysis [Fungi Topo. ana.].

l

Extraction of connected components in individual files [CC extraction].

IAM to Characterize T. reesei Morphology l

Skeleton by thinning [Skeleton].

l

Skeleton analysis [Sketeton analysis].

l

Batch [Batch Processing].

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4. Install the plug im! plugins by dragging and dropping the zipped files in the plug im! main window or copy the extracted folders directly into the “Plugins” folder in the plug im! installation folder. 3.3.2 Image Processing

In this part, plug im! will generate segmented (binary) images of fungi. 1. Launch plug im! (see Fig. 2), then start the image processing by clicking on the start button (see Fig. 3). 2. Click on the open button (see Fig. 4) and select one image from your folder where a stack of images are available. The selected image from the folder appears on the screen. 3. Navigate between images using a couple of arrows up/down (see Fig. 5). You can navigate between the plugins (working with the current image) using a couple of arrows down/up (see Fig. 6) or scrolling with left-clicking.

Fig. 2 plug im! start page

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Fig. 3 plug im! start button

Fig. 4 plug im! open button

Fig. 5 Arrows up/down to navigate between images in a folder

4. Start with the plugin 3D volume from TIFF images. Select it by clicking on “3D from list.” When processing is finished (the preview), an easily understandable message continuously flashes: “Operation completed.” 5. Validate your choice by clicking on the Execute button (see Fig. 7). You can now navigate in the 3D volume with a userfriendly interface (see Fig. 8). 6. Click on the module “Flowing Bilateral 3D” (see Note 7) and set parameters (Spatial Range at 3 and Tonal Range at 5), using the buttons plus or minus, or click on the number

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Fig. 6 Arrows to navigate between plugins available for the current image

Fig. 7 plug im! execute button

corresponding to the parameter of interest and type a new value when it is flashing (see Fig. 9). 7. Validate your choice by clicking on the Execute button (see Fig. 7). 8. Use the “FACE” plugin with FACE Algorithm and Patch size parameter at 8. Wait during the processing time. Here you can play with the interface in Preview mode (see Fig. 10). 9. Validate your choice by clicking on the Execute button (see Fig. 7). You will have a single focus image (see Fig. 11). 10. Execute the “Top Hat segmentation” (see Note 8) plugin with back TopHat, Radius at 20 and Height threshold MIV (if not

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Fig. 8 “3D from list” plug im! plugin interface with synoptic for navigation at the bottom

Fig. 9 “Flowig Bilateral 3D” plug im! plugin interface with parameters tuning

all options are visible on your screen, navigate with the arrows). The shape of your fungus will appear after the execution of the plugin.

Fig. 10 “FACE” plug im! plugin interface with synoptic for dynamic view of the treatment

Fig. 11 Example of a focused image obtained with the FACE algorithm

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Fig. 12 Example of a segmented image obtained with the “Back Top Hat” and cleared with the process describe here

11. Use the Morpho Maths plugin with the “Opening” Operator, Disk and Size of 2. Remember to click on “Execute.” 12. Execute the Closing ends plugin with a distance of 15. 13. Use the Border kill plugin to erase fungi partially available on your image. 14. Clear small objects with the Criteria opening module with length as criteria and minimum value at 50. Execute it. 15. You have now an image like in Fig. 12 (see Note 9). Save in FDA format. 3.4

Image Analysis

Topological analysis will be performed by means of morphological operators and by a complete evaluation of skeletons calculated from the binary images. 1. Use the “Fungi Topo. ana.” plugin with parameters Thickness (average thickness of fungi in pixels) to 10, Hole to 50 (minimum area in pixels of a hole inside fungi), and Distance to 10 (maximum distance in pixels below which fungi are merged). You can save the preview image (see Note 10), and the result of the analysis by executing the module and by using the Export button (top right in the interface). This analysis gives number of holes, surface area, estimation of mean thickness of fungi, etc. (see Table 1 for more details).

588

4

6

190

588

352

58

35

6570

6570

657

29,063 59,039 2906

352

5

1904

0

2

0

0

0

0

0

14,988

0

0

0

0

Mean Number area of Length of holes holes

14,723 14,723 1472

1904

3

2

1

Tag Area

Filled area

911.08

1924.83

98.5

55.83

1137.58

308.75

Geodesic extreme length

820.73

1506.88

92.76

55.04

389.67

285.57

Eutectic extreme length

222.59

399.06

35.58

16.39

326.32

95.12

462.8

907.68

79.62

38

691.03

224.54

Maximum Mean radius radius of filled of filled object object

Table 1 Data extracted from the topological analysis of the image. Numbers are given in pixels

7.07

52.01

3.61

3

17

3.61

774

1382

91

55

547

270

308

999

22

9

504

98

Minimum radius Frame Frame of filled object width height IAM to Characterize T. reesei Morphology 129

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Fig. 13 Example of a fungus transformed into skeleton and analysis. Red points are tips and green are nodes

2. Reload (open) the clear and segmented image. 3. Extract data with the CC extraction plugin by selecting an output folder and executing. You will have thumbnails of all your fungi saved in the output folder. You can also save the global image with windows and tags number over all fungi. 4. Go to the output folder set in the CC extraction plugin and load an image. 5. First execute the Skeleton plugin, then the Skeleton analysis for each miniature picture. You can use the Export Button at the end to have the CSV file (.txt extension) with complete analysis of the skeleton (see Fig. 13). 3.4.1 Automatizations of plug im!

You can use the plugin Batch to chain image processing directly in plug im!. You can also use a Python script (like Pyndigo or “Characterisation merger Conv” available in a near future on the website) to parallelize instances of plug im!, save time by executing one process per core of your computer, to convert pixel in the metric system or to class fungi in Unbranched, Branched, Entangled, Clumped, or Pellet.

3.4.2 Data Handling and Analysis

Plug im! outputs are different kinds of images and data tables. These tables are CSV comma or space separated data with dots as decimal separators. In the current version of the procedure, 31 morphological criteria are extracted by fungi. To compare data, it is

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recommended to evaluate at least 1000 fungal images per sample [3]. This amount of data imposes to use automatic processes, through scripting language, for data handling, statistical analysis, and to graph data. In previous works, the Python language was used with specialized libraries: Pandas for data handling [5], SciPy for statistical analysis [6] and Matplotlib for plotting [7]. It is also possible to manually process the data by using a simple spreadsheet, e.g., in Microsoft Excel. Hardy et al. (2017) [3] retained four parameters as the most relevant in relation to size and to damage due to fluid dynamic stress. They were the surface area (mm2), the total length (mm), the hyphal growth unit length (lm), and the number of holes (nH). For each criterion, it is possible to consider data as a distribution and to synthetize distribution by quantile. For example, the 90% quantile (q90) was relevant to summarize the number of holes. It is also possible to directly compare the distribution using statistical tests like two-sample Kolmogorov–Smirnov test (KS test). It allows the largest absolute difference between the two cumulative distribution functions as a measure of disagreement to be defined [8]. A probability value ( p-value) lower than 0.05 indicates that the hypothesis of equality of the distributions of the two samples should be rejected.

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Notes 1. This dilution minimizes artefacts caused by overlapping of hyphae. 2. This staining procedure aims at stopping the growth [9], coloring healthy regions of the fungi [10]. The commercial lactophenol blue solution can be diluted with a solution of lactophenol blue without cotton blue and without phenol to adapt the staining process. The concentration can be adjusted according to the capacity of fungi to fix the dye linked to his morphology and his nature. Basically, for Trichoderma reesei CL847 [11], we use the dilution by a factor of 2. 3. Problems with air bubbles occur from not applying the coverslip at an angle, not touching the liquid drop. The liquid drop must not be too large because the coverslip will float on the slide. 4. This method allows the conservation of the sample up to 1 week without impact on image analysis (unpublished personal observation). 5. Many of the routine controls such as measuring the exposure and automatic white reference are accessible from buttons on the toolbar in the live window or clicks on the live image.

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6. With this device, 60 stacks in Z axis separated by 2 μm of 60 mosaics were recorded. Each mosaic was composed of 16 pictures with a size corresponding to 560 μm per 419 μm for a total surface of 3.7 mm2. To take and save all 57,600 pictures (60 mosaics per slide), 8 h was needed. A calibration value (0.22 μm per pixel) was set to permit the conversion of pixels into micrometers. 7. The flowing bilateral filter is a noise reduction filter that preserves image transitions and does not blur them; spatial range allows to adjust the strength of the noise reduction; total range allows to set the minimum contrast between neighbor pixels to be preserved. 8. TopHat operation is the residue between the image after a morphological opening and the image itself. The morphological opening is processed here with a disc of radius 20 pixels as a structuring element. The image is then thresholded to obtain a binary image, this threshold being calculated on histogram, here automatically by Variance Interclass Maximization. 9. With plug im! you can undo or redo image analysis steps by clicking on the buttons. It is useful when you are trying to set parameters. 10. With plug im! you can save images in different formats (BMP, PNJ, TIFF, or the plug im! format FDA) using the save button. For the PNG format, the saved file is a screenshot of the software. References 1. Hardy N, Augier F, Nienow AW, Be´al C, Ben Chaabane F (2017) Scale-up agitation criteria for Trichoderma reesei fermentation. Chem Eng Sci 172:158–168. https://doi.org/10. 1016/j.ces.2017.06.034 2. Gabelle JC, Jourdier E, Licht RB, Ben Chaabane F, Henaut I, Morchain J, Augier F (2012) Impact of rheology on the mass transfer coefficient during the growth phase of Trichoderma reesei in stirred bioreactors. Chem Eng Sci 75:408–417. https://doi.org/10.1016/j. ces.2012.03.053 3. Hardy N, Moreaud M, Guillaume D, Augier F, Nienow A, Be´al C, Ben Chaabane F (2017) Advanced digital image analysis method dedicated to the characterization of the morphology of filamentous fungus. J Microsc 266 (2):126–140. https://doi.org/10.1111/jmi. 12523 4. Lecault V, Patel N, Thibault J (2007) Morphological characterization and viability assessment of Trichoderma reesei by image analysis.

Biotechnol Prog 23(3):734–740. https://doi. org/10.1021/bp0602956 5. McKinney W (2011) pandas: a foundational Python library for data analysis and statistics. In: Python for high performance and scientific computing, 14(9) 6. Walt SVD, Colbert SC, Varoquaux G (2011) The NumPy array: a structure for efficient numerical computation. Comput Sci Eng 13 (2):22–30. https://doi.org/10.1109/MCSE. 2011.37 7. Hunter JD (2007) Matplotlib: a 2D graphics environment. Comput Sci Eng 9(3):90–95. https://doi.org/10.1109/MCSE.2007.55 8. Lopes RHC, Hobson PR, Reid ID (2008) Computationally efficient algorithms for the two-dimensional Kolmogorov–Smirnov test. J Phys Conf Ser 119(4):042019. https://doi. org/10.1088/1742-6596/119/4/042019 9. Haack MB, Olsson L, Hansen K, Eliasson Lantz A (2006) Change in hyphal morphology of Aspergillus oryzae during fed-batch

IAM to Characterize T. reesei Morphology cultivation. Appl Microbiol Biotechnol 70 (4):482–487. https://doi.org/10.1007/ s00253-005-0085-8 10. Lecault V, Patel N, Thibault J (2009) An image analysis technique to estimate the cell density and biomass concentration of Trichoderma reesei. Lett Appl Microbiol 48(4):402–407.

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https://doi.org/10.1111/j.1472-765X. 2008.02544.x 11. Durand H, Clanet M, Tiraby G (1988) Genetic improvement of Trichoderma reesei for large scale cellulase production. Enzyme Microb Technol 10(6):341–346. https://doi.org/10. 1016/0141-0229(88)90012-9

Chapter 12 Measurement of Cellulase and Xylanase Activities in Trichoderma reesei Qing-Shan Meng, Fei Zhang, Chen-Guang Liu, Feng-Wu Bai, and Xin-Qing Zhao Abstract The microbial cellulase system is responsible for the generation of glucose from cellulose. Cellulases are comprised of at least three major groups of enzymes, namely endoglucanases, exoglucanases, and β-glucosidases. On the other hand, xylanases function in the degradation of hemicellulose and work synergistically with cellulases for the degradation of lignocellulosic biomass. Here, we describe the most commonly used methods for the activity measurement of cellulases and xylanases. Key words Cellulase, Xylanase, Measurement of enzyme activity, Lignocellulosic biomass

1

Introduction Cellulases produced by various microbial strains, especially filamentous fungi, have been shown to consist of three major types of enzymes: endo-β-1,4-glucanases (EG), exo-β-1,4-glucanases (CBH), and β-glucosidases (BGL). These components act synergistically to degrade insoluble cellulosic substrates to generate glucose [1, 2]. Although cellulase activities are always determined using insoluble cellulosic substrate, the physical heterogeneity of insoluble cellulosic substrates, the enzyme accessibility of substrates, and the complexity of cellulase mixtures are challenging for enzyme activity measurements [3–6]. Cellulase activity assays normally involve two parts: one is testing the total cellulase activity, and the other is determining the activities of individual enzymes [7]. The filter paper assay (FPA) measures the total cellulose hydrolyzing capacity of microbial cellulase preparations, and the standard method was established by the International Union of Pure and Applied Chemistry (IUPAC) [8]. Reliable and reproducible data can be obtained under standardized conditions following this method. The individual activities of

Astrid R. Mach-Aigner and Roland Martzy (eds.), Trichoderma reesei: Methods and Protocols, Methods in Molecular Biology, vol. 2234, https://doi.org/10.1007/978-1-0716-1048-0_12, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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cellulase components, including EG, CBH, and BGL, are measured using sodium carboxymethyl cellulose (CMC), p-nitrophenol-Dcellobioside ( pNPC), and p-nitrophenyl-β-D-glucopyranoside ( pNPG) as the substrates, respectively [9–11]. In this chapter, we describe the commonly used methods for the detection of cellulase and xylanase activities from Trichoderma reesei, which belongs to filamentous fungi and is commonly used for cellulase production [1]. Some useful guidance is provided here, which is beneficial for the reproducibility of the methods in different labs. In case that other microbial producers are used, a suitable pH value and adjusted temperatures for the assays should be considered.

2

Materials Prepare 25 mL glass colorimetric cylinders with stoppers and 96-well microplates for absorbance tests according to the number of samples. A spectrophotometer suitable for measuring absorbance at 540 nm and 420 nm should also be prepared in advance. Prepare all solutions using deionized water. Prepare the solutions at room temperature, and better results can be obtained using freshly prepared solutions. Solutions of CMC, pNPC, and pNPG should be stored at 4
C after preparation. The solutions needed are listed below for different enzymes in the cellulase complex.

2.1 Total Cellulase Activity Assay

1. DNS reagent: Weigh 10 g of NaOH and dissolve in 600–700 mL of deionized water, mix well to make sodium hydroxide solution in a 1 L glass beaker. Add 10 g of 3,5-dinitrosalicylic acid, 5 g of anhydrous sodium sulfite, 200 g of tartaric acid (sodium salt), and 2 g of melted phenol (50
C) (see Note 1) to the NaOH solution. After mixing completely, dilute the solution to 1 L with deionized water. Store in a brown reagent bottle, and place in the dark for over 12 h before use. After the preparation of the DNS reagent, it is better to finish using the solution within 1 month (see Note 2). 2. 50 mM citrate buffer: Weigh 4.83 g of citric acid monohydrate and dissolve it in about 750 mL of deionized water. While stirring, add 7.94 g of trisodium citrate to the solution and adjust to pH 4.8, then dilute to a final volume of 1 L with deionized water. Store at room temperature. 3. Fast qualitative filter paper strips: Cut Whatman No. 1 filter paper to 1  6 cm strips (approximately 50 mg) with a paper cutter (see Note 3). 4. Glucose standard stock solution: Dry anhydrous glucose to a constant weight at 60
C. Accurately prepare a 4 mg/mL glucose solution with 50 mM citrate buffer (see Note 4).

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2.2 Exoglucanase ( pNPCase) and β-Glucosidase ( pNPGase) Assays

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1. 50 mM citrate buffer (see Subheading 2.1). 2. 1 mg/mL 4-nitrophenyl β-D-cellobioside ( pNPC) in citrate buffer: Dissolve 10 mg of pNPC in 10 mL of 50 mM citrate buffer. 3. 1.5 mg/mL 4-nitrophenyl β-D-glucopyranoside ( pNPG) in citrate buffer: Dissolve 15 mg of pNPG in 10 mL of 50 mM citrate buffer. 4. 1 μg/μL 4-nitrophenol ( pNP) in citrate buffer: Dissolve 10 mg of pNP in 10 mL of 50 mM citrate buffer. 5. 10% w/v Na2CO3 buffer: Dissolve 10 g of Na2CO3 in 100 mL of 50 mM citrate buffer.

2.3 Endoglucanase (CMCase) and Xylanase Activity Assays

1. 50 mM citrate buffer (see Subheading 2.1). 2. Substrate solution for endoglucanase assays: 1% w/v carboxymethyl cellulose (CMC) in 50 mM citrate buffer (see Note 5). 3. Substrate solution for xylanase assays: Oat spelts xylan solution in 50 mM citrate buffer. 4. DNS reagent (see Subheading 2.1). 5. Glucose standard solution (see Subheading 2.1). 6. Xylose standard stock solution: Prepare 1 mg/mL xylose solution with 50 mM citrate buffer for the determination of xylanase activity.

3

Methods 1. Sample 1–3 mL of the liquid culture of T. reesei at different time points. 2. Centrifuge the samples at 8000  g for 10 min, collect the supernatant as crude enzyme, and preserve at 4
C until determination of the enzyme activity.

3.1 Total Cellulase Activity Assay

Total cellulase activities are always measured using insoluble cellulosic substrates, such as Whatman No. 1 filter paper, cotton, microcrystalline cellulose, and pretreated lignocellulose [12]. At present, the most accepted substrate for total cellulase activity assay is Whatman No. 1 filter paper. The assay requires a certain amount (at least 2 mg) of glucose released from a 50 mg Whatman No. 1 filter paper, which ensures that both crystalline and amorphous fractions of the filter paper are hydrolyzed. The advantages of FPA are: (1) the substrate is commonly and widely available in different labs; (2) the substrate is moderately susceptible to cellulase; and (3) the procedure is simple to compare cellulase activities. On the other hand, FPA should be performed with well-controlled parameters, since many factors such as complex reagents, operational

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steps, folded versus rolled filter paper, and filter paper cutting methods affect the final results [13, 14]. One unit of filter paper activity (FPase) is defined as the amount of enzyme required to release 1 μmol of reducing sugar per minute and is expressed as FPU/mL. 3.1.1 FPA Procedures

1. Put a rolled filter paper strip (see Subheading 2.1) at the bottom of a 25 mL colorimetric cylinder. 2. Add 1.5 mL of 50 mM citrate buffer to the cylinder. The filter paper strip should be submerged in the buffer. 3. Prepare a dilution series of the crude enzyme solution (see Note 6). 4. Pipette 0.5 mL of the crude enzyme dilutions into the cylinders and mix well. 5. Prepare the control group: 1.5 mL of 50 mM citrate buffer with filter paper strip. 6. Incubate all the cylinders in a 50
C water bath for exactly 1 h. 7. Add 3 mL of the DNS reagent to stop the reaction and mix well. 8. Add 0.5 mL of diluted enzyme solution to the control cylinder. 9. Boil all the cylinders for exactly 10 min (see Note 7). 10. Transfer all the cylinders to a tap water bath to cool to room temperature and dilute the reaction solutions to 25 mL with deionized water. Mix well by inverting the cylinders several times. 11. Pipette 0.2 mL of each reaction solution into a 96-well plate and measure absorbance at 540 nm. The absorbance of the control group is used as the blank.

3.1.2 Standard Sugar Curve

1. Prepare a serial dilution of glucose standard solutions using 50 mM citrate buffer in 25 mL colorimetric cylinders (see Table 1).

Table 1 Standard glucose solutions of different concentrations Tube No. Reagent

1

2

3

4

5

6

7

8

Glucose standard solution (mL)

0

0.2

0.3

0.4

0.5

0.6

0.7

0.8

Citric acid buffer (mL)

2

1.8

1.7

1.6

1.5

1.4

1.3

1.2

Glucose content (mg)

0

0.8

1.2

1.6

2.0

2.4

2.8

3.2

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Fig. 1 The relationship between absorbance at OD540 for the DNS assay and the standard glucose content

2. Add 3 mL of DNS reagent to each standard glucose solution and stir evenly. Put into boiling water and keep the reaction for exactly 10 min. 3. Transfer all the cylinders to a tap water bath and cool to room temperature. Dilute the reaction mixture to 25 mL with deionized water, and mix well. 4. Pipette 0.2 mL of each solution into a 96-well plate and determine the absorbance at 540 nm. 3.1.3 Calculation

1. Draw a standard glucose curve as shown in Fig. 1. 2. Calculate the delta absorbance of the diluted enzyme solutions (see Note 8). 3. Calculate the real reducing sugar concentrations released by the diluted crude enzyme according to the delta absorbance and the standard sugar curve. 4. Calculate the FPase of the crude enzyme. The equation is as follows: Δ reducing sugar ðmgÞ 5. FPase ðIU=mLÞ ¼ 0:18 ðmg=μmol Þ60 min 0:5 mL  DF

where DF refers to the dilution factor of the crude enzyme. For example, a DF of 100 means a 1:100 dilution. 3.2 Exoglucanase ( pNPCase) and β-Glucosidase ( pNPGase) Activity Assays

Exoglucanases or cellobiohydrolases (EC 3.2.1.91) act on cellulose chain ends releasing β-1,4-cellobiose which is then hydrolyzed to glucose by β-glucosidases. Microcrystalline cellulose (Avicel) is a good substrate for exoglucanase activity assays because of its relatively low degree of polymerization (DP) as well as relatively high accessibility [15, 16]. Therefore, avicelases were recognized to have exoglucanase activity before [4]. However, endoglucanases can also

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release reducing sugars from amorphous cellulose, which could affect the calculated accuracy of the exoglucanase activity [2]. Therefore, pNPC is commonly used to evaluate and reflect the exoglucanase activity [17]. β-Glucosidase (EC 3.2.1.21) can hydrolyze β-1,4-glycosidic bonds of soluble substrates, including cellobiose and other cellodextrins with a DP of up to 6 [15]. pNPG and cellobiose as substrates are usually used to indicate the β-glucosidase activity. Here, we describe the pNPG assay. One unit of exoglucanase ( pNPCase) or β-glucosidase ( pNPGase) activity is defined as the amount of enzyme required to release 1 μmol of 4-nitrophenol ( pNP) per minute. 3.2.1 Procedure

1. Add 50 μL of the pNPC or pNPG solution into a 1.5 mL microcentrifuge tube. 2. Equilibrate at 50
C until use. 3. Prepare a dilution series of the crude enzyme solution using 50 mM citrate buffer. 4. Add 100 μL of the diluted crude enzyme to the test tubes and mix vigorously. 5. For the control group, add 150 μL of 10% Na2CO3 solution to the tube. Then add 100 μL of the diluted crude enzyme to the tube and mix well. 6. For the reaction, incubate all the tubes in a 50
C water bath for exactly 30 min. 7. Add 150 μL of 10% Na2CO3 solution to the test tubes to stop the reaction (except the control tubes), then mix well. 8. Pipette 100 μL of each solution into a 96-well plate. 9. Measure the absorbance of liberated products of pNP at 420 nm, where the absorbance of the control is used as the blank (see Note 9).

3.2.2 Calculation

1. Draw a standard pNP curve as shown in Fig. 2. 2. Calculate the delta absorbance of the diluted enzyme solutions. 3. Calculate the real pNP concentrations released by the crude enzyme dilutions according to the delta absorbance and the standard pNP curve. 4. Calculate the pNPCase or pNPGase of the crude enzyme. The equation is as follows: Δ pNP ðμgÞ 5. pNPCase or pNPGase ðIU=mLÞ ¼ 139:11ðμg=μmol Þ30 min 0:1 mL  DF

where DF refers to the dilution factor of the crude enzyme.

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Fig. 2 The relationship between absorbance at OD420 and the standard pNP content

Table 2 Standard pNP solutions of different concentrations Tube No. Reagent

1

2

3

4

5

6

7

8

1 μg/μL pNP (μL)

0

5

10

20

30

40

50

60

Citrate buffer (μL)

150

145

140

130

120

110

100

90

0

5

10

20

30

40

50

60

pNP content (μg)

3.2.3 Standard pNP Curve

1. Prepare a 1 μg/μL pNP solution with 50 mM citrate buffer and then prepare a series of pNP standard solutions using 50 mM citrate buffer (see Table 2). 2. Add 150 μL of 10% Na2CO3 solution, and mix well. Then pipette 100 μL of each solution into a 96-well plate and measure absorbance at 420 nm. 3. Draw a standard pNP curve (see Fig. 2).

3.3 Endoglucanase (CMCase) Activity Assay

Endoglucanases (EC 3.2.1.4) randomly cleave intramolecular β-1,4-glycosidic linkages on the surface of amorphous cellulose. Through the soluble oligosaccharides and their derivatives as the substrate, endoglucanase activity can be measured by the formation of reducing sugars [18, 19]. However, at present, CMC is often used for determining endoglucanase activity according to the IUPAC protocol, called CMCase, since endoglucanases can specifically cleave the glycosidic bonds, resulting in a reduction in the DP of CMC. Furthermore, the procedure is simple [12].

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One unit of endoglucanase (CMCase) activity is defined as the amount of enzyme required to release 1 μmol of glucose per minute and is expressed as IU/mL. 3.3.1 Procedure

1. Add 1.5 mL of the CMC substrate solution to a 25 mL colorimetric cylinder. 2. Prepare the enzyme dilution series. 3. Equilibrate the substrate solution and enzyme solutions at 50
C. 4. Add 0.5 mL of diluted crude enzyme to the cylinders, and mix well. 5. Incubate the cylinders at 50
C for 30 min (water bath). 6. Add 3 mL of DNS reagent to the cylinders to stop the reaction. 7. Prepare the control group: First add 3 mL of DNS reagent to the cylinder containing 1.5 mL of CMC solution, then add 0.5 mL of diluted crude enzyme in the cylinder. Mix well. 8. Boil all the cylinders for exactly 10 min in vigorously boiling water. 9. Transfer the cylinders to a cold water bath and dilute to 25 mL with deionized water. Mix well. 10. Pipette 0.2 mL of each solution into a 96-well plate. 11. Measure the absorbance at 540 nm, where the absorbance of the control is used as the blank.

3.3.2 Calculation

1. Calculate the delta absorbance of the diluted enzyme solutions. 2. Calculate the real reducing sugar concentrations released by the diluted crude enzyme according to the standard sugar curve (see Fig. 1). 3. Calculate the CMCase activity of the crude enzyme. The equation is as follows: Δ reducing sugar ðmgÞ CMCase ðIU=mLÞ ¼ 0:18 ðmg=μmol Þ30 min 0:5 mL  DF

where DF refers to the dilution factor of the crude enzyme. 3.4 Xylanase Activity Assay

Xylanases are hydrolytic enzymes which can randomly cleave the β-1,4-glycosidic bond of the polysaccharide xylan. Xylan is the second most abundant polysaccharide in lignocellulosic biomass, demanding xylanases to degrade to xylose. Hemicelluloses from different plant sources have been used for D-xylanase activity assays. The most commonly used substrates are arabinoglucuronoxylan, arabinoxylan, glucuronoxylan and xylan. According to IUPAC recommendations [20], the xylanase activity was measured using oat spelts xylan as a substrate.

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Xylanase activity is expressed as the number of μmol of reducing sugars produced per minute of hydrolysis per mL of the crude enzyme used. 3.4.1 Procedure

1. Add 1.5 mL of the xylan substrate solution to a 25 mL colorimetric cylinder. 2. Prepare the enzyme dilution series. 3. Equilibrate the substrate solution and enzyme solutions at 50
C. 4. Add 0.5 mL of diluted crude enzyme to the cylinders. Mix well. 5. Incubate the cylinders at 50
C for 30 min (water bath) (see Note 10). 6. Add 3 mL of DNS reagent to the cylinders to stop the reaction. 7. Prepare the control group: add 3 mL of DNS reagent to the cylinder containing 1.5 mL of xylan substrate solution, then add 0.5 mL of the diluted crude enzyme to the cylinder. Mix well. 8. Boil all the cylinders for exactly 10 min. 9. Transfer the cylinders to a tap water bath to cool to room temperature and dilute the reaction mixture to 25 mL with deionized water. Mix well. 10. Pipette 0.2 mL of each solution into a 96-well plate and measure the absorbance at 540 nm, where the absorbance of the control is used as the blank.

3.4.2 Calculation

1. Prepare a series of xylose standard solutions diluted with 50 mM citrate buffer (see Table 3). 2. Add 3 mL of DNS reagent to each standard solution of xylose, stir evenly. To start the reaction, put into boiling water for 10 min.

Table 3 Standard xylose solutions of different concentrations Tube No. Reagent

1

2

3

4

5

6

7

8

Xylose standard solution (mL)

0

0.2

0.4

0.6

0.8

1.0

1.2

1.5

Citrate buffer (μL)

2.0

1.8

1.6

1.4

1.2

1.0

0.8

0.5

Xylose content (mg)

0

0.2

0.4

0.6

0.8

1.0

1.2

1.5

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Fig. 3 The relationship between absorbance at OD540 for the DNS assay and the standard xylose content

3. Transfer the cylinders to a tap water bath to cool to room temperature and dilute the mixture to 25 mL with deionized water. Mix well. 4. Pipette 0.5 mL of the supernatant into a 5 mL tube and dilute with 2 mL of deionized water, then mix completely. 5. Pipette 0.2 mL of each solution into a 96-well plate and determine the absorbance at 540 nm. Draw a standard sugar curve (see Fig. 3). 6. Calculate the delta absorbance of the diluted enzyme solutions. 7. Calculate the real reducing sugar concentrations released by the diluted crude enzyme according to the delta absorbance and the standard sugar curve. 8. Calculate the xylanase activity of the crude enzyme. The equation is as follows: Δ reducing sugar ðmgÞ xylanase ðIU=mLÞ ¼ 0:15 ðmg=μmol Þ30 min 0:5 mL  DF

where DF refers to the dilution factor of the crude enzyme.

4

Notes 1. Phenol is quite toxic. Be careful to handle the phenol safely. 2. The freshness of the DNS reagent is important. It could lose reducing ability after long time storage; therefore, DNS should be used within 1 month after preparation. Store the DNS reagent at 4
C.

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3. The exact weight of the filter paper strip is very important for the reproducible cellulase activity assay. Make sure that the strip weight variation is less than 1 mg. In addition, handle the strip with gloved hands. 4. The standard solution should be mixed well before starting the experiment. 5. CMC is very difficult to be dissolved in citrate buffer. Therefore, slowly add and stir until the solids are completely dissolved at 50
C. 6. Dilute the crude enzyme preparation to appropriate concentrations using 50 mM citrate buffer, and make sure that the correctly diluted solution releases about 2 mg of glucose from the filter paper strip (substrate amounts strongly influence enzyme activity). 7. Make sure that the volume of the boiling water bath is maintained above the level of the total liquid volume of the test cylinders. 8. The delta absorbance refers to the absorbance of the sample minus the absorbance of the control at 540 nm. 9. Properly control the solution concentration, and make sure that the absorbance at 420 nm does not exceed 1.2 during the assay. 10. Shake the cylinders gently from time to time during the reaction process in order to mix evenly.

Acknowledgments This work was supported from the Natural Science Foundation of China (grant number 21536006). References 1. Zhang F, Bunterngsook B, Li J-X, Zhao X-Q, Champreda V, Liu C-G, Bai F-W (2019) Chapter 3 - Regulation and production of lignocellulolytic enzymes from Trichoderma reesei for biofuels production. In: Li Y, Ge X (eds) Advances in bioenergy, vol 4. Elsevier, Cambridge, MA, pp 79–119. https://doi.org/10. 1016/bs.aibe.2019.03.001 2. Wang M, Li Z, Fang X, Wang L, Qu Y (2012) Cellulolytic enzyme production and enzymatic hydrolysis for second-generation bioethanol production. In: Bai F-W, Liu C-G, Huang H, Tsao GT (eds) Biotechnology in China III: biofuels and bioenergy. Springer, Berlin, pp

1–24. https://doi.org/10.1007/10_2011_ 131 3. Mullings R (1985) Measurement of saccharification by cellulases. Enzyme Microb Technol 7 (12):586–591. https://doi.org/10.1016/ 0141-0229(85)90025-0 4. Wood TM, Bhat KM (1988) Methods for measuring cellulase activities. Methods Enzymol 160:87–112. https://doi.org/10.1016/ 0076-6879(88)60109-1 5. Helbert W, Chanzy H, Husum TL, Schu¨lein M, Ernst S (2003) Fluorescent cellulose microfibrils as substrate for the detection of cellulase activity. Biomacromolecules 4

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(3):481–487. https://doi.org/10.1021/ bm020076i 6. Eveleigh DE, Mandels M, Andreotti R, Roche C (2009) Measurement of saccharifying cellulase. Biotechnol Biofuels 2(1):21. https://doi. org/10.1186/1754-6834-2-21 7. Dashtban M, Maki M, Leung KT, Mao C, Qin W (2010) Cellulase activities in biomass conversion: measurement methods and comparison. Crit Rev Biotechnol 30(4):302–309. https://doi.org/10.3109/07388551.2010. 490938 8. Ghose T (1987) Measurement of cellulase activities. Pure Appl Chem 59(2):257–268 9. Zhang YHP, Hong J, Ye X (2009) Cellulase Assays. In: Mielenz JR (ed) Biofuels: methods and protocols. Humana Press, Totowa, NJ, pp 213–231. https://doi.org/10.1007/978-160761-214-8_14 10. Jourdier E, Cohen C, Poughon L, Larroche C, Monot F, Chaabane FB (2013) Cellulase activity mapping of Trichoderma reesei cultivated in sugar mixtures under fed-batch conditions. Biotechnol Biofuels 6(1):79. https://doi.org/ 10.1186/1754-6834-6-79 11. Deshpande MV, Eriksson K-E, Go¨ran Pettersson L (1984) An assay for selective determination of exo-1,4,-β-glucanases in a mixture of cellulolytic enzymes. Anal Biochem 138 (2):481–487. https://doi.org/10.1016/ 0003-2697(84)90843-1 12. Percival Zhang YH, Himmel ME, Mielenz JR (2006) Outlook for cellulase improvement: screening and selection strategies. Biotechnol Adv 24(5):452–481. https://doi.org/10. 1016/j.biotechadv.2006.03.003 13. Coward-Kelly G, Aiello-Mazzari C, Kim S, Granda C, Holtzapple M (2003) Suggested improvements to the standard filter paper assay used to measure cellulase activity. Biotechnol Bioeng 82(6):745–749. https://doi. org/10.1002/bit.10620

14. Zhang YHP, Lynd LR (2005) Determination of the number-average degree of polymerization of cellodextrins and cellulose with application to enzymatic hydrolysis. Biomacromolecules 6(3):1510–1515. https:// doi.org/10.1021/bm049235j 15. Zhang Y-HP, Lynd LR (2004) Toward an aggregated understanding of enzymatic hydrolysis of cellulose: noncomplexed cellulase systems. Biotechnol Bioeng 88(7):797–824. https://doi.org/10.1002/bit.20282 16. Zhang Y-HP, Lynd LR (2006) A functionally based model for hydrolysis of cellulose by fungal cellulase. Biotechnol Bioeng 94 (5):888–898. https://doi.org/10.1002/bit. 20906 17. Bhat KM, Hay AJ, Claeyssens M, Wood TM (1990) Study of the mode of action and sitespecificity of the endo-(1-4)-beta-D-glucanases of the fungus Penicillium pinophilum with normal, 1-3H-labelled, reduced and chromogenic cello-oligosaccharides. Biochem J 266(2):371–378. https://doi.org/10.1042/ bj2660371 18. Bhat S, Kennedy JF, Goodenough PW, Owen E, Bhat MK (1997) Effect of d-glucono-1,4-lactone on the production of CMCase, pNPCase and true cellulase by Clostridium thermocellum. Carbohyd Polym 34 (1):95–99. https://doi.org/10.1016/S01448617(97)00049-0 19. Claeyssens M, Aerts G (1992) Characterisation of cellulolytic activities in commercial Trichoderma reesei preparations: an approach using small, chromogenic substrates. Bioresour Technol 39(2):143–146. https://doi.org/10. 1016/0960-8524(92)90133-I 20. Ghose T, Bisaria VS (1987) Measurement of hemicellulase activities: part I xylanases. Pure Appl Chem 59(12):1739–1751

Chapter 13 Flow Cytometry for Filamentous Fungi Matthias G. Steiger Abstract Flow cytometry is a powerful high-throughput method, which enables a fast and multi-parameter analysis of single cells and particles. A plethora of different dyes for flow cytometry are available to label different parts of a cell in addition to in vivo markers like fluorescent proteins. Trichoderma species as well as other filamentous fungi show hyphal growth, which makes analysis in a flow cytometer difficult. Nevertheless, conidia can be readily analyzed in conventional flow cytometers. Many different applications can be envisaged. This protocol describes how conidia can be prepared for flow cytometry and the occurrence of genetic markers such as GFP can be measured. Furthermore, a guideline how to fix and stain cells is given. Key words FACS, Conidia, Sorting, Fluorescence, Staining, Fixation, Filamentous fungi, Trichoderma reesei, GFP

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Introduction Flow cytometry is a powerful high-throughput method, which enables a fast and multi-parameter analysis of single cells and particles. Flow cytometers can be equipped with a sorter with which subpopulations or single cells with different optical properties can be separated. Although flow cytometry is an established technique to analyze bacteria, yeast, and mammalian cell cultures, it is rarely used for filamentous fungi. The obvious explanation can be found in the complex morphology of fungi. Elongated, hyphal growth makes it harder to analyze fungal cells in a flow cytometer than single and almost spherical cells. Nevertheless, it is possible to analyze conidia [1], germlings [2], and protoplasts [3] of Trichoderma reesei in a conventional flow cytometer. Another option is to encapsulate the fungal conidia into, e.g., calcium alginate microcapsules, which enable sorting after a prolonged cultivation period. Complex Object Parametric Analyzer and Sorter (COPAS) instruments must be used to sort such alginate particles [4]. A recent review provides an excellent overview on flow cytometry studies

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performed with filamentous fungi [5]. This universal technique can be used for different purposes including assessing viability, detection of genetic markers like GFP and DsRed but also cell size, morphology and DNA content measurements. In a flow cytometer, cells/particles are moved into a flow cell, which focuses cells into a single line and moving them through a laser beam measuring optical properties. Light scattering and fluorescence of each single particle can be analyzed at high speed, typically 1000–50,000 cells/ s depending on the capabilities of the instrument. Figure 1 schematically displays the parts of a flow cytometer and outlines the working principles. For further details, the reader is referred to excellent literature regarding the construction and function of flow cytometers and sorters [6, 7]. A plethora of different flow

Fig. 1 Schematic drawing of a flow cytometer with a cell sorter. The two boxes in light blue showing the sample port and the collection port are accessible to the user. The containers for the sheath fluid and for the waste are typically located beside the machine. In order to enable aseptic collection of the samples at the sample port, the entire machine should be placed into a laminar flow device. The sheath fluid is used to envelop the sample stream for the hydrodynamic focusing of the cells in the flow cell. After the flow, cell droplets are generated by a nozzle and charged in order to separate them between charged deflection plates. Based on the optical properties, cellular subpopulations can be sorted and collected. The optical system consists of one or more lasers, which are guided through the flow cell interacting with the cells. Light scattered in the forward direction is collected by a photomultiplier (PMT) detector behind the flow cell (forward scatter channel, FSC). Light scattered orthogonally is collected by a PMT detector and termed side scatter channel (SSC). Furthermore, the orthogonal beam coming from the flow cell is split up by an optical filter system into different fluorescence channels. Filters used are divided into bandpass (BP), shortpass (SP), and longpass filters (LP). The fluorescence channels are often abbreviated as FL1 (green), FL2 (orange), and FL3 (red). However, modern flow cytometers can have much more fluorescence channels, which should be better named according to respective bandpass filter, e.g., BP 525/50. This filter allows light to pass that is of a range of wavelengths of 500–550 nm (525  25 nm)

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cytometers are available on the market, which have different features and requirements. Therefore, before starting, it is necessary to carefully read the manual and receive instructions from the trained user or operator. In particular the optical capabilities of an instrument can be very different. An excessive catalogue of available fluorophores and their optical properties can be found in the Molecular Probes Handbook, which is distributed by Thermo Fisher Scientific [8]. This protocol describes how conidia are prepared for flow cytometry and how their size distribution and the occurrence of a genetic markers like GFP can be measured.

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Materials Prepare all solutions using distilled or deionized water and analytical grade reagents. All solutions used for flow cytometry are filtered through a 0.22 μm filter to reduce contamination with particles and to sterilize the solutions.

2.1 Conidia Harvesting and Washing

1. Glycerol stock of strain to be analyzed, e.g., Trichoderma reesei QM6a, QM9414. 2. Minimal medium agar plate or a plate with a desired growth medium (see Note 1). 3. 15 mL conical tubes. 4. Tween solution: 0.5% (w/v) Tween 20. Dissolve 5 g of Tween 20 in 1000 mL water by stirring. 5. PBS-Tween (1): 137 mM NaCl, 10 mM Na2HPO4, 2.7 mM KCl, 2 mM KH2PO4, and 0.5% (w/v) Tween 20. For 1000 mL of a 1 PBS solution, dissolve 8 g of NaCl, 1.44 g of Na2HPO4, 0.2 g of KCl, 0.24 g of KH2PO4, and 5 g of Tween 20 in 850 mL water by stirring. Adjust the pH to 7.4 with 0.1 M HCl and fill up to 1000 mL with water. 6. Cotton swabs: Sterile cotton swabs with a stick length of at least 15 cm. 7. Filter funnel with miracloth: Place cut miracloth squares (12  12 cm) into filter funnels with a diameter of 6 cm and fix it with a strip of autoclave tape at the outside of the funnel. Autoclave for 20 min at 1 bar (positive pressure). 8. Centrifuge with rotors for 15 mL and 1.5 mL reaction tubes. 9. Instrument for measuring conidia concentration (e.g., coulter counter, Thoma cell counting chamber, or an optical density measurement). 10. Mixer for reaction tubes. 11. Cell strainer (35 μm mesh). 12. Light microscope.

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2.2 Fixation of Conidia

1. 70% (v/v) ethanol.

2.3 DNA Staining with Propidium Iodide (PI)

1. PI staining solution: 10 mM Tris–HCl, 1 mM EDTA adjust pH to 7.0 with a final concentration of propidium iodide of 50 μg/ mL.

2. PBS-Tween (1) (see Subheading 2.1).

2. Tris–EDTA buffer (TE; 1): 10 mM Tris–HCl, 1 mM EDTA (pH 7.6), pH 7.6. 3. RNase solution: 0.1% (w/v) DNase-free Ribonuclease A. Dissolve 10 mg of DNase-free Ribonuclease A from bovine pancreas in 10 mL of 1 TE buffer (pH 7.6). 2.4

Flow Cytometry

1. Flow cytometer with at least one laser, typically with 488 nm, and a channel for forward scatter (FSC), side scatter (SSC), and two fluorescence channels (see Note 2). 2. Tubes for flow cytometer (see Note 3). 3. Sheath fluid depending on the requirements of the flow cytometer (see Note 4). 4. Rinse and clean solutions depending on the requirements of the flow cytometer.

2.5 Flow Cytometry and Sorting

1. Flow cytometer as specified under Subheading 2.4 with an integrated sorting device. Most sorters rely on a droplet sorting principle. Drops are generated by resonance, charged depending on their content, and distracted between deflection plates. 2. Sterile 96-well plates prefilled with 50 μL of PBS–Tween solution or a suitable growth medium.

3

Methods

3.1 Preparation of Conidia for Flow Cytometry

All working steps need to be carried out aseptically. 1. Inoculate the strain from a glycerol stock on a minimal medium plate or on a plate with a desired growth medium. 2. Incubate the plate at the optimal growth temperature of the strain (typically 30  C) until dense conidiation is observed after 3–8 days. 3. For each strain to be analyzed pipette 10 mL of Tween solution into a 15 mL conical tube. 4. Harvest the conidia by soaking a sterile cotton swab in Tween solution and scratching the conidia from the plate and transfer them to the conical tube. Vortex the suspension.

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5. Place a filter funnel with miracloth into a fresh 15 mL conical tube and pipette the conidial suspension into it. Allow it to pass through by gravitational flow. 6. Centrifuge the suspension at 2500  g, 10 min, 4  C and discard the supernatant. 7. Resuspend the conidia pellet in 10 mL of PBS–Tween solution and repeat steps 6 and 7. 8. Measure the conidia concentration by using a coulter counter, Thoma cell counting chamber, or an optical density measurement (see Note 5). 9. To obtain the conidia stock solution, dilute the conidia suspension in PBS–Tween solution to a final concentration of 1  107 conidia/mL. 10. Vortex the suspension for 1 min and filter through a cell strainer with a 35 μm mesh. Pipette 10 μL onto a microscope slide and apply a cover slip. 11. Check under a light microscope at a magnification of 400 (ocular 10, objective 40) whether the conidia are well separated and the suspension is ready for flow cytometry (see Note 6). In case conidia are aggregated, proceed to step 12. 12. Pulsing with an ultrasonic device enables to separate conidia. The conidia suspension is pulsed with five pulses, duty cycle 20%, output control 2 (~40 W) (using Branson Ultrasonics Sonifier 250). 3.2

Fixation of Cells

1. For each fixated sample, prepare a non-fixation control by adding 1.5 mL of PBS–Tween solution instead of the fixation solution at step 3. 2. Add 1.5 mL of the conidia stock solution (obtained at step 9, see Subheading 3.1) to a 2 mL reaction tube, centrifuge at 12,500  g, 5 min, 4  C, and discard the supernatant. 3. Add 1.5 mL of fixation solution and mix by vortexing. 4. Incubate at 4  C, shaking at 300 rpm for 20 min. 5. Centrifuge at 12,500  g, 5 min, 4  C and resuspend in 1.5 mL of PBS–Tween solution. Follow steps 10–12 (see Subheading 3.1).

3.3 DNA Staining of Conidia with PI

1. For each stained sample, prepare a non-stained control. 2. Add 1.5 mL of the conidia stock solution (obtained at step 9, see Subheading 3.1) to a 2 mL reaction tube, centrifuge at 12,500  g, 5 min, 4  C, and discard the supernatant. 3. Resuspend in 1.5 mL of PI staining solution and mix by vortexing.

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4. Incubate at room temperature for 20 min. 5. Centrifuge at 12,500  g, 5 min, 4  C and resuspend in 1.5 mL of PBS–Tween solution. Follow steps 10–12 (see Subheading 3.1). 3.4 Flow Cytometry and Sorting

1. Fill up sheath flow and empty waste container of the flow cytometer. 2. Perform the start-up and quality control routine of the flow cytometer. 3. The prepared conidia suspension is transferred to a test tube suitable for the sample port of the used flow cytometer. 4. Prepare acquisition diagrams in the software for FSC and SSC (see Note 7). 5. Start measuring the sample in the setup mode of the flow cytometer and adjust the gain settings of FSC and SSC and make sure that the entire population is visible (see Note 8). An example for a conidia population is provided in Fig. 2a. 6. Gate the conidia population and discriminate duplicates by plotting FSC-Area (FSC-A) against FSC-Height (FSC-H). Set a gate for single conidia (see Note 9).

Fig. 2 Example diagrams often used in flow cytometry. (a) Bivariate plot showing forward scatter data plotted against side scatter of a Trichoderma reesei population. The elliptical gate in red highlights single conidia. (b) Histogram of a fluorescence channel using a 488 nm excitation laser with a BP 530/30 filter, which can be used for the fluorophore of GFP. Two different samples are overlaid: in red on the left side of the histogram appears the negative control, a wild-type strain not expressing GFP. In green on the right side, the measurements of a strain expressing GFP is shown

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7. Prepare acquisition diagrams for the fluorescence parameter. A histogram plot showing fluorescence measurements of a strain expressing GFP and the respective parental strain is shown in Fig. 2b. 8. Start acquisition of a stained sample in the setup mode and adjust the gain settings of the acquisition channels. 9. Start measurement of samples and record at least 10,000 events (in the gate of single conidia) and save all measurements. 10. For cell sorting, specify a sorting gate and collection mode (amount of conidia sorted to each collection container). 11. For cell sorting, install a collection plate/tube and start sorting. 12. After sorting one cell population, clean the sample inlet port and clean the flow cell using the rinsing and cleaning agents recommended by the flow cytometer provider. 13. Run a sample with PBS–Tween solution in acquisition mode for 2 min to check for remaining cells in the system. Repeat the cleaning of the flow cell if more than 50 events are recorded in the cell gate. 14. Perform the shutdown routine of the flow cytometer rinsing and cleaning the flow cell.

4

Notes 1. If complex media components contain insoluble particles, e.g., mixed cereal agar plates, these particles are harvested together with the conidia and will be detected in the flow cytometer. Depending on the size of these particles, it can be difficult to remove them from a cytograph by gating. A possibility to remove this background is to cover the plate with cellophane foil and inoculate the fungal conidia on top. For that, cut cellophane foil in round pieces and autoclave them between filter paper soaked in dH2O. Transfer the round cellophane pieces aseptically with tweezers onto the agar plate and remove all air bubbles using a Drigalski spatula. Inoculate the conidia suspension on top of the cellophane foil. 2. Modern flow cytometers can be equipped with up to six lasers and 30 independent detection channels. 3. Test tubes have typically a total volume of 5 mL and a round bottom. Tubes are available, which have already a cell strainer integrated in the cap, which can be used at step 9 (see Subheading 3.1). Test tubes are filled with 2–3 mL of the conidia suspension. Some flow cytometers also accept standard 96-plate formats and are filled with 150–200 μL.

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4. If not specified by the provider, it is possible to use ultra-pure and particle-free water (e.g., MilliQ® water). 5. Conidia suspensions that are visually impermeable in a 15 mL tube need to be diluted prior to analyses in a Thoma chamber. If a strain is analyzed repeatedly, it is useful to measure the optical density of the conidia suspension at different dilutions at 600 nm and make a plotting cell count against optical density. Note that the accuracy of the calibration is strain specific and also depending on the growth medium of the conidia plate. 6. Prepared conidia suspensions must not be stored as gravitational forces will lead to sedimentation and agglomeration. Vortexing the conidia suspension every 3 min prevents sedimentation. 7. Create a bivariate plot of FSC and SSC to enable a visual inspection of the conidia population. Depending on the size distribution, it should be possible to visualize both at a linear scale. However, if in an experiment swollen conidia are tested, it might be necessary to apply a logarithmic scale on the diagram. 8. If different size distributions between samples are expected, adjust the gain settings for FSC and SSC with the sample expected to contain the largest conidia and make sure that these cells are still within the linear range of the detector. 9. For doublet discrimination, other plots like FSC-A against FSC-Width (FSC-W) or FSC-H against FSC-W can be used. In case a plethora of doublets can be observed, the conidial suspension can be further diluted and sonicated (proceed with step 12, see Subheading 3.1).

Acknowledgments This work has been supported by the Federal Ministry for Digital and Economic Affairs (bmwd), the Federal Ministry for Transport, Innovation and Technology (bmvit), the Styrian Business Promotion Agency SFG, the Standortagentur Tirol, Government of Lower Austria and ZIT—Technology Agency of the City of Vienna through the COMET-Funding Program α managed by the Austrian Research Promotion Agency FFG.

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References 1. Throndset W, Kim S, Bower B, Lantz S, Kelemen B, Pepsin M, Chow N, Mitchinson C, Ward M (2010) Flow cytometric sorting of the filamentous fungus Trichoderma reesei for improved strains. Enzym Microb Technol 47:335–341 2. Gao F, Hao Z, Sun X, Qin L, Zhao T, Liu W, Luo H, Yao B, Su X (2018) A versatile system for fast screening and isolation of Trichoderma reesei cellulase hyperproducers based on DsRed and fluorescence-assisted cell sorting. Biotechnol Biofuels 11:261 3. Wang G, Jia W, Chen N, Zhang K, Wang L, Lv P, He R, Wang M, Zhang D (2018) A GFP-fusion coupling FACS platform for advancing the metabolic engineering of filamentous fungi. Biotechnol Biofuels 11:232

4. Delgado-Ramos L, Marcos AT, Ramos-Guelfo MS, Sa´nchez-Barrionuevo L, Smet F, Cha´vez S, Ca´novas D (2014) Flow cytometry of microencapsulated colonies for genetics analysis of filamentous fungi. G3 (Bethesda) 4:2271–2278 5. Bleichrodt RJ, Read ND (2019) Flow cytometry and FACS applied to filamentous fungi. Fungal Biol Rev 33:1–15 6. Bu¨scher M (2019) Flow cytometry instrumentation – an overview. Curr Protoc Cytom 87:1–8 7. Shapiro HM (2003) Practical flow cytometry. Wiley, New York 8. Johnson I, Spence M (2010) The molecular probes handbook—a guide to fluorescent probes and labeling technologies, 11th edn. Life Technologies

Chapter 14 Molecular Identification of Trichoderma reesei Mohammad J. Rahimi, Feng Cai, Marica Grujic, Komal Chenthamara, and Irina S. Druzhinina Abstract Fungi comprise one of the most diverse groups of eukaryotes with many cryptic species that are difficult to identify. In this chapter, we detail a protocol for the molecular identification of the most industrially relevant species of Trichoderma—T. reesei. We first describe how a single spore culture should be isolated and used for the sequencing of the diagnostic fragment of the tef1 gene. Then, we provide two alternative methods that can be used for molecular identification and offer the diagnostic oligonucleotide hallmark of the tef1 sequence that is present in sequences of all T. reesei strains known to date and that is therefore suitable for reliable and straightforward identification. Key words DNA Barcoding, Identification phylogram, Internal transcribes spacers of the rRNA gene cluster, Sanger sequencing, Sequence similarity search, Translation elongation factor 1 alpha

1

Introduction Fungi comprise one of the most diverse groups of eukaryotes, with the estimated total number of species exceeding several millions [1]. Their simple diminutive bodies develop inside the substrate or directly on its surface, and their complex and frequently pleomorphic life cycle and superior metabolic plasticity make species identification difficult even for the taxonomists [2, 3]. However, many fungal species have unique ecophysiological properties that may either be used for applications or possess risks for agricultural plants or immunocompromised humans and other animals. Consequently, precise species identification is the crucial first step in every mycological study. Fortunately, the availability of DNA-based techniques offered the diversity of molecular tools suitable for species identification. More than a decade ago, DNA barcoding was introduced as the most reliable, simple, and precise method suitable for the identification of such highly diverse groups of organisms as bacteria, fungi, and insects [3]. DNA barcoding can

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be used to identify an organism based on the sequence of a diagnostic DNA fragment of a previously defined species. Although modern biology has experienced rapid growth of available DNA sequences through extensive whole genomic, metagenomic, or metatranscriptomic studies, the number of DNA fragments suitable for DNA barcoding remains limited. DNA loci that are appropriate for DNA barcoding should fulfill several strict criteria. First, such fragments should be universal, i.e., present in genomes of all targeted organisms. For instance, as all living cells contain ribosomes, genes encoding rRNA are suitable DNA barcoding loci. Second, these fragments should be sufficiently polymorphic as DNA barcoding requires sequence variability, and closely related species should have unequal sequences. As most protein-coding sequences in eukaryotes are aggressively conserved, they are unlikely to be used for molecular identification. Thus, DNA barcoding frequently relies on such fragments as intergenic spacers of the universal gene clusters, introns of housekeeping genes or genes present in genomes of endosymbiotic organelles (mitochondria or plastids) [4]. Third, the DNA barcoding loci should be accepted or standardized in the scientific community, and therefore respective sequences for a broad taxonomic diversity of a given group should be sufficiently represented in public databases of DNA sequences. Thus, although advances in genomics may aid the search for novel suitable DNA barcoding markers, their introduction may be inefficient if public databases do not contain sufficient reference sequences required for identification. In zoology, DNA barcoding relies on the sequence of mitochondrial genes encoding cytochrome oxidases I (COI) or II (COII) which are particularly suitable for the identification of such diverse groups as insects and other invertebrates [4, 5]. Prokaryotic organisms are most frequently identified based on the sequence of the fragments of the 16S rRNA gene; however, several additional loci are commonly used [6]. Similar to bacteria, fungi were also identified based on the sequences of genes encoding rRNA, such as the 28 S or 18 S rRNA genes [7]. However, due to an insufficient polymorphism of these loci in most fungal genera, the other region of the rRNA gene cluster was proposed for DNA barcoding. The fragment spanning from the 30 end of the 18 S rRNA gene (also known as SSU) over the internal transcribed spacer 1 (ITS1), the 5.8 S rRNA gene, the internal transcribed spacer 2 (ITS2) and ending in the 50 area of the gene encoding 28 S rRNA (LSU) is now standardized as the universal fungal DNA barcoding fragment named ITS1 and 2 of the rRNA gene cluster [3, 8]. ITS1 and 2 have numerous advantages and therefore are often used in fungal diversity survey. The highly reliable PCR primers developed almost three decades ago allow reliable and easy PCR amplification of the fragment [9]. Amplification is also facilitated by the fact that the rRNA operon has multiple copies in

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most fungal genomes, making PCR amplification typically successful. The presence of a highly conserved but short sequence of the 5.8 S rRNA gene inside the ITS1 and 2 fragments also aids multiple sequence alignment and makes the sequence similarity search reliable, which together helps identification. Consequently, public databases of nucleotide sequences have included comprehensive sets of fungal ITS1 and 2 sequences that are suitable for comparison and molecular identification. Moreover, several research communities have developed curated databases of reference ITS1 and 2 sequences for individual fungal groups [10, 11]. Nonetheless, in this chapter, we do not propose using ITS1 and 2 for molecular identification of the Trichoderma species (Hypocreales, Ascomycota). Recent studies revealed insufficient polymorphism of ITS1 and 2 in this genus, and the same phylotypes were shown to be shared by several species within large species complexes in Trichoderma [10, 12–19]. Therefore, in this genus, ITS1 and 2 can be used for the identification of most infrageneric species complexes such as clades and sections and only cautiously used for DNA barcoding of some species. The development of the field of molecular phylogeny along with several comprehensive diversity surveys has resulted in the sudden and dramatic taxonomic enlargement of several fungal genera including Trichoderma. A single species of the genus Trichoderma was discovered in 1794, and today it is comprised of more than 300 molecularly defined species, most of which were recognized based on the Genealogical Concordance Phylogenetic Species Recognition (GCPSR) method [20]. The loci used for the GCPSR method in Trichoderma include the coding fragments of housekeeping genes such as rpb2 encoding RNA polymerase subunit B II, cal and act genes encoding calmodulin and actin, respectively, as well as the partial sequence of endochitinase ech42 (chi185) and others [21]. However, the fragment spanning the fourth and the fifth introns of the tef1 gene encoding the translation elongation factor 1 alpha is widely accepted as the most polymorphic locus that may also be used for DNA barcoding (see Note 1) [9, 14]. Consequently, the sequence of tef1 is available for most molecularly recognized Trichoderma species [9]. Thus, in this chapter, we describe how to use the sequence of the tef1 gene to identify the most industrially relevant species of Trichoderma, T. reesei (see Notes 2 and 3). Although DNA barcoding is frequently introduced as an easy technique suitable for scientists without taxonomic training in a particular area, it also possesses numerous uncertainties. The sequence of the tef1 gene is more than 2000 bp long. It consists of six exons separated by five introns. For the purpose of DNA barcoding, usually a fragment of 500–600 bp is sufficient. First, Trichoderma taxonomists tested the usability of several introns in the 50 area of the gene or the short sequence including the fifth

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intron of tef1 for molecular phylogeny and later on for DNA barcoding of Trichoderma [12, 22–24]. Such attempts resulted in a collection of respective tef1 primers available in the scientific literature. However, it became apparent that the best result can be obtained if the large (fourth) intron of the tef1 gene is used [14]. Consequently, in this chapter, we describe how to amplify the tef1 gene that includes the diagnostic region and give practical advices on how to retrieve the diagnostic sequence from a sequence of the tef1 gene in Trichoderma. The most frequently used approach for DNA barcoding is based on the use of the sequence similarity search (BLAST) when the query sequence is compared to the reference sequences deposited in public databases such as NCBI GenBank. The species identity is then assigned based on the scores of the sequence similarity search, which is strongly influenced by the completeness and correctness of the reference database and by the length and composition of the query sequence. Thus, even though the DNA-based identification is considered to be the most precise, a portion of subjectivity remains also in this method. Therefore, we describe the standard procedure for the sequence similarity search that can be used to identify the T. reesei tef1 sequence. Alternatively, DNA barcoding can rely on the presence of diagnostic oligonucleotide hallmark sequences ultimately present in sequences of the DNA barcode locus of all known strains for each sequence [10]. In this chapter, we offer this diagnostic hallmark for the tef1 sequence of T. reesei.

2

Materials 1. Petri plates with solid cultivation medium [potato dextrose agar (PDA), malt extract (MEX) agar (see Note 4), or any other transparent complex solid medium (see Note 5)] supplemented with 0.1% Tween-80, and 200 mgL1 of chloramphenicol (see Note 6); the agar layer should be 2–3 mm thick. 2. Sterile cotton swabs. 3. Borosilicate glass test tubes (~2 cm in diameter) with 10–15 mL of sterile water supplemented with 0.1% Tween-80. 4. Basic microbiological equipment for sterile work including a L-shaped spatula. 5. Parafilm or any other transparent seal. 6. A basic light microscope with 10 or 20 objective (100- to 200-fold magnification). 7. Incubator.

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Fig. 1 Plates inoculated with different Trichoderma spp. (1–3) belonging to the section Longibrachiatum and secreting characteristic yellow pigment to the medium. The intensity of yellow pigment varies between species and strains. Plates were cultivated for 7 days on PDA at 25  C in darkness. The scale bar corresponds to 1 cm. T. reesei culture is shown under number 2

8. Initial culture (1–2 weeks old) with putatively mixed Trichoderma spp. cultures including a colony secreting bright yellow pigment to the medium (see Fig. 1) (see Note 7). 9. Young (36–72 h) single spore cultures of Trichoderma sp. (putative T. reesei). 10. Lysing buffer (Dilution Buffer from the Thermo Scientific™ Phire™ Plant Direct PCR Kit, F-130WH, is recommended). 11. Standard disposables and reagents for PCR. 12. Oligonucleotide primers for PCR (see below). 13. PCR thermocycler. 14. Standard equipment for agarose gel electrophoresis. 15. Standard equipment for DNA quantification. 16. Optional: Commercial kit for DNA extraction. 17. Optional: Commercial kit for PCR purification. 18. Arranged Sanger sequencing service for PCR products. 19. A partial nucleotide sequence of tef1 gene covering the large (fourth) intron, minimum 350 bp long [14]. 20. Internet connection.

3

Methods

3.1 Isolation of Single Spore Colonies of T. reesei

Steps 1–5 and 11 need to be carried out under sterile working conditions (see Note 8)

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1. Wet the cotton swab in sterile water and use it to harvest a small number of spores from the colony producing yellow pigment; one episode of contact is sufficient; avoid contact with other mold colonies. 2. Suspend spores in sterile water and mix thoroughly by gentle shaking. Do not use a vortex and avoid bubbles. 3. Prepare four sets of serial dilution in sterile water (1:10). Gently mix and label every dilution. 4. Transfer 100 μL of the 1:1000 dilution of the spore suspension to the center of the plate and spread on the entire surface of the medium using an L-shaped spatula. 5. Repeat step 4 using the 1:10,000 dilution. 6. Label and seal the Petri plates. 7. Incubate the plates at 28  C for 24 h (see Note 9). 8. Without opening the plates, use a microscope and check the plates for the presence of the single-spore colonies. It is convenient to observe plates by exposing the bottom of the plates to the microscope objective. 9. Use the thin laboratory marker and label the well-separated single-spore colonies on the bottom of the plate. 10. Incubate until the colonies become visible by naked eyes (12 h maximum). 11. Use a microbiological needle to transfer a targeted single spore colony to a fresh plate. Repeat at least five times. 12. Seal the plates with parafilm, label them, and incubate at 28  C for 70–100 h (3–4 days). 13. Note the production of the bright yellow pigment into the medium and the formation of green and slightly yellowish conidia over the entire colony (see Fig. 1) (see Note 10). 3.2 DNA Extraction, PCR Amplification, and Sequencing

1. Transfer 10–50 mg of fresh fungal mycelium (see Notes 11 and 12) to a clean 0.2 mL PCR tube and add 20 μL of lysing buffer (see Notes 13 and 14). 2. Crush the mycelium sample with a 100 μL pipette tip by gently pressing it against the tube wall. 3. Spin the fungal material down and incubate it at 98  C for 1 min. 4. Cool the mixture on ice. The supernatant can be directly used as the DNA template for PCR. 5. Prepare EF1-728F and TEF1-rev primers for the fragment of the tef1 gene (ca. 600 bp) at 10 μM (see Fig. 2) (see Notes 15–17).

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Fig. 2 (a) The map of the complete tef1 gene of T. reesei QM6a (NCBI Accession Number Z23012) showing the position of coding (orange) and non-coding (green) areas. (b) The sequences of the most common primers used for PCR amplification are provided. The dashed frame highlights the area containing the diagnostic large, the fourth intron of tef1 gene. The red bar shows the position of the oligonucleotide DNA barcode specific for T. reesei

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6. Prepare the PCR mixture on ice according to a standard protocol using following reagents: PCR components

Volume (μL)

2 Phire Plant PCR Buffer

10

EF1-728 F Primer 10 μM

1

TEF1-rev Primer 10 μM

1

Phire Hot Start II DNA Polymerase

0.5 (to be added at the end)

DNA template

0.5

Ultra-pure water

to 20

7. Run the PCR on a thermocycler: PCR step

Temperature ( C)

Time

Initial denaturation

98

5 min

30 cycles

98 55 72

5s 5s 20 s/kb

Final extension

72

1 min

Pause

4

1

8. Analyze PCR products by agarose gel electrophoresis (expected length of the product is 600 bp) (see Notes 18 and 19). 9. Use standard laboratory equipment for DNA quantification and follow the instructions of the sequencing service to prepare PCR products for Sanger sequencing (see Note 20). 3.3 Molecular Identification of T. reesei 3.3.1 By Oligonucleotide DNA Barcoding

1. Search your partial tef1 sequence for the presence of the following oligonucleotide hallmark: ATCACCCCGCTTTCT CCTACCCCTCCTTCGAGCGACGCAAA. Make sure that your sequence is not interrupted by line or paragraph breaks. 2. If your tef1 fragment contains this sequence, then your strain belongs to T. reesei sensu stricto (see Fig. 3). If the query sequence does not contain this oligonucleotide hallmark, then there is still a minor probability of it being attributed to T. reesei. In this case, perform the sequence similarity search analysis using the BLASTn algorithm available at https://blast. ncbi.nlm.nih.gov/Blast.cgi (see Subheading 3.3.2).

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Fig. 3 The location of the T. reesei DNA barcode (i.e., the diagnostic hallmark) on a multiple sequence alignment of several strains of T. reesei and the related species. Here, * and ** show sequences of T. gracile G.J.S. 10–263 and T. beinartii O.Y. 14707, respectively. The diagnostic fragment consists of 226 bp of the large, fourth intron based on the annotation of T. reesei QM6a Z23012 tef1 sequence (see Fig. 4 for details) 3.3.2 By Sequence Similarity Search Analysis Against Public Databases

Use any text editor such as Notepad and transform your sequence to FASTA format (avoid interrupting the sequence by paragraph or line breaks): >sequence title [paragraph mark] Sequence 1. Add the following sequence to your FASTA file: >DQ025754TreeseiQM6aDIAGNOSTIC AATCTTTGCCCATCTGCCCAGCATCTGGCGAAC G A AT G C T G T G C C G A C A C G AT T T T T T T T T T C AT CACCCCGCTTTCTCCTACCCCTCCTTCGAGCGACG CAAATTTTTTTTGCTGCCTTACGAGTTT TAGTGGGGTCGCACCTCACAACCCCAC TACTGCTCTCTGGCCGCTCCCCAGTCACCCAACGT CATCAACGCAGCAGTTTTCAATCAGCGATGCTAACCA TAT T C C C T C G A A C A G G A A G C C G C C G A A C T C G G CAAGGGTTCCTTCAAGTACGCGTGGGTTCTTGA CAAGCTCAAGGCCGAGCGTGAGCGTGGTATCAC CATCGACATTGCCCTCTGGAAGTTCGAGACTCC CAAGTACTATGTCACCGTCATTGGTATGTTGGCAGC CATCACC

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Example 1: >sample ATTGAGAAGTTCGAGAAGGTAAGCTTCGTTCCTT AAATCTCCAGACGCGAGCCCAATCTTTGCCCATCTG CCCAGCATCTGGCGAACGAATGCTGTGCCGACACG ATTTTTTTTTTCATCACCCCGCTTTCTCCTACCCCTCC TTCGAGCGACGCAAATTTTTTTTGCTGCCTTACGAGT TTTAGTGGGGTCGCACCTCACAACCCCACTACTGCTC TCTGGCCGCTCCCCAGTCACCCAACGTCATCAACGC AGCAGTTTTCAATCAGCGATGCTAACCATATTCCCTC GAACAGGAAGCCGCCGAACTCGGCAAGGGTTCCTTC AAGTACGCGTGGGTTCTTGACAAGCTCAAGGCCGAG CGTGAGCGTGGTATCACCATCGACATTGCCCTCTGG AAGTTCGAGACTCCCAAGTACTATGTCACCGTCATTG GTATGTTGGCAGCCATCACCTCACTGCGTCGTTGAC ACATCAAACTAACAATGCCCTCACAGACGCTCCCGGC CACCGTGACTTCATCAAGAACATGATCACTGGTACTT CCCAGGCCGACTGCGCTATCCTCATCATCGCTGCCG GTACTGGTGAGTTCGAGGCTGGTATT >DQ025754TreeseiQM6a AATCTTTGCCCATCTGCCCAGCATCTGGCGAAC G A AT G C T G T G C C G A C A C G AT T T T T T T T T T C AT CACCCCGCTTTCTCCTACCCCTCCTTCGAGCGACG CAAATTTTTTTTGCTGCCTTACGAGTTT TAGTGGGGTCGCACCTCACAACCCCAC TACTGCTCTCTGGCCGCTCCCCAGTCACCCAACGT CATCAACGCAGCAGTTTTCAATCAGCGATGCTAACCA TAT T C C C T C G A A C A G G A A G C C G C C G A A C T C G G CAAGGGTTCCTTCAAGTACGCGTGGGTTCTTGA CAAGCTCAAGGCCGAGCGTGAGCGTGGTATCAC CATCGACATTGCCCTCTGGAAGTTCGAGACTCC CAAGTACTATGTCACCGTCATTGGTATGTTGGCAGC CATCACC. 2. Copy both of the sequences to the clipboard of your computer. 3. Open an online alignment tool such as Clustal Omega (https://www.ebi.ac.uk/Tools/msa/clustalo/) or use any other alignment softwares. 4. Select “DNA” and paste your sequence in the sequence window. 5. Select “Output format” suitable for your alignment software or use “Pearson/FASTA” as a universally compatible format. 6. Click “Submit.”

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7. Your alignment in FASTA format will look like the example below: Example 2: >sample ATTGAGAAGTTCGAGAAGGTAAGCTTCGTTCCT TA A AT C T C C A G A C G C G A G C C C A AT C T T T G C C CATCTGCCCAGCATCTGGCGAACGAATGCTGTGCC GACACGATTTTTTTTTTCATCACCCCGCTTTCTCC TACCCCTCCTTCGAGCGACGCAAATTTTTTTTGCT G C C T TA C G A G T T T TA G T G G G G T C G C A C C T C A CAACCCCACTACTGCTCTCTGGCCGCTCCCCAGT CACCCAACGTCATCAACGCAGCAGTTTTCAATCAGC GATGCTAACCATATTCCCTCGAACAGGAAGCCGCC GAACTCGGCAAGGGTTCCTTCAAGTACGCGTGGGTT CTTGACAAGCTCAAGGCCGAGCGTGAGCGTGGTAT CACCATCGACATTGCCCTCTGGAAGTTCGAGACT C C C A A G TA C TAT G T C A C C G T C AT T G G TAT G T T GGCAGCCATCACCTCACTGCGTCGTTGACACAT CAAACTAACAATGCCCTCACAGACGCT CCCGGCCACCGTGACTTCATCAAGAACATGATCACT GGTACTTCCCAGGCCGACTGCGCTATCCTCATCAT CGCTGCCGGTACTGGTGAGTTCGAGGCTGGTATT >DQ025754TreeseiQM6aDIAGNOSTIC -----------------------------------------------------AATCTTTGCCCATCTGCCCAGCATCTGGCGAAC G A AT G C T G T G C C G A C A C G AT T T T T T T T T T C AT CACCCCGCTTTCTCCTACCCCTCCTTCGAGCGACG CAAATTTTTTTTGCTGCCTTACGAGTTT TAGTGGGGTCGCACCTCACAACCCCAC TACTGCTCTCTGGCCGCTCCCCAGTCACCCAACGT CATCAACGCAGCAGTTTTCAATCAGCGATGCTAACCA TAT T C C C T C G A A C A G G A A G C C G C C G A A C T C G G CAAGGGTTCCTTCAAGTACGCGTGGGTTCTTGA CAAGCTCAAGGCCGAGCGTGAGCGTGGTATCAC CATCGACATTGCCCTCTGGAAGTTCGAGACTCC CAAGTACTATGTCACCGTCATTGGTATGTTGGCAGC CATCACC-- - - - - - - - - - - - - - - - - - - - - - - - - - - - - - - - - - -----------------------------------------------------------------------------------------------------------------8. Copy these two aligned sequences in an alignment program and truncate flanking 50 and 30 areas of your sample sequence upstream and downstream of the position of “DQ025754 TreeseiQM6aDIAGNOSTIC.” The resulting area aligned to the reference fragment will result in a diagnostic region of the tef1 sequence that corresponds to the large, fourth intron and can be used for identification (see Note 21).

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9. Copy the resulting diagnostic fragment of your sample sequence to the clipboard of your computer (select “text, no gaps” option offered by most of the software programs). 10. Optional: You may wish to save the diagnostic tef1 region separately as only this tef1 fragment can be used for the analysis by sequence similarity search (BLAST). 11. Open the NCBI BLAST website https://blast.ncbi.nlm.nih. gov/Blast.cgi. 12. Select “Nucleotide BLAST.” 13. Paste your sequence. 14. Select “Nucleotide collection” database. 15. Select “blastn” program and click “BLAST.” 16. Investigate the results: T. reesei can be identified based on the high similarity to the type sequence for T. reesei QM6a Z230012 (DQ025754) and other reference sequences listed in Fig. 4. High sequence similarity is strictly defined as the combination of the following parameters: E-value ¼ 0 and similarity is >97% (“Per. Ident” or percent identity), and the Query Cover is >90%. If any of these criteria is not met, then the sequence cannot be reliably assigned to T. reesei. High similarity to other sequences may be suggestive of the attribution to the respective taxon in the case of a reliable annotation of the corresponding best hit sequences (see Notes 22–24). 17. Use the original sequence obtained from the sequencing company for deposition in a public database following the instructions on a respective website (see Note 25). 18. Prepare long-term, preservation stock cultures to maintain the culture after molecular identification (see Note 26).

4

Notes 1. Various names of the tef1 gene are available in scientific literature, tef1 is the correct name for the gene according to the nomenclature accepted for filamentous fungi; TEF1 and EF1 are the correct names for encoded proteins (Fungal Genetics Stock Center (FGSC), NEUROSPORA GENETIC NOMENCLATURE. Fungal Genetics Newsletter 46:31–41 http:// www.fgsc.net/fgn46/tofc46.htm). 2. The universal fungal DNA barcoding marker is the fragment of the rRNA gene cluster including the internal transcribed spacers 1 and 2 (ITS1 and 2 rRNA) [8], and it can only be used for the identification of infrageneric clades in

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Fig. 4 Identification phylogram showing the diversity of the diagnostic sequences of the tef1 gene in T. reesei and its closely related species. The tree was constructed using maximum parsimony algorithm based on the alignment of a 500 bp region covering the large, fourth intron from 69 strains. The numbers above the nodes correspond to bootstrap values obtained after 1000 repeats. The * indicates unpublished sequences from I. S. Druzhinina’s group. The tef1 sequences of the type strains were marked by a superscript T

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Trichoderma, but it is not suitable for the identification of all known species [10]. However, for now, all species of Trichoderma can be identified using the secondary DNA barcoding markers, i.e., the partial sequence of tef1 gene encoding translation elongation factor 1 alpha and the rpb2 gene encoding RNA Ploymerase subunite B II. 3. T. reesei and T. parareesei share the same phylotype of ITS1 and 2 rRNA sequence and cannot be distinguished [25, 26]. 4. Colonies of T. reesei cultivated on the most commonly used media such as PDA and MEX do not usually form conidial rings or pustules. 5. T. reesei does not utilize nitrate [27]. Therefore, if a synthetic cultivation medium is selected for the isolation, it should contain ammonium salts as the source of nitrogen. 6. Trichoderma spp. have different levels of resistance to antifungal compounds, and the use of fungicides for isolation of any Trichoderma species, including T. reesei, in pure culture is not recommended. 7. T. reesei shares morphological features with at least several related species such as T. parareesei and others [25, 26]. Therefore, it cannot be identified based on its phenotype. The production of yellow pigment in the medium is the characteristic feature of several species (such as T. reesei, T. parareesei, T. citrinoviride, and T. longibrachiatum) from the Longibrachiatum section of the genus [28]. This trait cannot be used for species identification, but it is suggestive for the abovementioned infrageneric group. 8. T. reesei is considered safe (GRAS status, A.B.S.A., Risk Group Database). According to safety/risk groups of microorganism classification, Trichoderma can be described as a risk group class 1 organisms (does not cause disease in healthy adult humans). The Centers for Disease Control and Prevention (CDC) and A.B.S.A. did not describe Trichoderma spp. as a risk group of microorganism. However, Trichoderma spp. are listed among opportunistic pathogens of humans, and thus, following the biosafety guidelines for handling microorganisms of the level 2 is recommended [29]. 9. T. reesei is a light insensitive or slightly photo-inhibited species [26]. Therefore, illumination conditions are not relevant for the isolation of T. reesei in pure cultures. However, darkness is recommended for preventing faster development of closely related photo-stimulated species such as T. parareesei [25, 26]. T. reesei and T. parareesei can have sympatric occurrence; therefore, the possibility of a mixed culture composed of both species cannot be excluded [25, 26].

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10. The age of the colony to obtain reasonable DNA from fungal mycelium is important because young mycelia have a thinner cell wall that is easier to break (36–72 h old mycelium in case of Trichoderma). Moreover, sampling from old mycelium with spores usually yields low DNA concentrations and high protein contents. Thus, the use of young hyphae is recommended. 11. The medium may be overlaid by sterile cellophane disks for improved sampling of hyphae. 12. If a larger amount of mycelium is used, then increase the volume of the lysing buffer (dilution buffer) to 50 μL. In any case, do not exceed 200 mg of mycelial biomass. 13. Add fungal samples into the liquid rather than onto the wall of an empty PCR tube. Make sure that you see the sample in the lysing buffer (dilution buffer). 14. There are numerous tef1 primers reported in scientific literature, which will amplify tef1 fragments of various lengths [14, 30]. In order to avoid further confusion, we do not provide a list of tef1 primers here. However, it is strongly recommended to check whether the selected primer pair will result in the amplification of the fragment that includes the large, fourth intron of the gene. Several primer pairs, such as tef1fw and tef1rev or EF1983F and EF1-2218R, or forward primers, such as EF-1002F and EF1-1-18F, will not amplify the diagnostic region and therefore should not be used for DNA barcoding. 15. Because tef1 gene is considered to be a constitutively expressed housekeeping gene, it is frequently used for the normalization of gene expression assays by quantitative PCR. Consequently, numerous primer pairs targeting usually short fragments of the large, sixth exon are available in the literature. None of them will amplify a fragment suitable for DNA barcoding for T. ressei. 16. Only one fragment of the tef1 gene sequence is suitable for DNA barcoding for T. reesei (see Figs. 2 and 3). It is critical to select the primer pair that will amplify the fragment covering the large, fourth intron sequence that can be used for T. reesei identification in this case and the fouth to fifth introns for identification of species other than T. reesei. Note that some primer pairs available for tef1 are not suitable for the DNA barcoding because they amplify the fragment that does not include a diagnostic area. No other variable regions of tef1 gene sequence can be used for DNA barcoding. 17. If the PCR does not work, then reduce the volume of the DNA template from 0.5 μL to 0.2 μL or dilute the lysate obtained at Subheading 3.2, step 3 by 1:10.

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18. If the PCR does not work, consider using commercial kits for DNA extraction or standard DNA extraction protocols. Several companies have introduced kits to extract DNA from plant tissues, but they can also be used for fungal DNA extraction. Plant/fungi DNA isolation kits offer a rapid method for the isolation and purification of total DNA from a wide range of plant and filamentous fungal species. 19. The sequencing of the PCR amplicon may result in a short fragment because the amplified tef1 region is enriched in mononucleotide polyT-stretches that may interfere with the sequencing reaction. In such cases, use the reverse primer for sequencing. Otherwise, sequencing from both directions is not necessary. 20. The detection of the diagnostic region of tef1 sequence can be done using the online tool TrichoMARK (http://www.isth. info/tools/blast/preblast.php) implemented in TrichoBLAST [14] available at www.isth.info; however, the portal is no longer supported. 21. The use of non-diagnostic fragments of the tef1 gene, such as the first introns or the last large exon, ultimately results in incorrect species identification. 22. DNA barcoding does not require phylogenetic analysis. Reliable identification of common and well-established species can be achieved based on oligonucleotide barcodes or using the sequence similarity search. 23. In the case of uncertain results of DNA barcoding, the species can be identified based on a phylogenetic analysis of the query sequence and an explicit collection of (a) most similar sequences from public databases and (b) voucher (reference) sequences of formally established taxa related to the best hits. Note that in Trichoderma and other filamentous fungi, the species can be recognized based on the GCPSR concept relying on the concordant phylogeny of at least three unlinked loci that are not contradicted by the other loci [31]. 24. If phylogenetic analysis is selected as a supplementary method for Trichoderma species identification, then maximum parsimony should be preferred over methods based on evolutionary modeling such as Bayesian phylogeny, maximum likelihood, or neighbor-joining analyses. 25. All sequences used for DNA barcoding should be deposited in public databases. It is customary to name senior scientists, faculty members, and PIs as sequence authors in order to facilitate future correspondence on sequence renaming if the name needs to be changed in future.

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26. For Trichoderma spp., the best preservation techniques include spore lyophilization or storage of a spore suspension in 25% (v/v) glycerol at a low temperature (80  C). Conidia are more stress tolerant compared to hyphae, and it is recommended to preserve conidiating (green) colonies rather than young hyphae.

Acknowledgments This work was supported by grants from the National Natural Science Foundation of China (KJQN201920), the Ministry of Science and Technology of Jiangsu Province (BK20180533), China, and the Postdoctoral Science Foundation (198162), all to FC. The work in Vienna (Austria) was supported by the Austrian Science Fund (FWF) P25613-B20 and P25745-B20, to ISD, and the Vienna Science and Technology Fund (WWTF), LS13-048, to ISD. References 1. Hawksworth DL, Lucking R (2017) Fungal diversity revisited: 2.2 to 3.8 million species. Microbiol Spectr 5 2. Begerow D, Nilsson H, Unterseher M, Maier W (2010) Current state and perspectives of fungal DNA barcoding and rapid identification procedures. Appl Microbiol Biotechnol 87:99–108 3. Xu J (2016) Fungal DNA barcoding. Genome 59:913–932 4. Fiser Pecnikar Z, Buzan EV (2014) 20 years since the introduction of DNA barcoding: from theory to application. J Appl Genet 55:43–52 5. Wilson JJ (2012) DNA barcodes for insects. Methods Mol Biol 858:17–46 6. Rossi-Tamisier M, Benamar S, Raoult D, Fournier PE (2015) Cautionary tale of using 16S rRNA gene sequence similarity values in identification of human-associated bacterial species. Int J Syst Evol Microbiol 65:1929–1934 7. Stockinger H, Kruger M, Schussler A (2010) DNA barcoding of arbuscular mycorrhizal fungi. New Phytol 187:461–474 8. Schoch CL, Seifert KA, Huhndorf S, Robert V, Spouge JL, Levesque CA, Chen W, Fungal Barcoding Consortium, Fungal Barcoding Consortium Author List (2012) Nuclear ribosomal internal transcribed spacer (ITS) region

as a universal DNA barcode marker for fungi. Proc Natl Acad Sci U S A 109:6241–6246 9. Stielow JB, Levesque CA, Seifert KA, Meyer W, Iriny L, Smits D, Renfurm R, Verkley GJ, Groenewald M, Chaduli D, Lomascolo A, Welti S, Lesage-Meessen L, Favel A, Al-Hatmi AM, Damm U, Yilmaz N, Houbraken J, Lombard L, Quaedvlieg W, Binder M, Vaas LA, Vu D, Yurkov A, Begerow D, Roehl O, Guerreiro M, Fonseca A, Samerpitak K, van Diepeningen AD, Dolatabadi S, Moreno LF, Casaregola S, Mallet S, Jacques N, Roscini L, Egidi E, Bizet C, Garcia-Hermoso D, Martin MP, Deng S, Groenewald JZ, Boekhout T, de Beer ZW, Barnes I, Duong TA, Wingfield MJ, de Hoog GS, Crous PW, Lewis CT, Hambleton S, Moussa TA, Al-Zahrani HS, Almaghrabi OA, Louis-Seize G, Assabgui R, McCormick W, Omer G, Dukik K, Cardinali G, Eberhardt U, de Vries M, Robert V (2015) One fungus, which genes? Development and assessment of universal primers for potential secondary fungal DNA barcodes. Persoonia 35:242–263 10. Druzhinina IS, Kopchinskiy AG, Komon M, Bissett J, Szakacs G, Kubicek CP (2005) An oligonucleotide barcode for species identification in Trichoderma and Hypocrea. Fungal Genet Biol 42:813–828

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Molecular Identification Beer ZW, Barnes I, Duong TA, Wingfield MJ, de Hoog GS, Crous PW, Lewis CT, Hambleton S, Moussa TAA, Al-Zahrani HS, Almaghrabi OA, Louis-Seize G, Assabgui R, McCormick W, Omer G, Dukik K, Cardinali G, Eberhardt U, de Vries M, Robert V (2015) One fungus, which genes? Development and assessment of universal primers for

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Chapter 15 In Vivo Footprinting Analysis in Trichoderma reesei Alice Rassinger Abstract The in vivo footprinting method identifies protein-targeted DNA regions under different conditions such as carbon sources. Dimethyl sulfate (DMS) generates methylated purine bases at DNA sites which are not bound by proteins or transcription factors. The DNA is cleaved by HCl, and the resulting DNA fragments are 50 -end [6-FAM]-labeled by a linker-mediated PCR (LM-PCR). Fluorescent fragments are separated and analyzed on a capillary sequencer, followed by automated data analysis using the software tool ivFAST. Key words In vivo footprint, DMS, Linker-mediated PCR, Blunt-end ligation, [6-FAM]-labeled DNA, Capillary gel electrophoresis, Peak normalization, ivFAST software tool

1

Introduction Protein–DNA interaction is one key element in gene regulation, which became a necessity to understand in respect to cellular mechanisms and their modulation. Particularly, transcription factors are responsible for the precise coordination of gene expression by binding to specific DNA-binding motifs under certain conditions, e.g., in response to different carbon sources. Hence, an unique pattern of DNA-bound proteins is left on the DNA. Already in the 1970s, specific methods for studying protein–DNA interaction were developed based on DNaseI protection [1] of the DNA, and DNA sequencing [2]. This chapter describes the in vivo and in vitro footprinting techniques in Trichoderma reesei. Both the footprinting methods share the following main steps: induced methylation of unprotected purine bases by the methylating agent dimethyl sulfate (DMS) [3], DNA fragmentation by HCl-cleavage, 50 -end fluorescent labeling by linker-mediated PCR (LM-PCR), DNA fragment analysis by a capillary sequencer, and automated data analysis using the software tool ivFAST [4]. For the in vivo footprint, DMS [5, 6] is incubated with living fungal mycelia. The agent is small enough to enter the fungal cell and is able to specifically methylate purine bases in those DNA regions that are not

Astrid R. Mach-Aigner and Roland Martzy (eds.), Trichoderma reesei: Methods and Protocols, Methods in Molecular Biology, vol. 2234, https://doi.org/10.1007/978-1-0716-1048-0_15, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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Fig. 1 Schematic overview of the linker-mediated PCR. In vivo as well as in vitro DMS-methylated DNA (black dots) is cleaved by HCl at methylated A and G bases on the coding and noncoding strand. The DNA strand is then extended by oligo 1 (1F, 1R) until the cleavage site (orange; forward primer on the left-hand side; reverse primer on the right-hand side) and is left with a blunt end. The asymmetric linker (black bars) is then ligated to the 50 end of the DNA fragments, which adds a commonly known sequence to the DNA fragments. The linkerligated fragments are amplified in several cycles of PCR using the gene-specific oligo 2 (2F, 2R; turquoise triangle) and oligo long (purple triangle), which anneals to the linker. Lastly, a fluorescent label (dark blue dot) is introduced by oligo 3 (3F, 3R; light blue triangle) in a separate PCR and creates single-stranded 50 -[6-FAM]labeled DNA fragments of various lengths and a common linker sequence

blocked by proteins (see Fig. 1). Thereby, the induced in vivo methylation by DMS mimics accessible elements of DNA. The methylated DNA is specifically cleaved by HCl at the methylated A and G bases and creates blunt-ended DNA fragments of various lengths. A linker-mediated PCR is used to amplify those DNA fragments with the help of a linker [7], followed by a 50 -end fluorescent labeling reaction using a 50 -[6-FAM]-labeled primer. The 50 -[6-FAM]-labeled DNA fragments are separated by size and are analyzed on a capillary sequencer. Fragment analysis and sequence data analysis are achieved automatically using the software tool ivFAST. The program performs a normalization of the generated in vivo footprinting data and shows statistical differences at specific DNA bases between different conditions (e.g., carbon sources) as a color-coded heatmap [4].

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Materials

2.1 In Vivo Methylation

1. 2-(N-Morpholino)ethanesulfonic acid (MES) buffer: 200 mM MES, pH 5.5. Weigh in 39.1 g of MES (Mw ¼ 195.24 g/L) and add 900 mL of distilled water. Stir with a magnetic stirrer until dissolved. Measure the pH and adjust with NaOH if necessary. Fill the solution into a measuring cylinder and add distilled water up to 1 L. Store at 4  C. Do not store longer than 1 month. 2. TLEβ-buffer: 10 mM Tris–HCl, pH 8.0, 300 mM LiCl, 1.0 mM EDTA, pH 8.0, 2% (v/v) β-mercaptoethanol. Prepare higher concentrated stock solutions for each solid compound (1 M Tris–HCl, pH 8.0, 0.5 M EDTA, pH 8.0, 1 M LiCl). Combine the stock solutions in appropriate volumes for the respective end concentrations. Fill up to 980 mL, add a magnetic stirrer bean and mix for 5 min, then leave the stirrer bean in the solution and autoclave at 120  C for 20 min. Before use, add 20 mL of β-mercaptoethanol (final volume is now 1 L) to the sterilized and cooled solution and let it stir for 10 min. Store at 4  C. 3. Methylation agent: Dimethyl sulfate (DMS). Commercially available as liquid.

2.2

DNA Extraction

1. DNA extraction buffer: 0.1 M Tris–HCl, pH 8.0, 1.2 M NaCl, 5 mM EDTA, pH 8.0. 2. Phenol, buffered, pH 8.0. 3. Chloroform. 4. Isoamylalcohol. 5. Isopropanol. 6. 10 mg/mL RNaseA. 7. 70% (w/v) ethanol. 8. 5 mM Tris–HCl, pH 7.5. 9. 0.8% (w/v) agarose gel in TAE buffer. 10. Xylene cyanol as (6) DNA loading dye. 11. GeneRuler 50 bp DNA ladder (Thermo Fisher Scientific, Waltham, MA, USA).

2.3 In Vitro Methylation

1. DMS reaction buffer: 0.05 M sodium cacodylate, 1 mM EDTA, pH 8.0. Adjust pH with sodium hydroxide or cacodylic acid. Store at 4  C. 2. Stop solution: 1.5 M sodium acetate, pH 7.0, 1.0 M β-mercaptoethanol. Store at 4  C. 3. Precipitation agent: 3 M sodium acetate, pH 5.0, 96% (w/v) ethanol.

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Acidic Cleavage

2.5 Fragment Recovery

1. Hydrochloric acid: 0.5 M HCl in distilled water. 1. Precipitation agent: 3 M sodium acetate, pH 5.0, 96% (w/v) ethanol. 2. Sodium hydroxide: 1 M NaOH in distilled water. 3. Hydrochloric acid: 1 M HCl in distilled water. 4. Washing ethanol: 70% (w/v) ethanol. 5. Resuspension buffer: 1 M and 10 mM Tris–HCl, pH 7.5. Prepare a 1 M Tris–HCl, pH 7.5 solution in distilled water and dilute to 10 mM with distilled water. 6. Purification of fragments: Use a commercial kit for the cleanup. Preferably, use the GeneJET PCR Purification Kit (Thermo Fisher Scientific, Waltham, MA, USA) or any similar commercial kit (see Note 1). 7. 1% (w/v) agarose gel in TAE buffer. 8. Xylene cyanol as (6) DNA loading dye. 9. GeneRuler 50 bp DNA ladder (Thermo Fisher Scientific, Waltham, MA, USA).

2.6

LM-PCR

1. Primers and probes: Dissolve primers in sterile bidistilled (sb) water to a stock concentration of 100 μM. Dissolve probes in 10 mM Tris–HCl, pH 7.5 prepared with sb water to a concentration of 100 μM. Dilute oligonucleotides for the elongation in either forward or reverse direction, oligo 1 forward (fwd) and oligo 1 reverse (rev), respectively, with sb water to a working stock concentration of 0.33 μM. Dilute the oligonucleotides for the amplification (also referred to as oligos 2 fwd and rev) to 12.5 μM in sb water. Keep the dilutions of the 50 -[6-FAM]-labeled oligonucleotides (also referred to as oligos 3 fwd and rev) in 10 mM Tris–HCl, pH 7.5 and a working concentration of 12.5 μM. Store at 20  C. 2. Asymmetric Linker: Mix 66 μL of oligo long (1 μg/μL, 50 -GCGGTGACCCGGGAGATCTGAATTC-30 ), 29 μL of oligo short (1 μg/μL, 50 -GAATTCAGATC-30 ), 100 μL of 1 M Tris–HCl, pH 7.7 (prepared with sb water), and 205 μL of sb water. Prepare four times 100 μL of aliquots in 0.2 mL PCR tubes and incubate for 5 min at 95  C. Let them gradually cool down to 30  C in a PCR cycler (0.01  C/s). Store aliquots at 20  C. 3. Other components: 2000 U/mL Vent-polymerase (New England Biolabs, Ipswich, MA, USA), 10 reaction buffer (New England Biolabs, Ipswich, MA, USA), 2 mM dNTP mix, T4 DNA Ligase, 10 Ligase buffer, 1 mg/mL tRNA from baker’s yeast.

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4. 1.2% (w/v) agarose gel in TAE buffer (see Note 2). 5. Xylene cyanol (XC) as (6) DNA loading dye. 6. GeneRuler 50 bp DNA ladder (Thermo Fisher Scientific, Waltham, MA, USA). 7. Typhoon FLA 9000 laser scanner (GE Healthcare, Chicago, IL, USA). 2.7 Fragment Analysis

1. Bidistilled water as diluent. 2. Capillary gel electrophoresis using an ABI 3730XL Genetic Analyser (Life Technologies, Carlsbad, CA, USA) using GeneScanTM 600-LIZ as internal size standard (Life Technologies, Carlsbad, CA, USA). 3. Peak Scanner™ Software v1.0 (Life Technologies, Carlsbad, CA, USA). 4. In vivo footprinting software tool (ivFAST).

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Methods

3.1 In Vivo Methylation

1. Prepare for each culture a 2 mL reaction tube with 2 mL of MES buffer. Add 2 μL of DMS per milliliter of culture to the MES buffer and mix vigorously (see Note 3). 2. Add the prepared DMS-MES buffer to the culture, distribute well by gently shaking the flasks, and incubate at the same temperature as the cultivation for 2 min. After incubation, proceed at room temperature. 3. Fill up the shaking flask with 5 volumes of the culture volume with ice-cold TLEβ buffer to stop the methylation reaction. Filter the mycelia over miracloth, wash the mycelia with water, dry press them between the sheets of Whatman paper, and freeze in liquid nitrogen. Store frozen mycelia until use at 80  C.

3.2

DNA Extraction

1. For each sample and DNA extraction, prepare a 2 mL reaction tube filled with 0.8 mL of DNA extraction buffer. 2. Homogenize the frozen mycelia in liquid nitrogen with mortar and pestle until a fine powder is obtained. 3. With a spatula, transfer roughly 50–60 mg to the tube containing the DNA extraction buffer and vortex until the mycelium is well resuspended without any clumps. 4. Add 1 volume of phenol–chloroform–isoamylalcohol (25:24:1) and mix gently. Let it set for 1 min and centrifuge at 4  C and 15,000  g for 10 min. Transfer the supernatant (roughly 750 μL) to a new 2 mL reaction tube and add again 1 volume of phenol–chloroform–isoamylalcohol. Repeat the

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centrifugation step. Add 1 volume of chloroform to the supernatant and mix. Repeat the centrifugation step as above. Transfer supernatant (roughly 650 μL) in a 1.5 mL reaction tube. 5. Treat the supernatant with 65 μL of RNaseA (10 mg/mL) for 30 min at 37  C. 6. Precipitate the DNA with 1 volume of isopropanol, mix by inversion, and incubate for 15 min at room temperature. Centrifuge at 20,000  g and 4  C for 15 min. Remove the supernatant and wash with 1 volume of 70% ethanol. Centrifuge again at 15,000  g and 4  C for 5 min and discard the washing ethanol. Use a pipette tip to remove the residual ethanol. 7. Let the DNA pellet dry at 50  C on a thermoblock for 2–5 min until the ethanol is evaporated (see Note 4). 8. Dissolve the pellet in 200 μL of 5 mM Tris–HCl, pH 7.5 and incubate at 50  C and 750 rpm on a thermoblock. The DNA is stored at 4  C until use. 9. Load 2 μL of DNA mixed with DNA loading dye onto a 0.8% (w/v) agarose gel prepared with TAE buffer and run at 80 V for 45 min (see Fig. 2a).

Fig. 2 Gel electrophoresis pictures of genomic, HCl-cleaved and labeled DNA. (a) The extracted genomic DNA from fungal mycelia shows no degradation or any RNA contaminations at the bottom of the gel. (b) The HCl-cleaved DNA shows a fragment size distribution ranging from 500 up to 900 bp if compared to the marker (M). (c) The 50 -[6-FAM]-labeled DNA fragments shown here are generated by the oligo set for cbh2 forward. Lanes 1–3 as well as 4 and 6 show successfully labeled DNA fragments. The DNA fragments show an even size distribution, which are suitable for genotyping and in vivo footprinting data analysis. As a negative example, lane 5 shows no labeled DNA fragments. Only the labeled probe (i.e., oligo 3 fwd) is visible at the bottom of the gel in lane 5. The marker (M) is not visible on the gel

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1. Use 100 μL (a total amount of 100 μg) of DNA prepared in Subheading 3.2. 2. Add 400 μL of DMS reaction buffer and 2 μL of DMS (1:20 dilution) and mix vigorously until the DMS is completely dissolved. 3. Combine the 100 μL of DNA and the DMS reaction buffer, mix well, and incubate at room temperature for 5 min. 4. Add 80 μL of stop solution and mix gently. 5. Precipitate the DNA with 0.1 volumes of precipitation agent and 1.5 volumes of 96% ethanol and incubate at 80  C for 10 min. 6. Recover the DNA pellet by centrifugation at 4  C and 14,000  g for 15 min and dissolve in 100 μL of 10 mM Tris–HCl, pH 7.5. 7. Repeat the precipitation and centrifugation step. 8. Wash the pellet with 1 mL of 70% ethanol and centrifuge at 4  C and 14,000  g for 10 min. 9. Let the pellet dry at 50  C for 2 min until the residual ethanol evaporated and dissolve the pellet in 100 μL of 10 mM Tris– HCl, pH 7.5. Store at 20  C until use.

3.4 Acidic Cleavage and Fragments Recovery

1. Add 6.25 μL of 0.5 M HCl to 100 μL of methylated DNA, mix gently, and incubate for 1.5 h on ice. 2. Add 143.75 μL of sb water to the sample and proceed with a sodium acetate/ethanol precipitation as described in Subheading 3.3, steps 5 and 6. 3. Dissolve the DNA pellet in 250 μL of sb water, add 10 μL of 1 M NaOH, mix gently, and incubate at 90  C on a thermoblock for 30 min. 4. Allow the sample to cool down on ice for 2 min and spin down. Add 25 μL of 1 M Tris–HCl, pH 7.5 and mix gently. 5. Neutralize the pH of the sample with 1 M HCl (see Note 5). 6. Repeat the precipitation, centrifugation, and washing as above, described in Subheading 3.3, steps 5 and 6. 7. Dry the pellet at 50  C on a thermoblock for 2–5 min until the residual ethanol is evaporated and resuspend in 100 μL of 10 mM Tris–HCl, pH 7.5. 8. Purify the DNA by using the GeneJET PCR Purification Kit. Elute in two steps: the first elution with 50 μL of the supplied elution buffer, then with 60 μL of elution buffer. Elute into the same tube. 9. Store at 4  C until use. 10. Prepare a 1% (w/v) agarose gel and load 5 μL of sample plus DNA loading dye. Run the gel at 80 V for 40 min (see Fig. 2b).

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Table 1 Overview of the primer design for in vivo footprinting Overlapping sequence

Oligo name

Coding strand

Noncoding strand

LM-PCR step

Number of GCs

Oligo 1

Rev

Fwd

Elongation

10

Oligo 2

Rev

Fwd

Amplification

13

x

Oligo 3

Rev

Fwd

Labeling

15

x

3.5

LM-PCR

3.5.1 Primer Design

Appropriate regions for protein–DNA interactions by in vivo footprinting have an approximate size of 350 bp without taking the flanking primer sequences on the coding and the noncoding strand (i.e., 50 -[6-FAM]-labeled oligo 3 rev and fwd, respectively) into account. The elements of interest such as transcription factor binding sites should be located in the middle of the 350 bp region. 1. Design three oligonucleotides (also referred to as oligos) for the coding (reverse oligo set) and noncoding strand (forward oligo set). 2. The numbering of the oligonucleotides indicates the chronological order of the use of the respective oligos: oligo 1 fwd/rev is used in the elongation step of the LM-PCR, oligo 2 fwd/rev in the amplification step and oligo 3 fwd/rev is a 50 -[6-FAM]labeled oligonucleotide used for the labeling reaction. 3. The three oligos for each strand differ in the number of GC bases and hence in the annealing temperature: 10 GC bases are present in oligo 1, 13 GCs in oligo 2, and oligo 3 has 15 GCs. 4. Design oligo 1 and oligo 2 without any overlaps in their 50 -30 sequences. A minimum distance of 10 bp is necessary between both oligonucleotides. The distance should not exceed 50–60 bp. 5. Oligo 2 and oligo 3 share an overlapping sequence of 15 bp, but the oligos differ in 2–5 bp. An overview of the primer design can be seen in Table 1.

3.5.2 PCR

To keep each PCR as homogenous as possible, work with master mixes for each LM-PCR step: elongation, ligation of linker, amplification, and labeling reaction (as summarized in Fig. 1). Prepare all reaction mixtures on ice and use bidistilled water for all reactions. 1. For the elongation reaction, combine the following reagents (final volume 30 μL): 5 μL of HCl-treated and purified DNA (prepared in 3.4), 3 μL of 10 reaction buffer, 1 μL of oligo 1 (forward or reverse), 3 μL of 2 mM dNTPs, 0.5 μL of Vent polymerase. Mix gently. On a PCR cycler, perform the

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following program: 95  C for 5 min, 55.5  C for 30 min, 75  C for 10 min, 4  C for 15 min. Keep the samples on ice until the next step. 2. Add the ligation mix to the elongated samples: 4 μL of 10 ligase buffer, 4.5 μL of linker, 1.5 μL of T4 DNA ligase. Mix gently (see Note 6). Incubate the samples at 17  C for 17 h in a PCR cycler. Precipitate the DNA by adding 10 μL of tRNA, 4 μL of sodium acetate, and 120 μL of 96% ethanol and incubate for 15 min at room temperature. Centrifuge at 14,000  g, 4  C for 15 min. Remove the supernatant carefully with a pipette tip and let the pellet air-dry. Dissolve the pellet in 10 μL of 10 mM Tris–HCl, pH 7.5 and incubate for 10 min at 50  C on a thermoblock. Keep the samples on ice until the next step. 3. Combine 10 μL of linker-ligated DNA, 2.5 μL of 10 reaction buffer, 2.5 μL of 2 mM dNTPs, 0.4 μL of oligo 2 (forward or reverse), 0.4 μL of oligo long (1:10 diluted in bidistilled water), 0.5 μL of Vent polymerase for amplifying the DNA fragments. Mix gently. The cycling conditions are the following: initial denaturation at 95  C for 2.5 min; 17 cycles of 95  C for 1 min, 60.5  C for 2 min, 75  C for 3 min; a final cooling step at 4  C for 20 min. 4. For the labeling reaction, add to the samples 0.4 μL of oligo 3 (forward or reverse), 0.5 μL of reaction buffer, 3.6 μL of 2 mM dNTPs, 0.5 μL of Vent polymerase. Mix gently and perform the following PCR program: initial denaturation at 95  C for 2.5 min; 5 cycles of 95  C for 1 min, 63.5  C for 2 min, 75  C for 3 min; a final cooling step at 4  C for 20 min. 5. Prepare a 1.2% agarose gel without any DNA dye. Use 5 μL of [6-FAM]-labeled DNA plus XC loading dye and load onto the gel. Use a 50 bp DNA ladder as size standard. Run the gel at 90 V for 45 min. 6. Scan the gel on a Typhoon FLA 9000 laser scanner at 473 nm and 850 V (see Fig. 2c). If the bands appear oversaturated, the excitation voltage should be reduced to 700 V. 3.6 Fragment Analysis 3.6.1 Capillary Gel Electrophoresis (CGE) 3.6.2 Data Analysis

For the CGE analysis, mix 4 μL of the [6-FAM]-labeled DNA with 16 μL of sb water. The labeled DNA fragments are separated and analyzed by Microsynth (https://www.microsynth.ch/home-ch. html). Detailed instructions of the use of ivFAST are found in the user’s manual in the download folder of the software tool. The following section gives hands-on instructions of the program, which are not explicitly mentioned in the user’s manual.

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Fig. 3 Electropherogram of the CGE separated DNA fragments of the cbh2 promoter using the forward oligo set, which yields the 50 -[6-FAM]-labeled coding strand. The dashed line indicates primer sequences and not distinct peaks. The triangle marks the manually identified beginning of the sequence and shows distinct peaks. The peaks in the chromatogram represent an A or G, and the sequence (A/G in blue) is mapped to the peaks accordingly

1. Download ivFAST: https://www.vt.tuwien.ac.at/ biochemische_technologie/synthetische_biologie_und_ molekulare_biotechnologie/. 2. Create a folder within the ivFAST folder, in which the input files are saved for the ivFAST analysis. 3. From data provided by the DNA fragment analysis (Microsynth), open the .fsa-files in Peak ScannerTM Software v1.0 (see Note 7). 4. Specify the start and end of the sequence in the electropherogram and omit the primer sequence at the very beginning of the chromatogram (see Fig. 3). Save the sequence in FASTA format as .txt-file and in 50 to 30 orientation of the coding strand in the folder, created within the ivFAST folder. 5. For each sample and replicate, prepare three separate .txt-files and name them accordingly to the user’s manual instructions. One of the three files contains only the specified sample peaks (p), which are determined manually based on the electropherograms (as described in step 4.). Additionally, prepare an all peaks sample file (a) and a standard file (s), containing the peaks of the internal size standard supplied by the sequencing service. 6. Save the sequence text file, which maps to the specified peaks, the specified sample peak file (p), the standard file (s), and the all peaks sample file (a) in the folder created within the ivFAST folder.

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7. Open the command line and direct to the location of ivFAST. jar using the Windows cmd.exe or Macintosh terminal.app. 8. Start the program:[space]java[space]-jar[space]ivFAST.jar [space]-d[space](name of the created folder containing three peak files and sequence as .txt-files written without the round brackets) and press enter. 9. Specify the start (-s) position of the sequence (which is 1, since the first specified sample peak maps to the first base in the sequence), the start position of the sequence where the plotted heatmap (-sd) should begin and the sequence position where the plotted heatmap should end (-e). Then enter which conditions should be compared (e.g., 1/0 if “1” is condition 1 and “0” is condition 2) as instructed by the ivFAST program and press enter. 10. An output folder is generated in the ivFAST folder. Open the mapping .txt-files and check if the assigned A or G base correlates with a high mass value, since the peaks in the electropherogram resemble A or G bases. If a high peak area is assigned incorrectly to a C or T instead, it can be due to mathematical rounding, e.g., statistical mass differences between two bases range from 1.0 to 1.4. In case of 1.5, the difference between assigned peaks is 2 and the sequence mapping jumps to the next base and not the immediately consecutive base. Change the peak’s mass value manually in the .txt-file (p) so that a difference of 1 is still maintained. 11. Re-run the ivFAST java-file and re-check again the mapping files. 12. Open the created heatmap to review differently targeted bases between different conditions (see Fig. 4).

Fig. 4 Heatmap of a cbh2 promoter footprint. The heatmap was generated from 3 replicates for each condition, which are compared to each other. The displayed data is statically tested and normalized. The user defines which conditions are compared (here carbon sources; G, glucose; S, sophorose; N, no carbon source) to each other and those are displayed as “sample/reference.” The colored boxes indicate protection or hypersensitivity. The shade intensity of the color represents the significant differences ranging from 1.1- to 1.5-fold (from l light shaded to dark shaded) as defined in the config file of the ivFAST program. From the genotyping of the forward primed footprint, the electropherograms corresponds to the 30 –50 sequence of the noncoding strand and is displayed in reverse orientation of the non-coding strand below the heatmap

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Notes 1. The choice for the commercial PCR clean-up kit should be made on the specifications of the product. The recovery rate of the fragment sizes from 100 bp up to 1000 bp should be at least 95%. Within one in vivo footprinting experiment, do not apply two different commercial kits to your sample set and controls. 2. Rinse the gel electrophoresis chamber carefully with water and soap. Any traces of DNA dyes (e.g., ethidium bromide, Midori Green) should be removed before preparing the DNA dye-free agarose gel. Traces of DNA dyes interfere with the fluorescence signal from your [6-FAM]-labeled DNA fragments. An indicator for contamination with DNA dye is the visual appearance of the DNA ladder on the gel. It is only visible in the fluorescence scan if the DNA dye stained it. 3. The DMS should be evenly distributed and eventually dissolved in the MES buffer. It can take several minutes of vigorous mixing (vortexing). 4. Do not incubate too long at 50  C. The pellet should not be over-dried. Otherwise the DNA can be damaged and decreases the quality of the footprint. Also, if the pellet is not dried long enough, ethanol remains in the sample. This decreases the consecutive PCR efficiency in the LM-PCR. 5. For neutralization, 10–13 μL of 1 M HCl is sufficient. Add 5–10 μL of 1 M HCl to the samples, mix gently, and check the pH by pipetting 1 μL of sample onto pH paper. Gradually adjust the volume of 1 M HCl added to the samples until the pH is neutral. 6. If the asymmetric linker for the linker ligation is freshly prepared and is directly used afterwards in the ligation reaction, it can be stored on ice until used. Otherwise, the frozen linker aliquot should be thawed on ice and then used. 7. Check the quality of the chromatograms of each sample and replicate. If the peaks are baseline separated and sharp, the file can be used for further data analysis. Otherwise, the in vivo footprint has to be repeated.

Acknowledgments This work was funded by two grants from the Austrian Science Fund (FWF): V232 and P24851 given to Astrid Mach-Aigner.

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References 1. Jackson PD, Felsenfeld G (1985) A method for mapping intranuclear protein-DNA interactions and its application to a nuclease hypersensitive site. Proc Natl Acad Sci U S A 82:2296–2300 2. Galas DJ, Schmitz A (1978) DNAase footprinting: a simple method for the detection of protein-DNA binding specificity. Nucleic Acids Res 5(9):3157–3170 3. Pfeifer GP, Tanguay RL, Steigerwald SD, Riggs AR (1990) In vivo footprint and methylation analysis by PCR-aided genomic sequencing: comparison of active and inactive X chromosomal DNA at the CpG island and promoter of human PGK-I. Genes Dev 4 4. Gorsche R, Jovanovic B, Gudynaite-Savitch L, Mach RL, Mach-Aigner AR (2013) A highly

sensitive in vivo footprinting technique for condition-dependent identification of cis elements. Nucleic Acids Res 42(1):e1. https:// doi.org/10.1093/nar/gkt883 5. Ephrussi A, Church GM, Tonegawa S, Gilbert W (1985) B lineage-specific lineage interactions of an immunoglobin enhancer with cellular factors in vivo. Science 227:134–140 6. Wolschek MF, Narendja F, Karlseder J, Kubicek CP, Scazzocchio C, Strauss J (1998) In situ detection of protein-DNA interactions in filamentous fungi by in vivo footprinting. Nucleic Acids Res 26(16):3862–3864 7. Mueller PR, Wold B (1989) In vivo footprinting of a muscle specific enhancer by ligation mediated PCR. Science 246(4931):780–786

Chapter 16 RNA Characterization in Trichoderma reesei Petra Till Abstract This chapter provides an overview on different methods for the characterization of RNAs in Trichoderma reesei. In the first section, protocols for the extraction of total RNA from fungal mycelia and the identification of 50 and 30 ends of certain RNAs of interest via rapid amplification of cDNA ends (RACE) are presented. In the next section, this knowledge on the transcriptional start and end points is used for in vitro synthesis and fluorescence labeling of the RNA of interest. The in vitro synthesized RNA can then be applied for in vitro analyses such as RNA electrophoretic mobility shift assays (RNA-EMSA) and RNA in vitro footprinting. RNA-EMSA is a method suitable for the identification and characterization of RNA–protein interactions or interactions of an RNA with other nucleic acids. RNA in vitro footprinting allows exact mapping of protein-binding sites on RNA molecules and also the determination of RNA secondary and tertiary structures at singe-nucleotide resolution. All protocols presented in this chapter are optimized for the analysis of noncoding RNAs (ncRNAs), especially long ncRNAs (lncRNAs) or other specific RNA species of more than 200 nt in length. Key words Trichoderma reesei, RNA, In vitro synthesis, Electrophoretic mobility shift assay, In vitro footprinting, RACE, 50 /30 end determination, RNA–protein interaction, RNase assay, RNA structure

1

Introduction

1.1 Rapid Amplification of cDNA Ends (RACE)

RACE is the most commonly used technique for the identification of mRNA 30 and 50 ends. For coding transcripts, which are translated into proteins, an exact definition of transcriptional start and end points might be of interest as noncoding terminal regions might contain regulatory signals (i.e., signals that have an impact on transcription initiation and translation, post-transcriptional processing, stability, or localization of the mRNAs [1, 2]). Anyway, this chapter is focused on the characterization of noncoding RNAs (ncRNAs), especially long ncRNAs (lncRNAs), for which an exact knowledge on the 30 and 50 ends is essential. As ncRNAs differ from coding transcripts in several regards, adaptions of the classical RACE protocol are required. RACE was first described by Frohman in 1988 [3]. The underlying mechanism is illustrated in Fig. 1. For defining the 30 end of a

Astrid R. Mach-Aigner and Roland Martzy (eds.), Trichoderma reesei: Methods and Protocols, Methods in Molecular Biology, vol. 2234, https://doi.org/10.1007/978-1-0716-1048-0_16, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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Fig. 1 Schematic illustration of 30 RACE (left) and 50 RACE (right). The RNA of interest (red) is reversely transcribed into cDNA (blue) and the terminal sequence is amplified in two PCR steps (black) based on the application of gene-specific primers (GSP) and an oligo-dT primer linked to an anchor sequence (Anchor). Primers are highlighted in bold. The direction of amplification is indicated by arrows

target transcript via classical 30 RACE (see Fig. 1, left), the RNA is reversely transcribed based on its poly(A)-tail using an oligo-dT primer linked to an anchor sequence. In a second step, the target sequence is amplified from the obtained cDNA via PCR applying a gene-specific primer and a primer binding to the anchor sequence. Specificity of the product is increased in a second amplification step applying a nested gene-specific primer and the anchor primer. For defining the 50 end of a target transcript via 50 RACE (see Fig. 1, right), the RNA is reversely transcribed using a gene-specific primer oriented in the reverse direction. In order to generate an anchor point for the 50 RACE, a poly(A)-tail is attached to the 30 end of the obtained cDNA. In a next step, the target sequence is amplified via PCR applying a nested gene-specific primer and an oligo-dT primer linked to an anchor sequence. Specificity of the product is increased in a second amplification step, applying a further nested genespecific primer and the anchor primer [3]. While the 50 RACE starts with a gene-specific reverse transcription step, the first step of 30 RACE yields a mixture of unspecific products resulting from reverse transcription of all poly(A)-tailed RNAs. This is relevant for the analysis of ncRNAs in two regards. First, the classical approach of 30 RACE requires the presence of a poly(A)-tail at the 30 end of the transcripts to be analyzed. Importantly, this is an optional feature for ncRNAs [4, 5]. Second, several ncRNAs such as lncRNAs are usually expressed at very low levels compared to (even poorly expressed) mRNAs [6, 7]. Therefore,

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transcripts of interest might get lost in the pool of highly abundant templates for the unspecific reverse transcription in the first step of 30 RACE. Moreover, PCR-based methods might be hindered by high-ordered secondary structures formed by some ncRNA species such as lncRNAs [8]. In this section, strategies to overcome these problems for the identification of ncRNA ends via RACE are outlined. 1.2 RNA In Vitro Synthesis and Labeling

Labeled or unlabeled in vitro synthesized RNA might be used for several applications such as function and interaction studies, structure analysis, microarray analysis, RNAi and anti-sense RNA experiments, RNA transfection, microinjection, in vitro translation, as hybridization probes for RNase protection assays or as functional molecules (e.g., ribozymes or aptamers). Kits for RNA in vitro synthesis are commercially available and make use of components of the bacteriophage systems. The T7 High Yield RNA Synthesis Kit presented in this section uses the T7 RNA polymerase from the T7 phage of E. coli, which is highly specific for its 23 bp promoter sequence [9]. It produces high yields and is suitable for in vitro transcription of RNAs ranging from less than 30–10,000 nt [10]. In this section we emphasize the production, purification, and analysis of RNAs of 200–400 nt in length which can be used for RNA-EMSAs or in vitro footprinting as described in the following sections. For some applications, labeling of the RNA is required. Radiolabels or fluorescence labels can be attached at the 50 end (catalyzed by the T4 polynucleotide kinase) and the 30 end (catalyzed by the T4 RNA ligase) or incorporated into the RNA body during in vitro synthesis [11]. Here we present a protocol for linking CY5 or CY3 labels to the 30 end of in vitro synthesized RNA using T4 RNA ligase. Other labeling strategies might be preferred depending on the properties of the RNA and the final application.

1.3 RNA Electrophoretic Mobility Shift Assay (RNA-EMSA)

The classical electrophoretic mobility shift assay (EMSA) evolved as a common and simple technique for studying DNA–protein interactions in vitro in a qualitative and quantitative manner [12, 13]. It can be used to analyze the relative affinity of a protein for one or more DNA target sites or, vice versa, to compare the affinities of different protein interaction partners for the same DNA target site [14]. Binding constants can be determined [14, 15]. Also higherordered complexes composed of different interaction partners can be investigated [16]. The method relies on the principle that DNA–protein complexes are larger and thus retarded during non-denaturing PAGE compared to unbound DNA [12, 13]. Approaches of the DNA with and without the protein are analyzed in parallel. They are separated on a native PAA gel in an electric field, and the DNA is visualized. The free DNA yields a signal at a certain position, while

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the protein–DNA complex appears as a discrete band shifted upwards on the gel. Formerly, visualization of the DNA was achieved by the application of radiolabeled nucleic acid fragments [12, 13], yet meanwhile also fluorescence-based imaging strategies have been established. Both strategies have been successfully conducted for studying DNA–protein interactions in T. reesei [17– 19]. Cell-free extracts as well as different purified proteins were applied, and binding to various DNA target sites was analyzed. However, no EMSA studies including RNA interaction partners have been performed in T. reesei. Here, we present a protocol for studying RNA interactions in T. reesei. The preparation of a 4% native PAA gel (30:0.36 acrylamide/bisacrylamide) and its application for RNA-EMSA analysis is described. The procedure is optimized for studying interactions of RNAs of 200–400 nt in length and proteins of approximately 100 kDa in size. However, also smaller or larger RNAs and proteins, other interaction partners, or high-ordered complexes may be analyzed. RNAs can form complexes with proteins as well as with nucleic acids, i.e., DNA or other RNAs. For studying RNA–DNA interactions using short dsDNA oligonucleotides, we recommend modifying the protocol for studying DNA–protein interactions described previously [17], as small oligonucleotides are not visible on the 4% PAA gel scheduled in this section. Yet, the current protocol might be adapted for RNA-EMSA studies with larger DNA or RNA interaction partners or for supershift experiments. Anyway, when studying interactions including more than one nucleic acid, labeling of the RNA and/or DNA is required in order to allow the distinction of the signals. A protocol for CY5-labeling of in vitro synthesized RNA is provided in Subheading 3.2.4. For ordinary RNA–protein interaction studies, unlabeled in vitro synthesized RNA is sufficient. An example for a classical RNA-EMSA is depicted in Fig. 2. 1.4 RNA In Vitro Footprinting

In vitro footprinting is a method suitable for defining the exact position of a protein-binding site on a DNA or RNA molecule [20, 21]. Moreover, it allows investigations on RNA secondary and tertiary structures at single-nucleotide resolution [22] and the analysis of ligand-induced conformational changes [23, 24]. The technique is based on the idea that RNA regions bound by a protein or regions forming an RNA duplex in highordered structures are protected from nuclease cleavage or chemical attacks. The concept originates from a nuclease protection assay published by Steitz in 1969 [25] and thereafter evolved as the classical footprinting technique for studying DNA–protein or RNA–protein interactions [20, 21]. For RNA–protein interaction studies, approaches of the unbound RNA and the RNA incubated with the protein are compared. Both of the approaches are treated with chemic or enzymatic reagents, which result in modification or

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Fig. 2 Result of an RNA–protein interaction study via RNA-EMSA. The signal of a free RNA (lane 1) is compared to RNA supplemented with increasing amounts of a protein (lanes 2-8). A 0.25- to four-fold molar excess of the 105 kDa protein relative to 1 μg of the in vitro synthesized 262-nt-long RNA was used

cleavage, respectively, of the RNA at accessible positions. The most commonly used chemical reagent is DMS, which causes methylation of the base-pairing sites of adenine and cytosine residues and of N-7 of guanidines [26]. The modified RNAs are finally analyzed by primer extension using radiolabeled or fluorescence-labeled primers (inhibition of reverse transcription at modified adenines and cytosines) [27–29] or by chemical treatment with sodium borohydride/aniline or hydrazine/aniline (strand scission at modified guanidines or cytosines, respectively) [22]. For footprinting based on enzymatic reactions, the treatment of an end-labeled RNA with enzymatic reagents yields RNA fragments of different sizes. Analysis of the patterns of end-labeled RNA fragments via denaturing gel electrophoresis or sequencing finally allows conclusions on the RNA structure and mapping of protein-binding sites. Various nucleases with different sequence or structure specificities have been employed for enzymatic footprinting. Among those are RNases T1, U2, and A, which specifically hydrolyze ssRNAs at 30 –50 phosphodiester bonds after guanosines, adenosines, and pyrimidines, respectively [30]. Structure-specific nucleases cleaving all nucleotides encompass the dsRNA-specific RNase V1 [31] and the ssRNA-specific RNase I and T2 and nucleases S1 and P1 [30]. However, many of those RNases are not available anymore. In this section, we focus on the application of RNase A from bovine pancreas [32] and RNase I from E. coli [33] for RNA structure analyses and investigations on RNA–protein interactions identified via EMSA. Two strategies for the analysis of end-labeled RNA fragments obtained from RNase digestion assays are presented,

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namely analysis via denaturing PAGE using Hoefer SE 600 and analysis via capillary gel electrophoresis.

2

Materials

2.1 Rapid Amplification of cDNA Ends (RACE)

2.1.1 Total RNA Extraction

Until PCR amplification of cDNA (PCR 1), all materials and solutions used in this section have to be RNase-free. RNase-free filter tips, reaction tubes, and further plastic ware are commercially available. Glassware can be prepared by incubation at 180–200  C for at least 4 h (see Note 1). Purchased kits and reagents for RNA applications are nuclease-free. RNase-free water and several solutions can be produced by treatment with diethyl pyrocarbonate (DEPC) (see Note 2). For DEPC treatment, the solution is stirred with 0.1% DEPC overnight, followed by autoclaving at 120  C for 1 h. Solutions including substances that contain primary or secondary amines (e.g., Tris-based buffers) have to be purchased or prepared by solving the solid components in DEPC-treated water. All purchased solutions, reagents, and kits are stored according to the manufacturer’s instructions. If not stated otherwise, self-prepared solutions and materials are stored at room temperature. 1. Desired T. reesei strain: Store plates at 4  C and glycerol stocks at 80  C. 2. Malt extract medium: 3% malt extract, 0.1% peptone dissolved in tap water. Alternatively, any other medium for cultivation. 3. Miracloth. 4. Funnel. 5. Tap water. 6. Whatman paper. 7. Liquid nitrogen. 8. Screw cap reaction tubes: 2 mL. 9. peqGOLD TriFast (PEQLAB Biotechnologie, Erlangen, Germany). 10. Glass beads: 0.01–0.1 mm, 1 mm, and 5 mm in diameter. 11. Teflon-Homogenisator: BIO 101/Savant FastPrep 120 (MP Biomedicals, Santa Ana, California, USA). 12. Chloroform/isoamylalcohol (24:1). 13. Isopropanol. 14. 70% ethanol. 15. RNase-free water. 16. Spectrophotometer to measure nucleic acid concentration, e.g., NanoDrop (Thermo Fisher Scientific, Waltham, MA, USA).

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1. RNase-free water. 2. DNase I and 10 DNase I Reaction Buffer: store at 20  C. 3. 50 mM EDTA: Supplied with DNase I or prepared by a 1:10 dilution of 0.5 M EDTA with RNase-free water. Store at 20  C. 4. RNase A: 10 mg/mL stock, diluted with deionized water to a final concentration of 100 μg/mL. Store at 20  C. 5. PCR Purification Kit: For example, GeneJET PCR Purification Kit (Thermo Fisher Scientific, Waltham, MA, USA). Contains purification columns, Binding Buffer, Wash Buffer, and Elution Buffer. Store according to the manufacturer’s instructions. 6. 50 /30 RACE Kit, second generation (Roche, Basel, Switzerland) (see Note 3): Contains 10 Reaction Buffer, 2 mM dATP, 10 mM dNTPs, 5 cDNA Synthesis Buffer, 37.5 μM Oligo dT-Anchor Primer, 12.5 μM PCR-Anchor Primer, Terminale Transferase, Reverse Transcriptase. Store at 20  C. 7. Polymerase and supplied reaction buffer: Store at 20  C. 8. Deoxynucleotide mixture (dNTPs): Mixture of dATP, dCTP, dGTP, dTTP, 10 mM each. Store at 20  C. 9. PCR anchor primer (if applicable): Purchased synthetic ssDNA oligonucleotide, sequence 50 -GACCACGCGTATCGATG TCGAC-30 . Resuspend in deionized water to a stock concentration of 100 μM and dilute 1:8 to a final concentration of 12.5 μM. Store at 20  C. 10. Gene-specific primer 1 (GSP1r): Synthetic 20- to 30-nt-long ssDNA oligonucleotide specific for the transcript of interest, in reverse orientation (see Note 4). Resuspend in deionized water to a stock concentration of 100 μM and dilute 1:16 to a final concentration of 6.25 μM for application in PCR. Store at 20  C. 11. Gene-specific primer 2 (GSP2r): Synthetic 20- to 30-nt-long ssDNA oligonucleotide specific for the transcript of interest, in reverse orientation, nested relative to GSP1r (see Note 4). Resuspend in deionized water to a stock concentration of 100 μM and dilute 1:16 to a final concentration of 6.25 μM for application in PCR. Store at 20  C. 12. Gene-specific primer 3 (GSP3r): synthetic 20- to 30-nt-long ssDNA oligonucleotide specific for the transcript of interest, in reverse orientation, nested relative to GSP2r (see Note 4). Resuspend in deionized water to a stock concentration of 100 μM and dilute 1:16 to a final concentration of 6.25 μM for application in PCR. Store at 20  C. 13. Gene-specific primer 3* (GSP3r*): see GSP2f (optional).

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14. 1.5% agarose gel: 1.5% agarose dissolved in 1 TAE Buffer. Melt the agarose in 1 TAE by heating in a microwave, cool to room temperature, add ethidium bromide, and cast the gel. 2.1.3 30 RACE

1. μMACS™ Streptavidin Kit (Miltenyi Biotec, Bergisch Gladbach, Germany): Contains μ Columns, Equilibration buffer for nucleic acids, and μMACS Streptavidine MicroBeads. Store according to the manufacturer’s instructions. 2. μMACS Separator (Miltenyi Biotec, Bergisch Gladbach, Germany). 3. Two to three thermomixers (for reaction tubes). 4. Biotinylated probe: 30-nt-long synthetic DNA oligonucleotide specific for the transcript of interest (sequence complementary to the RNA sequence), 50 biotinylated, HPLC-purified. Dissolve the oligonucleotide in 1 TE buffer to a final concentration of 0.5 μg/μL. Store at 20  C after resuspension. Estimate the annealing temperature of the biotinylated probe from the melting temperature. Calculate the melting temperature as follows TM ¼ 67 + 16.6 log10 ([Na+]/(1.0 + 0.7 [Na+])) + 0.8 (%(GC))  500/n where [Na+] is the concentration of Na+ ions in mol/L, (%(GC)) is the percentage of GC in the duplex, and n is the length of the duplex. Use an annealing temperature of 10  C below the calculated melting temperature for a probe with one biotin. 5. 10 TEN buffer: 100 mM Tris–HCl, 10 mM EDTA, 1 M NaCl, pH 8.0. Dissolve 1.21 g of Tris, 0.29 g of EDTA, and 5.844 g of NaCl in RNase-free water. Adjust pH to 8.0 with HCl and fill up to a total volume of 100 mL with RNase-free water. Alternatively, prepare the buffer from DEPC-treated 0.5 M EDTA and 4 M NaCl stocks and Tris dissolved in RNase-free water. 6. 1 TEN buffer: 10 mM Tris–HCl, 1 mM EDTA, 100 mM NaCl, pH 8.0. Prepare a 1:10 dilution of 10 TEN buffer with RNase-free water. 7. 1 TE buffer: 10 mM Tris–HCl, 1 mM EDTA, pH 8.0. Dissolve 0.121 g of Tris and 0.029 g of EDTA in RNase-free water. Adjust pH to 8.0 with HCl and fill up to a total volume of 100 mL with RNase-free water. Alternatively, prepare the buffer from DEPC-treated 0.5 M EDTA and 4 M NaCl stocks and Tris dissolved in RNase-free water. 8. RNase-free water. 9. DNase I and 10 DNase I Reaction Buffer: Store at 20  C. 10. 50 mM EDTA: Supplied with DNase I or prepared by 1:10 dilution of 0.5 M EDTA with RNase-free water. Store at 20  C.

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11. Gene JET RNA Clean-up and Concentration Micro Kit (Thermo Fisher Scientific, Waltham, MA, USA): Contains Gene JET RNA Purification Micro Columns, Nuclease-free water, Binding Buffer, Wash Buffer 1, Wash Buffer 2. Store according to the manufacturer’s instructions. 12. 96% ethanol. 13. Ribonuclease Inhibitor (optional): Store at 20  C. 14. 50 /30 RACE Kit, second generation (Roche, Basel, Switzerland) (see Note 3): Contains 10 Reaction Buffer, 2 mM dATP, 10 mM dNTPs, 5 cDNA Synthesis Buffer, 37.5 μM Oligo dT-Anchor Primer, 12.5 μM PCR-Anchor Primer, Terminale Transferase, Reverse Transcriptase. Store at 20  C. 15. Polymerase and supplied reaction buffer: Store at 20  C. 16. Deoxynucleotide mixture (dNTPs): Mixture of dATP, dCTP, dGTP, dTTP, 10 mM each. Store at 20  C. 17. PCR Anchor Primer (if applicable): Purchased synthetic ssDNA oligonucleotide, sequence 50 -GACCACGCGTATC GATGTCGAC-30 . Resuspend in deionized water to a stock concentration of 100 μM and dilute 1:8 to a final concentration of 12.5 μM. Store at 20  C. 18. Gene-specific primer 1 (GSP1f): Synthetic 20- to 30-nt-long ssDNA oligonucleotide specific for the transcript of interest, in forward orientation (see Note 5). Resuspend in deionized water to a stock concentration of 100 μM and dilute 1:16 to a final concentration of 6.25 μM for application in PCR. Store at 20  C. 19. Gene-specific primer 2 (GSP2f): Synthetic 20- to 30-nt-long ssDNA oligonucleotide specific for the transcript of interest, in forward orientation, nested relative to GSP1f (see Note 5). Resuspend in deionized water to a stock concentration of 100 μM and dilute 1:16 to a final concentration of 6.25 μM for application in PCR. Store at 20  C. 20. Gene-specific primer 2* (GSP2f*): see GSP2f (optional). 21. 1.5% agarose gel: 1.5% agarose dissolved in 1 TAE Buffer. Melt the agarose in 1 TAE by heating in a microwave, cool to room temperature, add ethidium bromide, and cast the gel. 2.1.4 Preparation and Analysis of RACE Libraries

1. PCR reagents and primers used in Subheadings 2.1.2 and 2.1.3. 2. PCR purification or gel extraction kit: Store according to the manufacturer’s instructions. 3. Blunt-end digested vector: E.g., pJET1.2/blunt (Thermo Fisher Scientific, Waltham, MA, USA). Store at 20  C.

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4. Blunting enzyme and reaction buffer (if applicable): Included in CloneJET PCR Cloning Kit (Thermo Fisher Scientific, Waltham, MA, USA). Store at 20  C. 5. DNA ligase and reaction buffer: Included in CloneJET PCR Cloning Kit (Thermo Fisher Scientific, Waltham, MA, USA). Store at 20  C. 6. Competent E. coli cells: Store at 80  C. 7. LB: Dissolve 5 g of NaCl, 5 g of yeast extract, and 10 g of peptone in deionized water. Adjust pH to 7.0. Fill up to 1 L with deionized water. For the preparation of LB agar plates, add 1.5% agar before autoclaving. 8. Antibiotics for cloning: Depending on the resistance gene. Store according to empirical knowledge. 9. Plasmid preparation kit: Store according to the manufacturer’s instructions. 2.2 RNA In Vitro Synthesis and Labeling

2.2.1 Template Preparation for RNA In Vitro Synthesis

All materials and solutions used in this section (except for those used for the template preparation for RNA in vitro synthesis) have to be RNase-free. Electrophoresis equipment and tanks might be washed with 0.5% SDS, thoroughly rinsed with RNase-free water, rinsed with ethanol, and allowed to dry; however, I never observed RNA degradation without this prior treatment. For further information on the preparation and acquisition of RNase-free material and solutions, refer to Subheading 2.1. If not stated otherwise, solutions and materials are stored at room temperature. 1. Chromosomal DNA of T. reesei: Store at 4  C. 2. Proof reading polymerase and standard PCR reagents: Store at 20  C. 3. Primers containing the T7 RNA Polymerase promoter and appropriate restriction sites: Store at 20  C after resuspension. 4. PCR purification or gel extraction kit: Store according to the manufacturer’s instructions. 5. Vector lacking the T7 RNA Polymerase promoter (e.g., pUC18): Store at 20  C. 6. DNA ligase and standard ligation reagents: Store at 20  C. 7. Competent E. coli cells: Store at 80  C. 8. LB: Dissolve 5 g of NaCl, 5 g of yeast extract, and 10 g of peptone in deionized water. Adjust pH to 7.0. Fill up to 1 L with deionized water. For the preparation of LB agar plates, add 1.5% agar before autoclaving. 9. Antibiotics for cloning: Depending on the resistance gene. Store according to empirical knowledge.

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10. Plasmid preparation kit: Store according to the manufacturer’s instructions. 11. Restriction enzymes and corresponding buffers: Store according to the manufacturer’s instructions. 12. Phenol/chloroform mixture (1:1): Prepare freshly. 13. Chloroform. 14. 3 M sodium acetate, pH 5.2. 15. 96% ethanol. 16. 70% ethanol. 17. RNase-free water. 18. NanoDrop or similar instrumentation for nucleic acid quantification. 2.2.2 RNA In Vitro Synthesis

1. T7 High Yield RNA Synthesis Kit (New England Biolabs, Ipswich, MA, USA): Contains 100 mM ATP, CTP, GTP, and UTP, 10 Reaction Buffer and T7 RNA Polymerase Mix. Store at 20  C. 2. DNase I and 10 DNase I Reaction Buffer: Store at 20  C. 3. 50 mM EDTA: Supplied with DNase I or prepared by 1:10 dilution of 0.5 M EDTA with RNase-free water. Store at 20  C. 4. Phenol/chloroform mixture (1:1): Prepare freshly. 5. Chloroform. 6. 3 M sodium acetate, pH 5.2. 7. 96% ethanol. 8. 70% ethanol, ice-cold. 9. EMSA Buffer: 10 mM tricine, 50 mM NaCl, pH 7.4. Weigh 1.7917 g of tricine and 2.922 g of NaCl in an RNase-free bottle and dissolve in 900 mL of RNase-free water. Adjust pH to 7.4 with NaOH. Fill up to 1 L with RNase-free water. 10. NanoDrop or similar instrumentation for nucleic acid quantification.

2.2.3 Quality Analysis

1. Mini-PROTEAN Tetra Cell System (Bio-Rad Laboratories, Hercules, CA, USA). 2. 50 mL beaker and stir bar: Bake at 180  C overnight. 3. Urea. 4. 30% acrylamide/bisacrylamide solution (19:1): A purchased solution is recommended. Alternatively, the acrylamide and bisacrylamide can be weighed in a hood (wear a respiratory mask!), dissolved in RNase-free water and filtered with a

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0.45 μm filter unit. Store at 4  C in the absence of light (bottle covered with aluminum foil). 5. 10 TBE: 0.89 M Tris, 0.89 M boric acid, 0.02 M EDTA, pH 8.0. For preparation of 1 L 10 TBE weigh 108 g of Tris base, 7.4 g of EDTA, and 60 g of boric acid. Dissolve in 800 mL of RNase-free water. Adjust pH 8.0 with boric acid (see Note 6). Fill up to 1 L with RNase-free water. 6. 0.5 TBE: 1:20 dilution of the 10 TBE stock solution. 7. RNase-free water. 8. Ammonium persulfate (APS): 10% (w/v) APS dissolved in RNase-free water, store at 4  C (see Note 7). 9. N,N,N,N0 -Tetramethyl-ethylenediamine (TEMED): Store at 4  C. 10. 1.5 RNA loading dye: 95% formamide, 0.025% bromophenol blue (BPB), 0.025% xylene cyanol FF (XC), 5 mM EDTA. Prepare a 0.25 M EDTA solution, pH 8.0 (see Note 8) and treat with DEPC as described in Subheading 2.1. Prepare stock solutions of BPB and XC by dissolving 2.5% (w/v) in RNasefree water. Mix 9.5 mL of formamide, 200 μL of 0.25 M EDTA, 100 μL of 2.5% BPB, and 100 μL of 2.5% XC. Store in aliquots at 20  C. 11. RNA marker (if required): Store at 20  C. 12. Pasteur pipette. 13. Ethidium bromide solution: 1 μg/mL in 0.5 TBE. Add 50 μL of a 1 mg/mL ethidium bromide stock solution to 50 mL of 0.5 TBE. Store at 4  C in the absence of light (bottle covered with aluminum foil). The solution can be reused. 14. Small tray (gel size): Ethidium bromide contaminated. 15. Aluminum foil. 16. Gel imaging system: UV transilluminator. 2.2.4 RNA Labeling and Purification

1. T4 RNA Ligase: 10 U/μL stock. Store at 20  C. 2. pCp-CY5: 1 mM Cytidine-50 -phosphate-30 -(6-aminohexyl) phosphate, labeled with Cy5 (Jena Bioscience, Jena, Germany). Dilute 1:5 with RNase-free water to get 0.2 mM pCp-CY5. Store at 20  C. 3. Sephadex G-25 Quick Spin Columns for radiolabeled RNA purification (Roche, Basel, Switzerland): Store according to the manufacturer’s instructions. 4. Swinging bucket rotor and appropriate centrifuge.

RNA Characterization

2.3 RNA Electrophoretic Mobility Shift Assay (RNA-EMSA)

2.3.1 Preparation for the RNA-EMSA

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All materials and solutions used in this section (except for those used for protein purification and analysis) have to be RNase-free. Electrophoresis equipment and tanks might be washed with 0.5% SDS, thoroughly rinsed with RNase-free water, rinsed with ethanol, and allowed to dry; however, I never observed RNA degradation without this prior treatment. For further information on the preparation and acquisition of RNase-free material and solutions, refer to Subheading 2.1. If not stated otherwise, solutions and materials are stored at room temperature. 1. Materials for RNA in vitro synthesis: Refer to Subheading 2.2. 2. Materials for protein expression and purification: Store according to empirical knowledge. 3. Materials for classical SDS-PAGE: Store according to empirical knowledge. 4. PD-10 column (GE Healthcare, Uppsala, Sweden). 5. Sodium azide: 0.02% in deionized water. 6. Bradford reagent: Prepare a 1:5 dilution with deionized water. Store at 4  C. 7. BSA standards: 0, 10, 30, 50, 75, 100, 150, 300, 500, and 700 ng/μL. Dissolve 1 mg of BSA in 2 mL of deionized water to prepare a stock solution of 0.5 mg/mL BSA. Make a dilution series to get to the final concentrations. Store at 20  C. 8. Cuvettes and spectrophotometer. 9. Materials for RNA labeling (for RNA–RNA or RNA–DNA interaction studies only): Refer to Subheading 2.2. 10. 50 FAM-labeled forward primer and unlabeled reverse primer (for RNA–DNA interaction studies only): Purchased ssDNA oligonucleotides. After resuspension, store at 20  C. 11. Standard PCR reagents (for RNA–DNA interaction studies only): Store at 20  C. 12. PCR purification kit (for RNA–DNA interaction studies only): Store according to the manufacturer’s instructions. 13. Mini-PROTEAN Tetra Cell System (Bio-Rad Laboratories, Hercules, CA, USA). 14. EMSA Buffer: 10 mM tricine, 50 mM NaCl, pH 7.4. Weigh 1.7917 g of tricine and 2.922 g of NaCl in an RNase-free bottle and dissolve in 900 mL of RNase-free water. Adjust pH to 7.4 with NaOH. Fill up to 1 L with RNase-free water. 15. 30% acrylamide/bisacrylamide solution (30:0.36): Weigh 0.036 g of bisacrylamide in a hood (wear a respiratory mask!) and dissolve it in 100 mL of a purchased 30% acrylamide solution. Alternatively dissolve both acrylamide and bisacrylamide in RNase-free water and filter with a 0.45 μm filter unit.

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Store at 4  C in the absence of light (bottle covered with aluminum foil). 16. 10 TBE: 0.89 M Tris, 0.89 M boric acid, 0.02 M EDTA, pH 8.0. For preparation of 1 L 10 TBE weigh 108 g of Tris base, 7.4 g of EDTA, and 60 g of boric acid. Dissolve in 800 mL of RNase-free water. Adjust pH 8.0 with boric acid (see Note 6). Fill up to 1 L with RNase-free water. 17. 0.5 TBE: 1:20 dilution of the 10 TBE stock solution. One-time use for EMSA applications. 18. RNase-free water. 19. Ammonium persulfate (APS): 10% (w/v) APS dissolved in RNase-free water, store at 4  C (see Note 7). 20. N,N,N,N0 -Tetramethyl-ethylenediamine (TEMED): Store at 4  C. 21. Freezer ice block. 2.3.2 RNA-EMSA

1. Mini-PROTEAN Tetra Cell electrophoresis module (Bio-Rad Laboratories, Hercules, CA, USA). 2. Ice box (large, flat) and ice. 3. EMSA buffer and materials prepared in Subheading 2.3.1 (i.e., RNA, protein, gel, freezer ice block, and 0.5 TBE). 4. 50% glycerol: Mix glycerol 1:1 with RNase-free water. 5. Pasteur pipette. 6. Ethidium bromide solution: 1 μg/mL in 0.5 TBE. Add 50 μL of a 1 mg/mL ethidium bromide stock solution to 50 mL of 0.5 TBE. Store at 4  C in the absence of light (bottle covered with aluminum foil). The solution can be reused. 7. Small tray (gel size): Ethidium bromide contaminated. 8. Aluminum foil. 9. Gel imaging system: UV transilluminator or imaging system suitable for the detection of fluorescence signals for RNA– DNA interaction studies (i.e., Gel Doc, ChemiDoc).

2.4 RNA In Vitro Footprinting

All materials and solutions used in this section (except for those used for protein purification and analysis) have to be RNase-free. Electrophoresis equipment and tanks might be washed with 0.5% SDS, thoroughly rinsed with RNase-free water, rinsed with ethanol and allowed to dry; however, I never observed RNA degradation without this prior treatment. For further information on the preparation and acquisition of RNase-free material and solutions, refer to Subheading 2.1. If not stated otherwise, solutions and materials are stored at room temperature.

RNA Characterization 2.4.1 Preparation for RNA In Vitro Footprinting

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1. Materials for RNA in vitro synthesis and labeling: Refer to Subheading 2.2. 2. Materials for protein expression, purification, analysis, and quantification: Refer to Subheading 2.3.1

2.4.2 RNase Assay

1. RNA: In vitro synthesized RNA, CY5 or CY3 labeled. Store at 80  C. 2. Protein: Heterologously expressed protein, purified, and exchanged to EMSA Buffer. Store at 4  C. 3. EMSA Buffer: 10 mM tricine, 50 mM NaCl, pH 7.4. Weigh 1.7917 g of tricine and 2.922 g of NaCl in an RNase-free bottle and dissolve in 900 mL of RNase-free water. Adjust pH to 7.4 with NaOH. Fill up to 1 L with RNase-free water. 4. RNase I: 100 U/μL stock, diluted 1:20 with RNase-free water to a final concentration of 5 U/μL. Store at 20  C. 5. 1% SDS (for RNase I assay only): Dissolve 1% SDS in deionized water and treat with DEPC. 6. RNase A: 10 mg/mL stock, diluted with deionized water to a final concentration of 100 μg/mL. Store at 20  C. 7. Ribonuclease Inhibitor (for RNase A assay only): 20 U/μL stock, purchased. Store at 20  C.

2.4.3 Analysis of In Vitro Footprinting Samples Via Denaturing PAGE Using Hoefer SE 600

1. Hoefer SE 600 electrophoresis and casting module (Hoefer, Inc., Holliston, MA, USA). 2. 250 mL beaker and stir bar: Bake at 180  C overnight. 3. Urea. 4. 30% acrylamide/bisacrylamide solution (29:1): A purchased solution is recommended. Alternatively, the acrylamide and bisacrylamide can be weighed in a hood (wear a respiratory mask!), dissolved in RNase-free water, and filtered with a 0.45 μm filter unit. Store at 4  C in the absence of light (bottle covered with aluminum foil). 5. 10 TBE: 0.89 M Tris, 0.89 M boric acid, 0.02 M EDTA, pH 8.0. Weigh 108 g of Tris base, 7.4 g of EDTA, and 60 g of boric acid. Dissolve in 800 mL of RNase-free water. Adjust pH 8.0 with boric acid (see Note 6). Fill up to 1 L with RNasefree water. 6. 1 TBE: 1:10 dilution of the 10 TBE stock solution. 7. RNase-free water. 8. Ammonium persulfate (APS): 10% (w/v) APS dissolved in RNase-free water, store at 4  C (see Note 7). 9. N,N,N,N0 -Tetramethyl-ethylenediamine (TEMED): Store at 4  C.

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10. 50% glycerol: Mix glycerol 1:1 with RNase-free water. 11. Autoclave. 12. RNA marker: Store at 20  C. 13. Pasteur pipette or syringe. 14. Ethidium bromide solution: 1 μg/mL in 0.5 TBE. Add 50 μL of a 1 mg/mL ethidium bromide stock solution to 50 mL of 0.5 TBE. Store at 4  C in the absence of light (bottle covered with aluminum foil). The solution can be reused. 15. Tray (gel size): Ethidium bromide contaminated. 16. Aluminum foil. 17. Gel imaging system: UV transilluminator and imaging system suitable for the detection of fluorescence signals (i.e., Gel Doc, ChemiDoc). 2.4.4 Analysis of In Vitro Footprinting Samples Via Capillary Gel Electrophoresis

1. 2.5 M NaCl: Dissolve 14.61 g of NaCl in deionized water and fill up to 100 mL. Treat with DEPC. 2. 96% ethanol. 3. 70% ethanol. 4. RNase-free water or buffer for analysis via capillary gel electrophoresis. 5. Genetic Analyzer or sequencer, appropriate analysis kit, and an RNA (or DNA) marker (see Note 9): Store according to the manufacturer’s instructions. 6. Peak Scanner™ Software (Thermo Fisher Scientific, Waltham, USA) or equivalent peak analysis software. 7. Command-line-based program ivFAST [37] or equivalent analysis tool.

3

Methods

3.1 Rapid Amplification of cDNA Ends (RACE) 3.1.1 Total RNA Extraction

1. Cultivate the T. reesei strain of interest in the desired medium (e.g., in 50 mL of malt extract medium at 30  C and 180 rpm for 24–48 h) to obtain biomass for RNA extraction. Harvest the fungal mycelium by passing the culture through a miracloth filter placed in a funnel. Rinse with tap water and dry the collected biomass in a piece of Whatman paper. Shock-freeze the mycelium in liquid nitrogen. 2. Put 0.37 g of small glassbeads (0.01–0.1 mm in diameter), 0.25 g of medium glass beads (1 mm in diameter), and one large glass bead (5 mm in diameter) in a 2 mL screw cap reaction tube (see Note 10) and add 1 mL of peqGOLD TriFast.

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3. Add 0.01–0.02 g of frozen mycelium from the cultivation in step 1. 4. Disrupt the mycelium using the Teflon-Homogenisator at level 6 for 30 s. Incubate at room temperature for 5 min. 5. Add 200 μL of chloroform/isoamylalcohol (24:1) and mix well by rigorous shaking in a horizontal position (see Note 11). Incubate at room temperature for 10 min. Centrifuge for 5 min at 12,000  g and 4  C. From now on every used reaction tube and tip must be RNase-free! 6. Transfer 500 μL of the aqueous top phase to a new 1.5 mL reaction tube. Add an equal volume of chloroform/isoamylalcohol (24:1) and mix well by rigorous shaking in a horizontal position. Centrifuge for 5 min at 12,000  g and 4  C. 7. Transfer 350 μL of the aqueous top phase to a new 1.5 mL reaction tube. Add 0.7 volumes (245 μL) of isopropanol. Incubate at room temperature for 10–15 min. 8. Pellet the RNA by centrifugation for 15 min at 22,000  g and 4  C. Carefully remove the supernatant. 9. Wash the pellet with 900 μL of 70% ethanol and centrifuge for 5 min at 22,000  g and 4  C. Carefully remove the supernatant (see Note 12). 10. Dry the pellet at 50  C for 10 min. 11. Resuspend the pellet in 30 μL of RNase-free water by incubation at 50  C for 10 min (shaking). 12. Quantify the RNA using a suitable spectrophotometer. 3.1.2 50 RACE

If possible, proceed until finalization of the step “PCR Amplification of cDNA (PCR1)” in 1 day as RNA and cDNA are much more fragile than dsDNA. Keep RACE reactions and all components on ice during working. Store RNAs and cDNAs at 80  C and PCR at 20  C. PCR 1 and PCR 2 are performed in a PCR instrument, whereas DNase I digest, reverse transcription, RNase A digest, and poly(A)-tailing are performed in a PCR instrument or thermomixer. The RACE protocol is optimized for rare, highly structured transcripts (see Note 13).

DNase I Digest

1. Mix 1 μg of total RNA extract (from Subheading 3.1.1), 1 μL of 10 DNase I Reaction Buffer, 1 μL of DNase I and RNasefree water in a total reaction volume of 10 μL. 2. Incubate at 37  C for 30 min. 3. Stop the reaction by the addition of 1 μL of 50 mM EDTA and incubation at 65  C for 10 min.

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Reverse Transcription

1. Mix 11 μL of DNase I-digested RNA, 4 μL of cDNA Synthesis Buffer, 2 μL of 10 mM dNTPs, 2 μL of GSP1r, and 1 μL of Reverse Transcriptase (20 μL total reaction volume). 2. Incubate at 55  C for 60 min. 3. Stop the reaction by incubation at 85  C for 5 min.

RNase A Digest

1. Add 200 ng of RNase A (2 μL of a 100 μg/mL stock) to obtain a final concentration of 10 μg/mL. 2. Incubate at 37  C for 30 min.

Purification of the cDNA

As an example, DNA purification using the GeneJET PCR Purification Kit from Thermo Fisher Scientific is presented. The purification protocol is adapted according to the instructions provided in the manual of the RACE kit. Other PCR clean-up kits might be used, yet purification should be performed according to the modified purification protocol presented in this section. 1. Add Binding Buffer to the RNase digested cDNA according to the instructions of the kit protocol (equal volumes for GeneJET PCR Purification Kit) and mix by pipetting. 2. Transfer the total reaction mixture to a purification column preassembled with a collection tube. Centrifuge for 30 s at 8000  g. Discard the flow-through. 3. Add 500 μL of Wash Buffer and centrifuge for 30 s at 8000  g. Discard the flow-through. 4. Add 200 μL of Wash Buffer and centrifuge for 30 s at 8000  g. Discard the flow-through. 5. Centrifuge the empty column for 2 min at top speed to completely remove the Wash Buffer. 6. Place the column in a new 1.5 mL reaction tube. Add 40 μL of Elution Buffer to the column and incubate for 1 min. Centrifuge for 1 min at 8000  g to elute the cDNA.

Poly(A)-Tailing of cDNA

1. Mix 19 μL of purified cDNA, 2.5 μL of 10 Reaction Buffer and 2.5 mL of 2 mM dATP. 2. Denature at 95  C for 3 min. Cool on ice. 3. Add 1 μL of Terminale Transferase. 4. Incubate at 37  C for 20–30 min. 5. Stop the reaction by incubation at 70  C for 10 min.

PCR Amplification of cDNA (PCR 1)

Initial amplification is based on binding of the oligo(dT) contained in the Oligo(dT)-Anchor Primer to the poly(A)-tail of the cDNA, which allows a maximum annealing temperature of 55  C. For further rounds of amplification, the total sequence of the Oligo (dT)-Anchor Primer can hybridize, thus an annealing temperature

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of up to 65  C can be applied. Consequently, for the 50 RACE, the amplification of poly(A)-tailed cDNA is performed in two stages of repeating cycles. 1. Apply 1 μL of undiluted or 1:10-diluted cDNA as a template in a 25 μL PCR using the Oligo(dT)-Anchor Primer and the gene-specific primer GSP2r. For the preparation of a 50 μL master mix, combine 1 μL of the Oligo(dT)-Anchor Primer, 2 μL of GSP2r, 1 μL of 10 mM dNTPs, Reaction Buffer, polymerase, and deionized water in a PCR tube, distribute on two tubes and add the template. The exact composition of the PCR depends on the polymerase (see Note 14) and must be determined individually. 2. Run a PCR program. The optimal reaction conditions depend on the gene-specific primer and the polymerase. Use an annealing temperature of 55  C or below for the first 10 cycles, followed by an annealing temperature of 60  C to 65  C for 30 cycles. The elongation time depends on the polymerase and the estimated transcript length (see Note 15). An example of a PCR program for PCR 1 is given below: 95  C 3 min. 95  C 30 s/55  C 30 s/72  C 1 min (10 cycles). 95  C 30 s/60  C 30 s/72  C 1 min (30 cycles). 72  C 5 min. 3. Analyze 10 μL of PCR 1 on a 1.5% agarose gel. The first amplification step of the 50 RACE usually yields a weak smear signal rather than a discrete band (see Note 16). Nested PCR (PCR 2)

1. Apply 1 μL of a 1:50 dilution (see Note 17) of the two approaches from PCR 1 (resulting from application of undiluted or 1:10-diluted cDNA) as a template in a 25 μL PCR using the PCR Anchor Primer and the nested gene-specific primer GSP3r. For the preparation of a 50 μL master mix, combine 1 μL of the PCR Anchor Primer, 2 μL of GSP3r, 1 μL of 10 mM dNTPs, Reaction Buffer, polymerase, and deionized water in a PCR tube, distribute on two tubes, and add the template. The exact composition of the PCR depends on the polymerase (see Note 14) and must be determined individually. 2. Optional: To verify the specificity of the product generated in PCR 2, the same PCR as described in step 1 but using GSP3r* instead of GSP3r might be performed in parallel (see Note 18). 3. Run a PCR program. The optimal reaction conditions depend on the gene-specific primer and the polymerase. Use an annealing temperature from 60  C to 65  C. The elongation time

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depends on the polymerase and the estimated transcript length (see Note 15). 4. Analyze 10 μL of PCR 2 on a 1.5% agarose gel. Specific products should appear as one or more discrete bands (see Note 16). If a second nested PCR was performed, products from the PCR using GSP3r* should be shifted relative to the PCR using GSP3r. Sequence Analysis

Determine the 50 end of the transcript of interest by preparation and analysis of a RACE library as described in Subheading 3.1.4.

3.1.3 30 RACE

If possible, proceed until finalization of the step “PCR Amplification of cDNA (PCR1)” on 1 day as RNA and cDNA are much more fragile than dsDNA. Keep RACE reactions and all components on ice during working. Store RNAs and cDNAs at 80  C and PCR at 20  C. PCR 1 and PCR 2 are performed in a PCR instrument, whereas DNase I digest, poly(A)-tailing, and reverse transcription are performed in a PCR instrument or thermomixer. The RACE protocol is optimized for rare, highly structured transcripts (see Note 13).

Enrichment of Rare Transcripts of Interest

For poorly expressed RNAs such as lncRNAs, the transcript of interest may be enriched from 2 mg of total RNA (a pool of approximately 20–30 total RNA extracts prepared according to the protocol in Subheading 3.1.1, see Note 19) by hybridization with a biotinylated target-specific DNA probe, which can then be isolated using streptavidin-linked magnetic beads and a magnetic separator unit (e.g., μMACS Separator). Prepare all solutions in advance and pre-heat the thermomixers to the appropriate temperatures. Pre-heat an aliquot of RNase-free water to 80  C. Work quickly during the following procedure. 1. Mix 2 mg of total RNA extract (contained in a maximal volume of 1.3 mL) with 10 TEN buffer to obtain 1 TEN in your reaction. 2. Incubate at 75–85  C (see Note 20) for 5 min for denaturation. Then cool the reaction to the calculated annealing temperature of the target-specific biotinylated probe. 3. Add 1.6 μL of the biotinylated probe (see Note 21) and mix well. Incubate at the calculated annealing temperature for 15 min for hybridization. 4. Meanwhile, prepare the column supplied in the μMACS™ Streptavidin Kit. Unpack the column and place it in the μMACS Separator. Place a 2 mL reaction tube beyond the outlet of the column to collect the flow-through. Wash the column one time with 100 μL of Equilibration buffer for

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nucleic acid applications and twice with 1 TEN buffer. Remove the flow-through. 5. Add 160 μL of μMACS Streptavidine MicroBeads (vortexed!) to the hybridization reaction and mix well (see Note 22). Incubate at the calculated annealing temperature for 2 min. 6. Immediately transfer the hybridization reaction mixture to the column. Remove the flow-through. Do not interrupt magnetic attachment at any time until elution of your sample! 7. Wash the column five times with 200 μL of 1 TE buffer. 8. Elute the enriched RNA from the magnetic beads with 150 μL of RNase-free 80  C (pre-heated) water. Collect the flowthrough in a fresh 1.5 mL reaction tube. 9. The RNA concentration in the final eluate is usually too low for the detection via NanoDrop. Proceed to DNase I digest. DNase I Digest

1. Add 17.2 μL of 10 DNase I Reaction Buffer and 5 μL of DNase I to the 150 μL of RNA eluate. 2. Incubate at 37  C for 30 min. 3. Stop the reaction by the addition of 5 μL of 50 mM EDTA and incubate at 65  C for 10 min.

RNA Purification

Here, a protocol for the purification of DNase I digested RNA using the Gene JET RNA Cleanup and Concentration Micro Kit is presented. Alternatively, the RNA might be precipitated with isopropanol as described for the total RNA extraction in Subheading 3.1.1 and resuspended in 20 μL of RNase-free water. 1. Adjust the volume of the DNase I digest to 200 μL with RNase-free water and split the reaction to two reaction tubes (100 μL per tube). Add 500 μL of Binding Buffer to each reaction. Mix thoroughly by pipetting. 2. Add 600 μL of 96% ethanol per reaction and mix by pipetting. 3. Transfer the total reaction mixture to two Gene JET RNA Purification Micro Columns preassembled with collection tubes (600 μL per column). Centrifuge for 1 min at 14,000  g. Discard the flow-through. Repeat the step until the whole reaction mixture is loaded to the columns. 4. Add 700 μL of Wash Buffer 1 and centrifuge for 1 min at 14,000  g. Discard the flow-through. 5. Add 700 μL of Wash Buffer 2 and centrifuge for 1 min at 14,000  g. Discard the flow-through. 6. Repeat step 5. 7. Centrifuge the empty column for 1 min at 14,000  g to completely remove the Wash Buffer. 8. Add 10 μL of nuclease-free water to the column and centrifuge for 1 min at 14,000  g to elute the RNA. Pool the eluates from the 2 columns.

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Poly(A)-Tailing (for ncRNAs Lacking a Poly(A)-Tail Only)

Noncoding transcripts lacking a poly(A)-tail can be analyzed by poly(A)-tailing of the enriched RNA of interest prior to 30 RACE using a Terminale Transferase. Except for the optional Ribonuclease Inhibitor, all reagents are included in the 50 /30 RACE Kit, second generation from Roche. 1. Mix 19 μL of purified RNA, 2.5 μL of 10 Reaction Buffer and 2.5 mL of 2 mM dATP. 2. Denature at 95  C for 3 min. Cool on ice. 3. Add 1 μL of Terminale Transferase and 1 μL of Ribonuclease Inhibitor (optional). 4. Incubate at 37  C for 20 min. 5. Stop the reaction by incubation at 70  C for 10 min.

Reverse Transcription

1. Mix 12 μL of poly(A)-tailed RNA, 4 μL of cDNA Synthesis Buffer, 2 μL of 10 mM dNTPs, 1 μL of Oligo(dT)-Anchor Primer, and 1 μL of Reverse Transcriptase (20 μL total reaction volume). 2. Incubate at 55  C for 60 min. 3. Stop the reaction by incubation at 85  C for 5 min.

PCR Amplification of cDNA (PCR 1)

1. Apply 1 μL and 5 μL of cDNA as a template in a 25 μL PCR using the PCR Anchor Primer and the gene-specific primer GSP1f. For the preparation of a 50 μL master mix, combine 1 μL of the PCR Anchor Primer, 2 μL of GSP1f, 1 μL of 10 mM dNTPs, Reaction Buffer, polymerase, and deionized water in a PCR tube, distribute on two tubes, and add the template. The exact composition of the PCR depends on the polymerase (see Note 14) and must be determined individually. 2. Run a PCR program. The optimal reaction conditions depend on the gene-specific primer and the polymerase. Use an annealing temperature from 60  C to 65  C. The elongation time depends on the polymerase and the estimated transcript length (see Note 15). 3. Analyze 10 μL of PCR 1 on a 1.5% agarose gel. The first amplification step of the 30 RACE usually yields a continuous smear rather than a specific product (see Note 16).

Nested PCR (PCR 2)

1. Apply 1 μL of a 1:50 dilution (see Note 17) of the two approaches of PCR 1 (resulting from application of either 1 μL or 5 μL of cDNA) as a template in a 25 μL PCR using the PCR Anchor Primer and the nested gene-specific primer GSP2f. For the preparation of a 50 μL master mix, combine 1 μL of the PCR Anchor Primer, 2 μL of GSP2f, 1 μL of 10 mM dNTPs, Reaction Buffer, polymerase, and deionized water in a

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PCR tube, distribute on two tubes and add the template. The exact composition of the PCR depends on the polymerase (see Note 14) and must be determined individually. 2. Optional: To verify the specificity of the product generated in PCR 2, the same PCR as described in step 1 but using GSP2f* instead of GSP2f might be performed in parallel (see Note 18). 3. Run a PCR program. The optimal reaction conditions depend on the gene-specific primer and the polymerase. Use an annealing temperature from 60  C to 65  C. The elongation time depends on the polymerase and the estimated transcript length (see Note 15). 4. Analyze 10 μL of PCR 2 on a 1.5% agarose gel. Specific products should appear as one or more discrete bands (see Note 16). If a second nested PCR was performed, products from the PCR using GSP2f* should be shifted relative to the PCR using GSP2f. Sequence Analysis

Determine the 30 end of the transcript of interest by preparation and analysis of a RACE library as described in Subheading 3.1.4.

3.1.4 Preparation and Analysis of RACE Libraries

Commonly, PCR products resulting from RACE are not one defined product, but a mixture of fragments originating from slightly different versions of the target transcript and unspecific products. Therefore, a library must be prepared for the final analysis of 50 and 30 RACE products. Where applicable, a sequencing instrument might be used for the analysis of RACE libraries. In this case, a library has to be prepared depending on the sequencing instrument and according to the manufacturer’s instructions of the used kit. Alternatively, the PCR products can be cloned into a vector, transformed into Escherichia coli, and the plasmids of single colonies can be sent to a sequencing service for final analysis. This strategy is especially suitable for the analysis of ncRNAs with a single predominating end as it is relatively cheap and does not require complex library preparation or specific instrumentations. It also yields insights into the ratio of different transcript versions in complex libraries, yet it has to be considered that the analyzed fragments are only samples of the total library. 1. Preparation of specific RACE products for the final analysis: Prepare 2 50 μL of the PCR 2 yielding specific RACE products as described in Subheadings 3.1.2 and 3.1.3. 2. Purification: Extract the product bands of interest from a gel using a commercially available kit. Alternatively, the PCR products might be purified using a PCR purification kit, but usually extraction from a gel is more efficient (i.e., yields a higher number of specific sequences).

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3. Blunting (if applicable): Depending on the polymerase, blunt ends or sticky ends are obtained from PCR amplification. For polymerases producing short overhangs such as Taq polymerases, perform blunting of the PCR products prior to ligation according to the instructions supplied with the blunting enzyme. When the CloneJET PCR Cloning Kit is used, blunting and ligation can be performed in the same reaction buffer, and all reagents required for the two reaction steps are included in the kit. 4. Ligation: Ligate the PCR product into a blunt-end digested vector such as the commercially available pJET1.2/blunt. 5. Transformation in E. coli: Transform the ligation approach into competent E. coli cells and plate the cells on selection media (i.e., LB plates containing antibiotics). 6. Plasmid preparation: Grow a few transformants in LB containing antibiotics overnight and isolate the plasmids of using a commercially available plasmid preparation kit. 7. Sequencing and final analysis: Send the plasmids to a sequencing service for final analysis. Use primers located close to the target site of integration of the RACE fragments. Compare the obtained sequences to the known sequence of the RNA of interest. 3.2 RNA In Vitro Synthesis and Labeling

As a template for RNA in vitro synthesis, a plasmid containing the RNA encoding gene under the control of the T7 RNA Polymerase promoter is prepared (see Note 23).

3.2.1 Template Preparation for RNA In Vitro Synthesis Vector Construction and Cloning

1. Amplify the RNA encoding gene from chromosomal DNA of T. reesei with primers containing the T7 RNA Polymerase promoter (see Note 24) and appropriate restriction sites (see Note 25) via standard PCR techniques. 2. Purify the PCR product or extract it from a gel using a commercially available kit. 3. Ligate it into a vector lacking the T7 RNA Polymerase promoter (e.g., pUC18). 4. Transform the ligation approach into competent E. coli cells and plate the cells on selection media (i.e., LB plates containing antibiotics).

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5. Grow a few transformants in LB containing antibiotics overnight and isolate the final vector via plasmid prep using a commercially available kit. Linearization of the Vector

Phenol/Chloroform Extraction (See Note 26)

Linearize 20–200 μg of the vector in a 100 μL reaction by digestion with a restriction enzyme that generates blunt ends or 50 -overhangs (e.g., XbaI) and cuts immediately downstream of the cloned gene. The restriction site might be attached via the reverse primer used for PCR. Follow the manufacturer’s instructions for digestion and inactivation of the restriction enzyme. 1. Add an equal volume (100 μL) of phenol/chloroform (1:1). Mix by rigorous shaking for 15 s. Incubate at room temperature for 10 min. Centrifuge for 10 min at 12,000  g and 4  C. Transfer the aqueous top phase to a new 1.5 mL reaction tube. 2. Add an equal volume of chloroform and mix by rigorous shaking for 30 s. Centrifuge for 10 min at 12,000  g and 4  C. Transfer the aqueous top phase to a new 1.5 mL reaction tube. 3. Repeat extraction with chloroform and centrifugation (step 2). 4. Transfer the aqueous top phase to a new 1.5 mL reaction tube and add 1/10 volume of 3 M sodium acetate pH 5.2 and two volumes of 96% ethanol. Mix by inversion. Incubate at 20  C for at least 30 min. 5. Pellet the DNA by centrifugation for 15 min at top speed and 4  C. Carefully remove the supernatant. 6. Wash the pellet with 500 μL of 70% ethanol and centrifuge for 15 min at top speed and 4  C. Carefully remove the supernatant. 7. Dry the pellet at 50  C for 10 min. 8. Resuspend the pellet in 50 μL of RNase-free water by incubation at 50  C for 10 min (shaking). 9. Quantify the DNA via NanoDrop. You should end up with a final concentration of at least 125 ng/μL (preferentially 500–1000 ng/μL).

3.2.2 RNA In Vitro Synthesis Standard RNA In Vitro Synthesis Procedure DNase I Digest

1. Mix 2 μL of 100 mM ATP, CTP, GTP, UTP, and 10 Reaction Buffer (see Note 27). Add 1 μg of template DNA and fill up to 18 μL with RNase-free water. Add 2 μL of T7 RNA Polymerase Mix. Mix thoroughly and pulse-spin in a microfuge. 2. Incubate at 37  C for 2 h in a PCR instrument (see Note 28). 1. Add 68 μL of nuclease-free water, 10 μL of 10 DNase I Reaction Buffer, and 2 μL of DNase I to the reaction approach. 2. Incubate at 37  C for 15 min. 3. Stop the reaction by adding 5 μL of 50 mM EDTA and incubation at 65  C for 10 min.

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Phenol/Chloroform Extraction

1. Adjust the reaction volume to 180 μL by the addition of 80 μL of RNase-free water. Add 20 μL (1/10 volume) of 3 M sodium acetate, pH 5.2, and mix by resuspension. 2. Add 200 μL of phenol/chloroform (1:1). Mix by rigorous shaking for 15 s. Centrifuge for 5 min at 12,000  g and 4  C. Transfer the aqueous top phase to a new 1.5 mL reaction tube. 3. Add an equal volume of chloroform and mix by rigorous shaking for 30 s. Centrifuge for 5 min at 12,000  g and 4  C. Transfer the aqueous top phase to a new 1.5 mL reaction tube. 4. Repeat extraction with chloroform and centrifugation (step 3). 5. Transfer the aqueous top phase to a new 1.5 mL reaction tube and add two volumes of 96% ethanol. Mix by inversion. Incubate at 20  C for at least 30 min. 6. Pellet the DNA by centrifugation for 30 min at top speed and 4  C. Carefully remove the supernatant. 7. Wash the pellet with 500 μL of ice-cold 70% ethanol and centrifuge for 10 min at top speed and 4  C. Carefully remove the supernatant. 8. Dry the pellet at 50  C for 10 min. 9. Resuspend the pellet in 50 μL of EMSA buffer by incubation at 50  C for 10 min (shaking). 10. Quantify the DNA via NanoDrop. RNA in vitro synthesis usually yields final concentrations of 2000–5000 ng/μL.

3.2.3 Quality Analysis

The quality of the in vitro synthesized RNA can be analyzed by denaturing polyacrylamide gel electrophoresis (PAGE) (see Note 29). RNAs of 150–500 nt in length are separated on a 5% polyacrylamide (PAA) gel (19:1 acrylamide/bisacrylamide). For longer or shorter RNAs, the percentage of acrylamide has to be adjusted as specified in Table 1. In this case, also the running conditions might be altered to achieve an optimal resolution.

Preparation of Four Denaturing 5% PAA Gels (Cassette Size 7.3 cm  10.1 cm  1 mm)

1. Assemble the Mini-PROTEAN Tetra Cell casting apparatus. 2. Weigh in 12 g of urea in an RNase-free 50 mL beaker. 3. Add 4.167 mL of 30% acrylamide/bisacrylamide solution (19:1) and 2.5 mL of 10 TBE. Fill up with RNase-free water to approximately 23 mL (500

5

150–500

10

61–150

15

30–60

20

Save Report from the main menu. By default a report will be created using the name of the fastq file with _fastqc.zip appended to the end. 3.4.2 Read Trimming [8]

1. Install Trimmomatic using the link: http://www.usadellab. org/cms/?page¼trimmomatic (see Note 23). 2. Run Trimmomatic for the Paired End Mode: java -jar PE [-threads CWD (1) /blast/executables/blast+/LATEST ... done. ==> SIZE ncbi-blast-2.10.0+-x64-linux.tar.gz ... 233258021 ==> PASV ... done.

==> RETR ncbi-blast-2.10.0+-x64-linux.tar.gz ... done.

Length: 233258021 (222M) (unauthoritative) 100%[======================================>] 233,258,021 2.11MB/s in 4m 6s 2020-03-13 15:08:56 (925 KB/s) - ‘ncbi-blast-2.10.0+-x64-linux.tar.gz’ saved [233258021] [name of user account app]$ tar zxvf ncbi-blast-2.10.0+-x64-linux.tar.gz

# List all BLAST files

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[name of user account app]$ tar zxvf ncbi-blast-2.10.0+-x64-linux.tar.gz ncbi-blast-2.10.0+/ ncbi-blast-2.10.0+/ChangeLog ncbi-blast-2.10.0+/LICENSE ncbi-blast-2.10.0+/README ncbi-blast-2.10.0+/bin/ ncbi-blast-2.10.0+/bin/blast_formatter ncbi-blast-2.10.0+/bin/blastdb_aliastool ncbi-blast-2.10.0+/bin/blastdbcheck ncbi-blast-2.10.0+/bin/blastdbcmd ncbi-blast-2.10.0+/bin/blastn ncbi-blast-2.10.0+/bin/blastp ncbi-blast-2.10.0+/bin/blastx ncbi-blast-2.10.0+/bin/cleanup-blastdb-volumes.py ncbi-blast-2.10.0+/bin/convert2blastmask ncbi-blast-2.10.0+/bin/deltablast ncbi-blast-2.10.0+/bin/dustmasker ncbi-blast-2.10.0+/bin/get_species_taxids.sh ncbi-blast-2.10.0+/bin/legacy_blast.pl ncbi-blast-2.10.0+/bin/makeblastdb ncbi-blast-2.10.0+/bin/makembindex ncbi-blast-2.10.0+/bin/makeprofiledb ncbi-blast-2.10.0+/bin/psiblast ncbi-blast-2.10.0+/bin/rpsblast ncbi-blast-2.10.0+/bin/rpstblastn

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ncbi-blast-2.10.0+/bin/segmasker ncbi-blast-2.10.0+/bin/tblastn ncbi-blast-2.10.0+/bin/tblastx ncbi-blast-2.10.0+/bin/update_blastdb.pl ncbi-blast-2.10.0+/bin/windowmasker ncbi-blast-2.10.0+/doc/ ncbi-blast-2.10.0+/doc/README.txt ncbi-blast-2.10.0+/ncbi_package_info

3

Methods

3.1 Genome Sequence File Formats

The format of genome sequence files analyzed by TSETA is “fasta.” QM6a.genome.fa CBS1-1.genome.fa WTH109.genome.fa WTH111.genome.fa WTH115.genome.fa WTH119.genome.fa The order of each homologous chromosome in all six fasta genome sequence files must be the same.

3.2

TSETA Options

3.2.1 Tetrad Analysis

3.2.2 SNP Calling

TSETA was originally designed for genome-wide mapping of meiotic recombination products and RIP mutations. Accordingly, it requires the users to upload six genome sequences, two parental genomes in a hybrid zygote and the four F1 progeny generated from the same meiotic event. TSESA can be applied to determine SNPs (e.g., SNVs and Indels) of two or more query genomes to a reference genome.

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Running TSETA

3.3.1 Starting TSETA by Changing Directory to TSETA

# setup TSETA [name of user account TSETA]$ cd TSETA [name of user account TSETA]$ node setup.js path to blastn or blastn command (default blastn): (text) > ~/app/ncbi-blast-2.10.0+/bin/blastn ~/app/ncbi-blast-2.10.0+/bin/blastn -version blastn: 2.10.0+ Package: blast 2.10.0, build Dec 3 2019 18:03:18

path to mafft or mafft command (default mafft): (text) > ~/app/mafft-linux64/mafft.bat v7.450 (2019/Aug/23) ~/app/mafft-linux64/mafft.bat --version

save setting { blastn_bin: '~/app/ncbi-blast-2.10.0+/bin/blastn', mafft_bin: '~/app/mafft-linux64/mafft.bat' } 3.3.2 Running the “SNP Calling” Mode

On-screen instructions will be given to users to select the two reference genome sequence files [e.g., QM6a and CBS999.97 (MAT-1)]. # SNP mode [name of user account TSETA]$ node src/prepare.js Task name: (text)

All input parameters will be saved into DemoSNP.json All output files are kept at DemoSNP

Tetrad Analysis Using PacBio SMRT Technology

> DemoSNP mode (default tetrad): (tetrad, SNP) > SNP ref name: (text) > QM6a ref genome fasta file (default QM6a.genome.fa): (file path) > QM6a.genome.fa number of subject: (number) >1 subject 1 name: (text) > CBS1-1 subject 1 fasta file (default CBS1-1.genome.fa): (file path) > CBS1-1.genome.fa GenomeDataSet { name: 'DemoSNP', mode: 'SNP', ref: 'QM6a', parental_list: [ 'QM6a' ], parental: { QM6a: 'QM6a.genome.fa' }, progeny_list: [ 'CBS1-1' ], progeny: { 'CBS1-1': 'CBS1-1.genome.fa' } }

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output: DemoSNP/QM6a.length.txt output: DemoSNP/CBS1-1.length.txt output: QM6a 1 DemoSNP/tmp/fasta/QM6a_ChI_QM6a.fa output: QM6a 2 DemoSNP/tmp/fasta/QM6a_ChII_QM6a.fa output: QM6a 3 DemoSNP/tmp/fasta/QM6a_ChIII_QM6a.fa output: QM6a 4 DemoSNP/tmp/fasta/QM6a_ChIV_QM6a.fa output: QM6a 5 DemoSNP/tmp/fasta/QM6a_ChV_QM6a.fa output: QM6a 6 DemoSNP/tmp/fasta/QM6a_ChVI_QM6a.fa output: QM6a 7 DemoSNP/tmp/fasta/QM6a_ChVII_QM6a.fa output: CBS1-1 1 DemoSNP/tmp/fasta/CBS1-1_Ch1_CBS1-1_unitig_0RV_consensus.fa output: CBS1-1 2 DemoSNP/tmp/fasta/CBS1-1_Ch2_CBS1-1_unitig_1_consensus.fa output: CBS1-1 3 DemoSNP/tmp/fasta/CBS1-1_Ch3_CBS1-1_unitig_2RV_consensus.fa output: CBS1-1 4 DemoSNP/tmp/fasta/CBS1-1_Ch4_CBS1-1_unitig_3RV_consensus.fa output: CBS1-1 5 DemoSNP/tmp/fasta/CBS1-1_Ch5_CBS1-1_unitig_5_consensus.fa output: CBS1-1 6 DemoSNP/tmp/fasta/CBS1-1_Ch6_CBS1-1_unitig_6RV_consensus.fa output: CBS1-1 7 DemoSNP/tmp/fasta/CBS1-1_Ch7_CBS1-1_unitig_4RV_consensus.fa save: DemoSNP.json next step: detect GC content %, AT-rich blocks, centromere, telomeres command: node ./src/make_GC_AT_table.js -dataset DemoSNP.json -w 500 -m 6

(see Note 1) 3.3.3 Running the “Tetrad Analysis” Mode

On-screen instructions will be given to users to select the two parental genome sequence files [e.g., QM6a and CBS999.97 (MAT-1)] and the four representative F1 progeny genome sequences (e.g., WTH109, WTH111, WTH115, and WTH119).

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1. Step 1: Tetrad Analysis Mode [name of user account TSETA]$ node src/prepare.js Task name: (text) > DemoTetrad mode (default tetrad): (tetrad, SNP) > tetrad ref (parental 1) name: (text) > QM6a ref (parental 1) genome fasta file (default QM6a.genome.fa): (file path) > QM6a.genome.fa parental 2 name: (text) > CBS1-1 parental 2 genome fasta file (default CBS1-1.genome.fa): (file path) > CBS1-1.genome.fa progeny 1 name: (text) > WTH109 progeny 1 fasta file (default WTH109.genome.fa): (file path) > WTH109.genome.fa progeny 2 name: (text) > WTH111 progeny 2 fasta file (default WTH111.genome.fa): (file path) > WTH111.genome.fa progeny 3 name: (text) > WTH115

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progeny 3 fasta file (default WTH115.genome.fa): (file path) > WTH115.genome.fa progeny 4 name: (text) > WTH119 progeny 4 fasta file (default WTH119.genome.fa): (file path) > WTH119.genome.fa GenomeDataSet { name: 'DemoTetrad', mode: 'tetrad', ref: 'QM6a', parental_list: [ 'QM6a', 'CBS1-1' ], parental: { QM6a: 'QM6a.genome.fa', 'CBS1-1': 'CBS1-1.genome.fa' }, progeny_list: [ 'WTH109', 'WTH111', 'WTH115', 'WTH119' ], progeny: { WTH109: 'WTH109.genome.fa', WTH111: 'WTH111.genome.fa', WTH115: 'WTH115.genome.fa', WTH119: 'WTH119.genome.fa' } } output: DemoTetrad/QM6a.length.txt output: DemoTetrad/CBS1-1.length.txt output: DemoTetrad/WTH109.length.txt output: DemoTetrad/WTH111.length.txt

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output: DemoTetrad/WTH115.length.txt output: DemoTetrad/WTH119.length.txt output: QM6a 1 DemoTetrad/tmp/fasta/QM6a_ChI_QM6a.fa output: QM6a 2 DemoTetrad/tmp/fasta/QM6a_ChII_QM6a.fa output: QM6a 3 DemoTetrad/tmp/fasta/QM6a_ChIII_QM6a.fa output: QM6a 4 DemoTetrad/tmp/fasta/QM6a_ChIV_QM6a.fa output: QM6a 5 DemoTetrad/tmp/fasta/QM6a_ChV_QM6a.fa output: QM6a 6 DemoTetrad/tmp/fasta/QM6a_ChVI_QM6a.fa output: QM6a 7 DemoTetrad/tmp/fasta/QM6a_ChVII_QM6a.fa output: CBS1-1 1 DemoTetrad/tmp/fasta/CBS1-1_Ch1_CBS1-1_unitig_0RV_consensus.fa output: CBS1-1 2 DemoTetrad/tmp/fasta/CBS1-1_Ch2_CBS11_unitig_1_consensus.fa output: CBS1-1 3 DemoTetrad/tmp/fasta/CBS1-1_Ch3_CBS1-1_unitig_2RV_consensus.fa output: CBS1-1 4 DemoTetrad/tmp/fasta/CBS1-1_Ch4_CBS1-1_unitig_3RV_consensus.fa output: CBS1-1 5 DemoTetrad/tmp/fasta/CBS1-1_Ch5_CBS11_unitig_5_consensus.fa output: CBS1-1 6 DemoTetrad/tmp/fasta/CBS1-1_Ch6_CBS1-1_unitig_6RV_consensus.fa output: CBS1-1 7 DemoTetrad/tmp/fasta/CBS1-1_Ch7_CBS1-1_unitig_4RV_consensus.fa

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output: WTH109 1 DemoTetrad/tmp/fasta/WTH109_Ch1_hexadecad_008A.fa output: WTH109 2 DemoTetrad/tmp/fasta/WTH109_ch2_hexadecad_008A.fa output: WTH109 3 DemoTetrad/tmp/fasta/WTH109_ch3_hexadecad_008A.fa output: WTH109 4 DemoTetrad/tmp/fasta/WTH109_ch4_hexadecad_008A.fa output: WTH109 5 DemoTetrad/tmp/fasta/WTH109_ch5_hexadecad_008A.fa output: WTH109 6 DemoTetrad/tmp/fasta/WTH109_Ch6_hexadecad_008A.fa output: WTH109 7 DemoTetrad/tmp/fasta/WTH109_ch7_hexadecad_008A.fa output: WTH111 1 DemoTetrad/tmp/fasta/WTH111_ch1_hexadecad_008C.fa output: WTH111 2 DemoTetrad/tmp/fasta/WTH111_ch2_hexadecad_008C.fa output: WTH111 3 DemoTetrad/tmp/fasta/WTH111_ch3_hexadecad_008C.fa output: WTH111 4 DemoTetrad/tmp/fasta/WTH111_ch4_hexadecad_008C.fa output: WTH111 5 DemoTetrad/tmp/fasta/WTH111_ch5_hexadecad_008C.fa output: WTH111 6 DemoTetrad/tmp/fasta/WTH111_ch6_hexadecad_008C.fa output: WTH111 7 DemoTetrad/tmp/fasta/WTH111_ch7_hexadecad_008C.fa output: WTH115 1 DemoTetrad/tmp/fasta/WTH115_ch1_hexadecad_008G.fa output: WTH115 2 DemoTetrad/tmp/fasta/WTH115_ch2_hexadecad_008G.fa output: WTH115 3 DemoTetrad/tmp/fasta/WTH115_ch3_hexadecad_008G.fa output: WTH115 4 DemoTetrad/tmp/fasta/WTH115_ch4_hexadecad_008G.fa output: WTH115 5 DemoTetrad/tmp/fasta/WTH115_ch5_hexadecad_008G.fa output: WTH115 6 DemoTetrad/tmp/fasta/WTH115_ch6_hexadecad_008G.fa output: WTH115 7 DemoTetrad/tmp/fasta/WTH115_ch7_hexadecad_008G.fa output: WTH119 1 DemoTetrad/tmp/fasta/WTH119_ch1_hexadecad_008K.fa output: WTH119 2 DemoTetrad/tmp/fasta/WTH119_ch2_hexadecad_008K.fa

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output: WTH119 3 DemoTetrad/tmp/fasta/WTH119_ch3_hexadecad_008K.fa output: WTH119 4 DemoTetrad/tmp/fasta/WTH119_ch4_hexadecad_008K.fa output: WTH119 5 DemoTetrad/tmp/fasta/WTH119_ch5_hexadecad_008K.fa output: WTH119 6 DemoTetrad/tmp/fasta/WTH119_ch6_hexadecad_008K.fa output: WTH119 7 DemoTetrad/tmp/fasta/WTH119_ch7_hexadecad_008K.fa save: DemoTetrad.json next step: detect GC content %, AT-rich blocks, centromere, telomeres command: node ./src/make_GC_AT_table.js -dataset DemoTetrad.json -w 500 -m 6

2. Step 2: Determination of chromosome-wide guanine-cytosine (GC) contents. [name of user account TSETA]$ node ./src/make_GC_AT_table.js -dataset DemoTetrad.json -w 500bp -m ≥ 6%

3. Step 3: Sequential (50 to 30 ) slicing of the query chromosome into smaller fragments (~10 kb) [name of user account TSETA]$ node ./src/slice_all.js -dataset DemoSNP.json

4. Step 4: Execute MAFFT [name of user account TSETA]$ node ./src/run_mafft.js -dataset DemoSNP.json -mafft-algorithm localpair --mafft-maxiterate 1000 --mafft-thread 20

5. Step 5: Define and align the 50S rDNA loci, which locate at ChrVI [name of user account TSETA]$ node ./src/re_align_rDNA.js -dataset DemoSNP.json -rDNA rDNA.fa -chr 6

6. Step 6: Generation of the final output files # The output files of the “SNP calling” mode. [name of user account TSETA]$ node ./src/snp_summary.js -dataset DemoSNP.json Output snp: Task name + _snp.txt Output snv: Task name + _snv.txt Output viewer: Task name + _snp.html

(see Note 5)

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# The output files of the “Tetrad analysis” mode. DemoTetrad.json -min-co 5000 (see Note 5) Output viewer: Task name + .html (see Note 6) Output table 1: Task name + _final_table.txt ( (see Notes 7–9)

4

Notes 1. The parameters used to determine genome-wide GC contents and AT-rich blocks: – w: a sliding window of 500 bp (default) is used to calculate the GC content. – m: The GC contents of AT-rich blocks are
6% lower than that of the average genome sequence. 2. We reported that the NGS-based SNP calling method (i.e., MUMmer [2]) only revealed 604,578 SNVs and 331,491 Indels between QM6a and CBS999.97(MAT1-1) [5]. Thus, TSETA is about eight times more powerful than MUMmer (7,375,810 SNPs versus 936,069 SNPs, respectively) in revealing intraspecific sequence diversity between genetically distinct strains or isolates. 3. It is important to note that the “Recombine Analyzer” function in TSETA ignores all markers harboring hyphen symbols (1n:3, 2n:2, 3n:1 and 4n:0), RIP mutations, or illegitimate mutations [4] to avoid misidentification of false-positive CO and NCO products. Remember, the hyphen symbols were deliberately incorporated by TSETA to create user-friendly and comprehensive graphical representations of all six chromosomes. 4. For mapping genome-wide meiotic recombination products of fungal genomes (genome size ¼ 34 Mb), we recommend having at least 200 GB of RAM for TGS-based genome assembly and TSETA analysis, respectively, as well as 10 GB of free disk space. The TSETA software package was written in JavaScript or ECMAScript. TSETA has two different functional modes: (1) The “SNP” model is a powerful analytic tool for single-nucleotide-resolution comparison of different intraspecies genome. (2) The “Tetrad” mode of TSETA aims for genome-wide identification of genetic variation before and after a single meiotic events. A stable Internet connection is required for the installation and configuration of TSETA, as well as for retrieving the test data. 5. The closest distance between the two adjacent crossovers is 5000 bp.

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Fig. 2 Intuitive visualization of the large rDNA loci. (a) The ChrVI chromosomes of QM6a, CBS999.97(MAT1-1), and the representative F1 progeny are shown as described in Fig. 1. The large internal deletion (57,946 bp) within the ChrVI centromere of the second F1 representative progeny is marked by a vertical black arrow. (b) The large rDNA loci are shown as an array of red arrows, with each red arrow representing an 18S-5.8S-26S rRNA gene cluster

6. The copy numbers of the 18S-5.8S-26S rRNA gene cluster in the four representative F1 progeny are not always identical to those of parental QM6a or CBS999.97(MAT1-1). To this end, TSETA displays the large rDNA loci as an array of red arrows. Each red arrow represents a single 18S-5.8S-26S rRNA gene cluster (see Fig. 2). 7. TSETA Viewer has a zoom function (
105-fold magnification) that can be manually controlled by turning a mouse wheel. The TSETA viewer allows users to conduct instantaneous and continuous visualization of all six near-complete chromosomes from the scale of the full-length chromosome landscape to individual nucleotides (see Fig. 1). For global visualization, regions with sequences identical to QM6a and CBS999.97(MAT1-1) are colored “magenta” or “cyan,” respectively. GC tracts associated with a CO event are

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labeled with “red” bars. Gapped (deletion) regions are colored “white.” Chromosomal regions harboring RIP mutations are colored “black” (see Fig. 1). TSETA also displays all the positions with non-2:2 markers, including 3:1 (in yellow), 4:0 (in green), 1n:3 (in royal blue), 2n:2 (in blue), 3n:1 (in purple), and 4n:0 (in grey), and illegitimate mutations (in orange) (see Fig. 1). For local nucleotide sequence visualization, QM6a-like and CBS999.97(MAT1-1)-like sequences are colored “pale turquoise” and “pink,” respectively (see Fig. 1e). All non-2:2 markers and the 2:2 markers located next to the boundaries of COs are also highlighted in local nucleotide sequences (in “cyan” and “magenta,” respectively) (see Fig. 1e). 8. Accuracy assessment of TSETA has been described in details recently [4]. 9. Assessment of the quality of TGS assembly has also been described in details recently [4].

Acknowledgments This work was supported by Academia Sinica, Taiwan, Republic of China [AS-105-TP-B07 and AS108-TP-B07 to TFW] and the Ministry of Science and Technology, Taiwan, Republic of China [MOST106-2311-B-001-016-MY3 and MOST109-2311-B-001008-MY3 to TFW]. Hou-Cheng Liu and Wan-Chen Li contributed equally to this work. References 1. Li WC, Huang CH, Chen CL, Chuang YC, Tung SY, Wang TF (2017) Trichoderma reesei complete genome sequence, repeat-induced point mutation, and partitioning of CAZyme gene clusters. Biotechnol Biofuels 10:170. https://doi.org/10.1186/s13068-017-0825x 2. Marcais G, Delcher AL, Phillippy AM, Coston R, Salzberg SL, Zimin A (2018) MUMmer4: A fast and versatile genome alignment system. PLoS Comput Biol 14(1): e1005944. https://doi.org/10.1371/journal. pcbi.1005944 3. Lin HN, Hsu WL (2019) MapCaller – An integrated and efficient tool for short-read mapping and variant calling using highthroughput sequenced data. BioRxiv. https:// doi.org/10.1101/783605 4. Li WC, Liu HC, Lin YJ, Tung SY, Wang TF (2020) Third-generation sequencing-based

mapping and visualization of singel nucleotide polymorphism, meiotic recombination, illegitimate mutation and repeat-induced point mutation. NAR Genomics Bioinformatics 2 (3). https://doi.org/10.1093/nargab/ lqaa056. 5. Li W-C, Chuang Y-C, Chen C-L, Wang T-F (2016) Hybrid infertility: the dilemma or opportunity of applying sexual development to improve Trichoderma reesei industrial strains. In: Schmoll M, Dattenbo¨ck C (eds) Gene expression systems in fungi: advancements and applications. Springer International Publishing, Cham, pp 351–359. https://doi. org/10.1007/978-3-319-27,951-0_16 6. Li WC, Chuang YC, Chen CL, Timofejeva L, Pong WL, Chen YJ, Wang CL, Wang TF (2019) Two different pathways for initiation of Trichoderma reesei Rad51-only meiotic recombination. 2019/05/21 edn., BioRxiv. https://doi.org/10.1101/644443

Tetrad Analysis Using PacBio SMRT Technology 7. Seidl V, Seibel C, Kubicek CP, Schmoll M (2009) Sexual development in the industrial workhorse Trichoderma reesei. Proc Natl Acad Sci U S A 106(33):13909–13914. https://doi. org/10.1073/pnas.0904936106 8. Chuang YC, Li WC, Chen CL, Hsu PW, Tung SY, Kuo HC, Schmoll M, Wang TF (2015) Trichoderma reesei meiosis generates segmentally aneuploid progeny with higher xylanaseproducing capability. Biotechnol Biofuels 8:30. https://doi.org/10.1186/s13068-015-02026 9. Aramayo R, Selker EU (2013) Neurospora crassa, a model system for epigenetics research. Cold Spring Harb Perspect Biol 5(10): a017921. https://doi.org/10.1101/ cshperspect.a017921 10. Gladyshev E (2017) Repeat-induced point mutation and other genome defense mechanisms in fungi. Microbiol Spectr 5 (4). https:// doi.org/10.1128/microbiolspec.FUNK0042-2017 11. Li WC, Chen CL, Wang TF (2018) Repeatinduced point (RIP) mutation in the industrial workhorse fungus Trichoderma reesei. Appl Microbiol Biotechnol 102(4):1567–1574. https://doi.org/10.1007/s00253-017-87315 12. Katoh K, Standley DM (2013) MAFFT multiple sequence alignment software version 7: improvements in performance and usability. Mol Biol Evol 30(4):772–780. https://doi. org/10.1093/molbev/mst010 13. Nakamura T, Yamada KD, Tomii K, Katoh K (2018) Parallelization of MAFFT for largescale multiple sequence alignments.

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Bioinformatics 34(14):2490–2492. https:// doi.org/10.1093/bioinformatics/bty121 14. Katoh K, Rozewicki J, Yamada KD (2019) MAFFT online service: multiple sequence alignment, interactive sequence choice and visualization. Brief Bioinform 20 (4):1160–1166. https://doi.org/10.1093/ bib/bbx108 15. Lassmann T, Sonnhammer EL (2005) Kalign-an accurate and fast multiple sequence alignment algorithm. BMC Bioinformatics 6:298. https://doi.org/10.1186/1471-2105-6-298 16. Altschul SF, Gish W, Miller W, Myers EW, Lipman DJ (1990) Basic local alignment search tool. J Mol Biol 215(3):403–410. https:// doi.org/10.1016/S0022-2836(05)80360-2 17. Mount DW (2007) Using the Basic Local Alignment Search Tool (BLAST). CSH Protoc 2007:pdb.top17. https://doi.org/10.1101/ pdb.top17 18. Nambiar M, Smith GR (2016) Repression of harmful meiotic recombination in centromeric regions. Semin Cell Dev Biol 54:188–197. https://doi.org/10.1016/j.semcdb.2016.01. 042 19. Sims J, Copenhaver GP, Schlogelhofer P (2019) Meiotic DNA repair in the nucleolus employs a nonhomologous end-joining mechanism. Plant Cell 31(9):2259–2275. https:// doi.org/10.1105/tpc.19.00367 20. Anderson CM, Chen SY, Dimon MT, Oke A, DeRisi JL, Fung JC (2011) ReCombine: a suite of programs for detection and analysis of meiotic recombination in whole-genome datasets. PLoS One 6(10):e25509. https://doi.org/10. 1371/journal.pone.0025509

INDEX A Ankyrins ............................................................................. 9 Ascospores .........................................80, 82–84, 332, 339 Automating ...................... 120, 121, 130–132, 177, 275, 282–284, 290, 292, 298, 304, 305, 313, 326, 333, 335, 336, 340 Auxotrophic markers ................................................63–71

B Batch cultivation .................................................. 114, 115 Big data.......................................................................... 252 Bio-based economy............................................ 24, 37–40 Bioreactors....................................................113–118, 121 BLASTs ........... 160, 164, 168, 172, 299, 301, 305, 308, 347–349 Blunt-end ligation ...............................178, 199, 214, 215 Bradford assay ...................................................... 115–117

C Capillary gel electrophoresis (CGE) .................. 181, 185, 196, 206, 223, 225, 226 cDNAs .............................. 105, 110, 192, 195, 197, 199, 207–210, 212, 213, 255–257, 264, 267, 298, 300, 302, 304–306, 308 Cellulolytic enzymes .................................................1, 5, 8 Chromosome-level genome sequences........................ 332 Conidia swollen ........................................................73, 74, 154 Copper ...................................................47, 100, 104, 105 CRISPR/Cas9.......................................32, 33, 47, 87, 88

D

cellulase .................................................................... 135 xylanase .................................................................... 136 Erythritol ...............................................49, 114, 116, 118 Exons ........................ 159, 171, 172, 298, 300, 302–304 Expression vectors................................................ 297–309

F Filamentous fungi ..................... 1, 36, 39, 45–47, 73, 79, 87, 88, 119, 120, 135, 136, 147–154, 166, 172, 272, 273, 287, 297, 298, 332 Fixation ................................................................. 150, 151 Fluorescence ..............................148, 150, 153, 188, 192, 195, 204, 206, 220, 222, 225, 226 Fruiting bodies ..............4, 15, 80, 82–84, 317, 332, 339 Fusions.................................................31, 34–36, 48, 298

G Gene duplications ................................................................. 9 expression .............................. 31–33, 47–50, 63, 104, 105, 171, 177, 251, 252, 297, 302 gun ................................................................ 56, 59, 60 integrations................................................... 31, 56, 64 knockdown ..................................................... 100, 104 losses ................................................................. 8, 9, 48 targeted insertions..................................................... 64 tcu1 .......................................................................... 104 Genetic modification....................................................... 87 Genome annotation ................... 312, 320, 322–324, 326 Green fluorescent protein (GFP) ............... 148, 149, 153 gRNA transcription......................................................... 88

H

De novo assembly ............................................................. 9 Dimethyl sulfate (DMS) ....................177–179, 181, 183, 188, 195 DNA barcoding.......3, 12, 157–160, 164, 166, 170–172

Hygromycin............................ 30, 56, 58, 60, 63, 67, 74, 89, 96, 105, 108

E

Identification phylogram .............................................. 169 Image processing.................................122, 123, 128, 130 In vitro footprinting ......................... 177, 192, 194, 195, 204–206, 218, 222–226 In vitro synthesis ............... 192, 200–203, 205, 214–218 In vivo footprint ................................................... 177, 188 [6-FAM]-labeled DNA .................178, 182, 185, 188

Electrophoretic mobility shift assay (EMSA) ........ 192, 194, 195, 201, 203–205, 216, 218–223 Electroporations .......................30, 46, 55, 57, 63, 73–77 Enzyme activity measurement

I

Astrid R. Mach-Aigner and Roland Martzy (eds.), Trichoderma reesei: Methods and Protocols, Methods in Molecular Biology, vol. 2234, https://doi.org/10.1007/978-1-0716-1048-0, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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REESEI:

METHODS

AND

PROTOCOLS interference................................................................ 46 probing with RNAse ...................................... 210, 326 sequencing .....................................195, 214, 251, 320 structures ....................... 88, 192, 194, 195, 223, 251

Internal transcribed spacers of the rRNA gene cluster ..................................................12, 158, 166 Introns ..........................12, 88, 101, 104, 105, 158–161, 167, 171, 172, 298, 300, 302–304

L

S

Lateral gene transfer ......................................................... 7 Lignocellulosic biomass .................................49, 142, 238 Linker-mediated PCR .......................................... 177, 178 Liquid chromatography-high-resolution mass spectrometry (LC-MS/MS) .......... 238, 241, 244, 246–248

Secretome ................................................ 32, 36, 237–248 Sequence similarity search ......... 159, 160, 164–168, 172 Sequencing PacBio long-read............................................ 317, 319 Sanger ............................................................. 161, 164 tags .................................................................. 251, 305 third generation .................................... 312, 331, 332 Sexual crossing .......................................... 7, 80, 316, 332 SNP calling ........................ 332, 336, 349–352, 357, 358 Sodium hydroxide soluble protein...................... 114, 116 Software tools Funannotate ................................................... 322, 323 ivFAST ............................................................ 177, 178 Maker 2.................................................................... 326 Specific codon optimization ........................................... 88 Stable isotope-assisted workflow .................................. 278 Staining ..................... 104, 109, 110, 131, 150–152, 220 Strain engineering .....................................................35, 46 Synthetic biology ............................................... 30, 45–50

M Mating assays.............................................................82, 83 Measurement of enzyme activity.................................. 135 Meiotic recombination products......................... 331–360 Metabolomics .............................273, 274, 276, 287, 291 Multiplexed genome editing .......................................... 87 Mycoparasitism.................................................4, 9, 10, 15

O Oligonucleotides ............................... 160, 161, 164, 172, 180, 184, 194, 197–199, 203, 219, 230, 297–309 Orphan genes .................................................................... 9

P Particle bombardment ................................ 29, 56, 58, 63 Peak normalization ....................................................... 291 Phylogenomics .................................................................. 4 Protein production heterologous expression .................. 1, 31, 33–37, 48, 50, 87, 205 Proteomics proteomic profiling ........................................ 237–248

R Rapid amplification of cDNA ends (RACE) ..................191–193, 195–200, 206–214, 226, 227 Recombinant DNA technology .......................................... 26, 29, 33 expression ............................................................30, 35 proteins ....................................26, 29–31, 33–36, 297 Repeat-induced point mutation ...............................9, 311 Resistance markers .......................................................... 80 Reversible................................................................ 47, 100 RNA interaction with proteins...................... 194, 195, 218, 219, 222

T Tools for genetic engineering...................................46, 47 Transcription factors .............................46, 177, 184, 272 Transcriptomics ......................................... 5, 10, 251, 273 Transformation protoplasts .........................29, 46, 56, 57, 63, 73, 74, 88, 91, 96, 97 Translation elongation factor 1 alpha ...........12, 159, 170 Trichoderma parareesei ...............................................................8, 12 QM6a.......................... 2, 5, 9, 23, 80, 149, 311–328, 332, 339, 340 reesei ............................... 1–15, 23–40, 45–50, 55–61, 63–77, 79–84, 87–97, 113–132, 135–145, 147, 149, 157–173, 177–188, 191–233, 251–268, 271–292, 311–327, 332, 339, 340

W Whole-cell biocatalyst ........................................ 46, 49, 50

X Xylanases .......................... 39, 40, 48, 135–145, 272, 283