Plant Cell and Tissue Differentiation and Secondary Metabolites: Fundamentals and Applications 3030301842, 9783030301842

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Reference Series in Phytochemistry Series Editors: J.-M. Mérillon · K. G. Ramawat

Kishan Gopal Ramawat Halina Maria Ekiert Shaily Goyal  Editors

Plant Cell and Tissue Differentiation and Secondary Metabolites Fundamentals and Applications

Reference Series in Phytochemistry Series Editors Jean-Michel Mérillon Faculty of Pharmaceutical Sciences Institute of Vine and Wine Sciences University of Bordeaux Villenave d’Ornon, France Kishan Gopal Ramawat Department of Botany University College of Science M. L. Sukhadia University Udaipur, Rajasthan, India

This series provides a platform for essential information on plant metabolites and phytochemicals, their chemistry, properties, applications, and methods. By the strictest definition, phytochemicals are chemicals derived from plants. However, the term is often also used to describe the large number of secondary metabolic compounds found in and derived from plants. These metabolites exhibit a number of nutritional and protective functions for human wellbeing and are used e.g. as colorants, fragrances and flavorings, amino acids, pharmaceuticals, hormones, vitamins and agrochemicals. The series offers extensive information on various topics and aspects of phytochemicals, including their potential use in natural medicine, their ecological role, role as chemo-preventers and, in the context of plant defense, their importance for pathogen adaptation and disease resistance. The respective volumes also provide information on methods, e.g. for metabolomics, genetic engineering of pathways, molecular farming, and obtaining metabolites from lower organisms and marine organisms besides higher plants. Accordingly, they will be of great interest to readers in various fields, from chemistry, biology and biotechnology, to pharmacognosy, pharmacology, botany and medicine. The Reference Series in Phytochemistry is indexed in Scopus. More information about this series at http://www.springer.com/series/13872

Kishan Gopal Ramawat • Halina Maria Ekiert • Shaily Goyal Editors

Plant Cell and Tissue Differentiation and Secondary Metabolites Fundamentals and Applications

With 190 Figures and 93 Tables

Editors Kishan Gopal Ramawat Department of Botany University College of Science M. L. Sukhadia University Udaipur, Rajasthan, India

Halina Maria Ekiert Department of Pharmaceutical Botany Faculty of Pharmacy, Jagiellonian University Medical College Kraków, Poland

Shaily Goyal Amarillo, TX, USA

ISSN 2511-834X ISSN 2511-8358 (electronic) ISBN 978-3-030-30184-2 ISBN 978-3-030-30185-9 (eBook) ISBN 978-3-030-30186-6 (print and electronic bundle) https://doi.org/10.1007/978-3-030-30185-9 © Springer Nature Switzerland AG 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. This Springer imprint is published by the registered company Springer Nature Switzerland AG. The registered company address is: Gewerbestrasse 11, 6330 Cham, Switzerland

Preface

Plant cell and tissue differentiation are closely associated with the differentiation of speciality metabolites. We have seen the development of culturing plant cells and tissues from a mere technique to a full biotechnological tool involving various production systems for metabolites production, understanding the production process, and their biosynthesis in a plant cell. There are several books on plant tissue culture describing the fundamental process and its applications for various biological problems. However, there are a few books on the production of secondary metabolites in plant cell and tissue cultures, but there is no book to describe the process of cell and tissue differentiation vis-a-vis production of speciality metabolites. Therefore, this is an attempt to describe the fundamental process of cell and tissue differentiation as involved with speciality metabolites production. The differentiation of tissue and organ leads to root and shoot formation, and it is evident that this process affects the quality of metabolites production, particularly hairy root cultures, transformed with Agrobacterium rhizogenes. Plant cell and tissue cultures, whether grown in shake flasks or bioreactors, are indispensable tools to explore biosynthesis, accumulation, and production of secondary metabolites. Bottlenecks in technology development are being removed to augment the production of these speciality metabolites. This book describes the most recent technological developments in this field, which are grouped into four parts: 1. Cell and Tissue Differentiation and Secondary Metabolites, 2. Production of Secondary Metabolites in Shoot Cultures, 3. Production of Secondary Metabolites in Normal and Hairy Root Cultures, and 4. In Vitro Production Systems for Secondary Metabolites. In all, there are 32 chapters, which include cell cultures grown in shake flasks, differentiation of glands and trichomes, resin canal differentiation and production of resin, untransformed and transformed organized cultures, and various in vitro systems developed for production of secondary metabolites of importance. The book will be useful to postgraduate students studying plant biotechnology in botany, agricultural sciences, and pharmacy. Biotechnological research scientists in biotechnology, drug industries, and academia will find this book very useful.

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Preface

The editors wish to thank all the contributors for their excellent contributions and the staff at Springer, particularly Drs. S. Costa, S. Blago, and J. Klute, for their expert technical cooperation in the completion of this book. India Poland USA 2021

Professor Kishan Gopal Ramawat Professor Halina Maria Ekiert Dr. Shaily Goyal

Contents

1

An Introduction to the Process of Cell, Tissue, and Organ Differentiation, and Production of Secondary Metabolites . . . . . . Kishan Gopal Ramawat

Part I Cell and Tissue Differentiation and Secondary Metabolites . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2

3

4

Glandular Trichomes on the Leaves of Nicotiana tabacum: Morphology, Developmental Ultrastructure, and Secondary Metabolites . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Branka Uzelac, Dragana Stojičić, and Snežana Budimir The Structural Peculiarities of the Leaf Glandular Trichomes: A Review . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . L. E. Muravnik Accumulation of Secondary Metabolites and Improved Size of Glandular Trichomes in Artemisia annua . . . . . . . . . . . . . . . . . . Neha Pandey, Anupam Tiwari, Sanjay Kumar Rai, and Shashi Pandey-Rai

5

A Model for Resin Flow . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Paulo Cabrita

6

Research Progress on the Resin Canal and Raw Lacquer Synthesis of Toxicodendron vernicifluum (Stokes) F.A. Barkley . . . Meng Zhao

Part II 7

Production of Secondary Metabolites in Shoot Cultures . . .

Shoot Organogenesis, Genetic Stability, and Secondary Metabolite Production of Micropropagated Digitalis purpurea . . . Elizabeth Kairuz, Naivy Pérez-Alonso, Geert Angenon, Elio Jiménez, and Borys Chong-Pérez

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Bioreactor-Grown Shoot Cultures for the Secondary Metabolite Production . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Agata Krol, Adam Kokotkiewicz, Agnieszka Szopa, Halina Maria Ekiert, and Maria Luczkiewicz Production of Specific Flavonoids and Verbascoside in Shoot Cultures of Scutellaria baicalensis . . . . . . . . . . . . . . . . . . . . . . . . . . Beata Kawka, Inga Kwiecień, and Halina Maria Ekiert

10

Secondary Metabolites in Shoot Cultures of Hypericum . . . . . . . . Ana Coste, Carmen Pop, Adela Halmagyi, and Anca Butiuc-Keul

11

Different Types of In Vitro Cultures of Schisandra chinensis and Its Cultivar (S. chinensis cv. Sadova): A Rich Potential Source of Specific Lignans and Phenolic Compounds . . . . . . . . . . Agnieszka Szopa, Adam Kokotkiewicz, Marta KlimekSzczykutowicz, Maria Luczkiewicz, and Halina Maria Ekiert

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High Production of Depsides and Other Phenolic Acids in Different Types of Shoot Cultures of Three Aronias: Aronia melanocarpa, Aronia arbutifolia, Aronia 3 prunifolia . . . . . . . . . . Halina Maria Ekiert, Agnieszka Szopa, and Paweł Kubica Neuroprotective Xanthones and Their Biosynthesis in Shoot Cultures of Hoppea fastigiata (Griseb.) C.B. Clarke . . . . . . . . . . . . Utkarsh Ravindra Moon, Pritam Kumar Dey, and Adinpunya Mitra

14

Bioreactor Technology for In Vitro Berry Plant Cultivation . . . . . Ilian Badjakov, Vasil Georgiev, Maria Georgieva, Ivayla Dincheva, Radka Vrancheva, Ivan Ivanov, Diyan Georgiev, Denitsa Hristova, Violeta Kondakova, and Atanas Pavlov

15

Secondary Metabolites of Various Eleuthero (Eleutherococcus senticosus/Rupr. et Maxim./Maxim) Organs Derived from Plants Obtained by Somatic Embryogenesis . . . . . . . . . . . . . . . . . Katarzyna Bączek, Anna Pawełczak, Jarosław L. Przybył, Olga Kosakowska, and Zenon Węglarz

Part III Production of Secondary Metabolites in Normal and Hairy Root Cultures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 16

17

Biotechnological Production of Useful Phytochemicals from Adventitious Root Cultures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Hosakatte Niranjana Murthy, Dayanand Dalawai, Medha A. Bhat, Vijayalaxmi S. Dandin, Kee-Yoeup Paek, and So-Young Park Mass Scale Hairy Root Cultivation of Catharanthus roseus in Bioreactor for Indole Alkaloid Production . . . . . . . . . . . . . . . . . . . Dhara Thakore and Ashok Kumar Srivastava

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365 383

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Bioproduction of Anticancer Podophyllotoxin and Related Aryltretralin-Lignans in Hairy Root Cultures of Linum flavum L. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Sara Mikac, Lucija Markulin, Samantha Drouet, Cyrielle Corbin, Duangjai Tungmunnithum, Reza Kiani, Atul Kabra, Bilal Haider Abbasi, Sullivan Renouard, Avninder Bhambra, Eric Lainé, Randolph R. J. Arroo, Elisabeth Fuss, and Christophe Hano Development of Taxus spp. Hairy Root Cultures for Enhanced Taxane Production . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Katarzyna Sykłowska-Baranek, Grażyna Sygitowicz, and Agnieszka Pietrosiuk Biosynthesis of Biological Active Abietane Diterpenoids in Transformed Root Cultures of Salvia Species . . . . . . . . . . . . . . . . . Łukasz Kuźma Bonediol Production in Bonellia macrocarpa Hairy Root Culture . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Peggy Elizabeth Alvarez-Gutiérrez, Luis Alberto Ruiz-Ramirez, Gregorio del Carmen Godoy-Hernández, and Federico Antonio Gutiérrez-Miceli Withanolide Production in Hairy Root Culture of Withania somnifera (L.) Dunal: A Review . . . . . . . . . . . . . . . . . . . . . . . . . . . Ganeshan Sivanandhan, Natesan Selvaraj, Andy Ganapathi, and Yong Pyo Lim Enhanced Secondary Metabolite Production in Hairy Root Cultures Through Biotic and Abiotic Elicitors . . . . . . . . . . . . . . . . Gurminder Kaur, Pravin Prakash, Rakesh Srivastava, and Praveen Chandra Verma

Part IV Other In Vitro Production Systems for Secondary Metabolites . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 24

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Morphogenesis, Genetic Stability, and Secondary Metabolite Production in Untransformed and Transformed Cultures . . . . . . . Mihir Halder and Sumita Jha

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Secondary Metabolism in Tissue and Organ Cultures of Plants from the Tribe Cichorieae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Anna Stojakowska and Janusz Malarz

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Plant Liquid Cultures as a Source of Bioactive Metabolites Izabela Grzegorczyk-Karolak, Renata Grąbkowska, and Ewelina Piątczak

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Propagation of Southern Sweet-Grass Using In Vitro Techniques as a Method for the Production of Plants Being a Source of Standardized Raw Material . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Katarzyna Bączek, Anna Pawełczak, Ewelina Pióro-Jabrucka, Jarosław L. Przybył, Olga Kosakowska, and Zenon Węglarz Approaches for Modeling and Optimization of the Secondary Metabolite Production by Plant Biotechnology Methods . . . . . . . . Ryad Amdoun, El-Hadi Benyoussef, Ahcene Benamghar, Fatiha Sahli, Nassim Bendifallah, and Lakhdar Khelifi Relationship of Phenolic Metabolism to Growth in Plant and Cell Cultures Under Stress . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Vincenzo Lattanzio In Vitro Systems of Selected Eryngium Species (E. planum, E. campestre, E. maritimum, and E. alpinum) for Studying Production of Desired Secondary Metabolites (Phenolic Acids, Flavonoids, Triterpenoid Saponins, and Essential Oil) . . . . . . . . . . . . . . . . . . . Małgorzata Kikowska and Barbara Thiem

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Biosynthesis and Biotechnological Production of Anticancer Drug Camptothecin in Genus Ophiorrhiza . . . . . . . . . . . . . . . . . . . Ganesan Mahendran and Laiq ur Rahman

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Production of Cholinesterase-Inhibiting Compounds in In Vitro Cultures of Club Mosses . . . . . . . . . . . . . . . . . . . . . . . . . . Wojciech J. Szypuła and Agnieszka Pietrosiuk

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

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About the Editors

Prof. Dr. Kishan Gopal Ramawat is a former professor and head of the Botany Department, M.L. Sukhadia University, Udaipur, India, with long-standing research experience. He received his Ph.D. in plant biotechnology in 1978 from the University of Jodhpur, India, and afterwards joined the university as a faculty member. In 1991, he moved to M.L. Sukhadia University in Udaipur as associate professor and became professor in 2001. Prof. Ramawat served as head of the Department of Botany (2001–2004, 2010–2012), was in charge of the Department of Biotechnology (2003–2004), was a member of the task force on medicinal and aromatic plants in the Department of Biotechnology, Government of India, New Delhi (2002–2005), and coordinated UGC-DRS and DST-FIST programs (2002–2012). Prof. Ramawat had completed his postdoctoral studies at the University of Tours, France, from 1983 to 1985, and later returned to Tours as visiting professor (1991). He also visited the University of Bordeaux 2, France, several times as visiting professor (1995, 1999, 2003, 2006, 2010), and in 2005, he visited Poland for an academic exchange program (2005). Through these visits in France, Prof. Ramawat and Prof. Mérillon established a strong connection, which has resulted in productive collaborations and several book and reference work publications. Prof. Ramawat has published more than 170 well-cited peer-reviewed papers and articles and has edited several books and reference works on topics such as the biotechnology of medicinal plants, secondary metabolites, bioactive molecules, herbal drugs, and many other topics. His research was funded by several funding agencies. In his research group, Prof. Ramawat has supervised doctoral thesis of 25 students. xi

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About the Editors

He is an active member of several academic bodies, associations, and editorial boards of journals. Professor Halina Maria Ekiert since 2015 is the head of chair and Department of Pharmaceutical Botany, Pharmaceutical Faculty, Medical College, Jagiellonian University, Kraków (Poland). Her scientific career was associated first with the Pharmaceutical Faculty at the Medical Academy in Kraków (Poland) and after reorganization (since 1993) with the Pharmaceutical Faculty at the Medical College at Jagiellonian University. In the years 1999–2014, she was acting head of chair and Department of Pharmaceutical Botany. Her areas of scientific interest are associated mainly with pharmaceutical sciences with strong background in plant biotechnology, phytochemistry, analysis of natural products, and biological activity of plant secondary metabolites. Her biotechnological interests include medicinal and/or cosmetic plant in vitro cultures, endogenic production of bioactive plant secondary metabolites, and biotransformations of exogenic substrates in in vitro cultures. Coumarins, phenolic acids, flavonoids, schisandra lignans, phenylpropanoid glycosides, iridoids, catechins, and arbutin are the special objects of her interest. Throughout her career, Prof. Ekiert received a few postdoctoral internships at German Universities (Bonn – 1998, Würzburg – 1998, Marburg am Lahn – 2000, two trainings). The trainings in Bonn and in Marburg were supported by DAAD – German Academic Exchange Service. Her scientific achievements include approximately more than 120 published articles with total number of citation of approximately 1040 and H-index of 20 (according to Web of Science), few book chapters (published by Springer, Science Publisher, and Studium Press), and the role of co-editor and/or editor at Springer and also guest editor in the MDPI journal – Molecules. Prof. Ekiert cooperated with Würzburg University, and currently she cooperates with Technical University of Braunschweig (Germany) and with the University of Messina (Sicily, Italy). She is an academic teacher with extensive and broad experience in pharmaceutical botany, plant biotechnology, and phytochemistry. She has guided Ph.D. students and candidates for habilitation in the field of plant biotechnology.

About the Editors

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Dr. Shaily Goyal is currently affiliated with West Texas A&M University, Canyon, Texas, USA. She is actively working on pm2.5 and other aeroallergens. She had her Ph.D. in plant biotechnology. After receiving her Ph.D. in 2008 from Mohanlal Sukhadia University (India), she worked as research associate in the Laboratory of Biomolecular Technology, Department of Botany, M.L. Sukhadia University, and also as a teaching assistant of biology, botany, and biotechnology-related courses. Dr. Goyal has been collaborating with Prof. Ramawat since 2005 and has published several book chapters and research articles in peer-reviewed journals. She also co-authored two textbooks for graduate and postgraduate students.

Contributors

Bilal Haider Abbasi Laboratoire de Biologie des Ligneux et des Grandes Cultures, INRA USC1328, Orleans University, Orléans Cedex 2, France Department of Biotechnology, Quaid-i-Azam University, Islamabad, Pakistan Peggy Elizabeth Alvarez-Gutiérrez Tecnológico Nacional de México/Instituto Tecnológico de Tuxtla Gutiérrez, Tuxtla Gutiérrez, Chiapas, México CONACYT-TecNM-Instituto Tecnológico de Tuxtla Gutiérrez, Tuxtla Gutiérrez, Chiapas, México Ryad Amdoun Institut National de Recherche Forestière (INRF), Baïnem, Alger, Algeria Geert Angenon Laboratory of Plant Genetics, Vrije Universiteit Brussel (VUB), Brussels, Belgium Randolph R. J. Arroo Faculty of Health and Life Sciences, De Montfort University, Leicester, UK Katarzyna Bączek Laboratory of New Herbal Products, Department of Vegetable and Medicinal Plants, Warsaw University of Life Sciences – SGGW, Warsaw, Poland Ilian Badjakov AgroBioInstitute, Agricultural Academy, Sofia, Bulgaria Ahcene Benamghar ENSTP, Alger, Algeria Nassim Bendifallah Institut National de Recherche Forestière (INRF), Baïnem, Alger, Algeria El-Hadi Benyoussef Laboratoire de Valorisation des Energies Fossiles, Ecole Nationale Polytechnique, Alger, Algeria Avninder Bhambra Faculty of Health and Life Sciences, De Montfort University, Leicester, UK Medha A. Bhat Department of Botany, Karnatak University, Dharwad, India xv

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Contributors

Snežana Budimir Department of Plant Physiology, Institute for Biological Research “Siniša Stanković” - National Institute of Republic of Serbia, University of Belgrade, Belgrade, Serbia Anca Butiuc-Keul Department of Molecular Biology and Biotechnology, Faculty of Biology and Geology, Babeş-Bolyai University, Cluj-Napoca, România Paulo Cabrita IAPN – Institute of Applied Plant Nutrition, Georg-August University of Göttingen, Göttingen, Germany Borys Chong-Pérez Sociedad de Investigación y Servicios BioTECNOS Ltda, San Javier, Chile Cyrielle Corbin Laboratoire de Biologie des Ligneux et des Grandes Cultures, INRA USC1328, Orleans University, Orléans Cedex 2, France Ana Coste Department of Experimental Biology and Biochemistry, Institute of Biological Research Cluj-Napoca, Cluj-Napoca, România Dayanand Dalawai Department of Botany, Karnatak University, Dharwad, India Vijayalaxmi S. Dandin Department of Botany, Karnatak University, Dharwad, India Pritam Kumar Dey Agricultural and Food Engineering Department, Natural Product Biotechnology Group, Indian Institute of Technology Kharagpur, Kharagpur, West Bengal, India Ivayla Dincheva AgroBioInstitute, Agricultural Academy, Sofia, Bulgaria Samantha Drouet Laboratoire de Biologie des Ligneux et des Grandes Cultures, INRA USC1328, Orleans University, Orléans Cedex 2, France Halina Maria Ekiert Department of Pharmaceutical Botany, Faculty of Pharmacy, Jagiellonian University, Medical College, Kraków, Poland Elisabeth Fuss Interfaculty Institute of Biochemistry, University of Tübingen, Tübingen, Germany Andy Ganapathi Department of Biotechnology, School of Life Sciences, Bharathidasan University, Tiruchirappalli, Tamil Nadu, India Diyan Georgiev Research Institute of Mountain Stockbreeding and Agriculture, Agricultural Academy, Triyan, Bulgaria Vasil Georgiev Laboratory of Applied Biotechnologies, Institute of Microbiology, Bulgarian Academy of Sciences, Plovdiv, Bulgaria Maria Georgieva Research Institute of Mountain Stockbreeding and Agriculture, Agricultural Academy, Triyan, Bulgaria

Contributors

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Gregorio del Carmen Godoy-Hernández Unidad de Bioquímica y Biología Molecular de Plantas, Centro de Investigación Científica de Yucatán A.C., Mérida, Yucatán, México Renata Grąbkowska Department of Biology and Pharmaceutical Botany, Medical University of Lodz, Lodz, Poland Izabela Grzegorczyk-Karolak Department of Biology and Pharmaceutical Botany, Medical University of Lodz, Lodz, Poland Federico Antonio Gutiérrez-Miceli Tecnológico Nacional de México/Instituto Tecnológico de Tuxtla Gutiérrez, Tuxtla Gutiérrez, Chiapas, México Mihir Halder Department of Botany, Barasat Government College, Kolkata, West Bengal, India Adela Halmagyi Department of Experimental Biology and Biochemistry, Institute of Biological Research Cluj-Napoca, Cluj-Napoca, România Christophe Hano Laboratoire de Biologie des Ligneux et des Grandes Cultures, INRA USC1328, Orleans University, Orléans Cedex 2, France Denitsa Hristova Research Institute of Mountain Stockbreeding and Agriculture, Agricultural Academy, Triyan, Bulgaria Ivan Ivanov Department of Organic Chemistry and Inorganic Chemistry, University of Food Technologies, Plovdiv, Bulgaria Sumita Jha Department of Botany, Centre of Advanced Study, Calcutta University, Kolkata, West Bengal, India Elio Jiménez Florida Crystals Corp, Belle Glade, FL, USA Atul Kabra I.K. Gujral Punjab Technical University, Kapurthala, Punjab, India Elizabeth Kairuz Departamento de Biología, Facultad de Ciencias Agropecuarias, Universidad Central Marta Abreu de Las Villas (UCLV), Santa Clara, Cuba Instituto de Biotecnología de las Plantas (IBP), Universidad Central Marta Abreu de Las Villas, Santa Clara, Cuba Laboratory of Plant Genetics, Vrije Universiteit Brussel (VUB), Brussels, Belgium Gurminder Kaur Institute of Bioscience and Technology, Shri Ramswaroop Memorial University, Barabanki, Uttar Pradesh, India Beata Kawka Chair and Department of Pharmaceutical Botany, Jagiellonian University, Medical College, Kraków, Poland Lakhdar Khelifi Laboratoire des Ressources Génétiques et Biotechnologie, ENSA, Ecole Nationale Supérieure Agronomique – Kasdi merbah, Alger, Algeria Reza Kiani Department of Horticultural Sciences, College of Agriculture and Natural Resources, University of Tehran, Karaj, Iran

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Contributors

Małgorzata Kikowska Department of Pharmaceutical Botany and Plant Biotechnology, Poznan University of Medicinal Sciences, Poznań, Poland Marta Klimek-Szczykutowicz Department of Pharmaceutical Jagiellonian University, Collegium Medicum, Kraków, Poland

Botany,

Adam Kokotkiewicz Department of Pharmacognosy, Faculty of Pharmacy, Medical University of Gdańsk, Gdańsk, Poland Violeta Kondakova AgroBioInstitute, Agricultural Academy, Sofia, Bulgaria Olga Kosakowska Laboratory of New Herbal Products, Department of Vegetable and Medicinal Plants, Warsaw University of Life Sciences – SGGW, Warsaw, Poland Agata Krol Department of Pharmacognosy, Faculty of Pharmacy, Medical University of Gdańsk, Gdańsk, Poland Paweł Kubica Department of Pharmaceutical Botany, Jagiellonian University, Collegium Medicum, Kraków, Poland Łukasz Kuźma Department of Biology and Pharmaceutical Botany, Medical University of Łódź, Łódź, Poland Inga Kwiecień Chair and Department of Pharmaceutical Botany, Jagiellonian University, Medical College, Kraków, Poland Eric Lainé Laboratoire de Biologie des Ligneux et des Grandes Cultures, INRA USC1328, Orleans University, Orléans Cedex 2, France Vincenzo Lattanzio Department of Sciences of Agriculture, Food and Environment, University of Foggia, Foggia, Italy Yong Pyo Lim Molecular Genetics and Genomics Laboratory, Department of Horticulture, College of Agriculture and Life Sciences, Chungnam National University, Daejeon, South Korea Maria Luczkiewicz Department of Pharmacognosy, Faculty of Pharmacy, Medical University of Gdańsk, Gdańsk, Poland Ganesan Mahendran Plant Biotechnology Division, Central Institute of Medicinal and Aromatic Plants (CSIR-CIMAP), Lucknow, India Janusz Malarz Maj Institute of Pharmacology, Polish Academy of Sciences, Krakow, Poland Lucija Markulin Laboratoire de Biologie des Ligneux et des Grandes Cultures, INRA USC1328, Orleans University, Orléans Cedex 2, France Department of Molecular Biology, Faculty of Science, University of Zagreb, Zagreb, Croatia

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Sara Mikac Laboratoire de Biologie des Ligneux et des Grandes Cultures, INRA USC1328, Orleans University, Orléans Cedex 2, France International Centre for Cancer Vaccine Science, University of Gdansk, Gdansk, Poland Adinpunya Mitra Agricultural and Food Engineering Department, Natural Product Biotechnology Group, Indian Institute of Technology Kharagpur, Kharagpur, West Bengal, India Utkarsh Ravindra Moon Agricultural and Food Engineering Department, Natural Product Biotechnology Group, Indian Institute of Technology Kharagpur, Kharagpur, West Bengal, India Department of Microbiology, Mahatma Gandhi College of Science, Gadchandur, Maharashtra, India L. E. Muravnik Laboratory of Plant Anatomy and Morphology, Komarov Botanical Institute of the Russian Academy of Sciences, St. Petersburg, Russia Hosakatte Niranjana Murthy Department of Botany, Karnatak University, Dharwad, India Department of Horticulture, Division of Animal, Horticultural and Food Sciences, Chungbuk National University, Chenogju, Republic of Korea Kee-Yoeup Paek Department of Horticulture, Division of Animal, Horticultural and Food Sciences, Chungbuk National University, Chenogju, Republic of Korea Neha Pandey Laboratory of Morphogenesis, Centre of Advanced Study in Botany, Institute of Science, Banaras Hindu University, Varanasi, Uttar Pradesh, India Department of Botany, CMP Degree College (A Constituent PG College of University of Allahabad), Prayagraj, Uttar Pradesh, India Shashi Pandey-Rai Laboratory of Morphogenesis, Centre of Advanced Study in Botany, Institute of Science, Banaras Hindu University, Varanasi, Uttar Pradesh, India So-Young Park Department of Horticulture, Division of Animal, Horticultural and Food Sciences, Chungbuk National University, Chenogju, Republic of Korea Atanas Pavlov Laboratory of Applied Biotechnologies, Institute of Microbiology, Bulgarian Academy of Sciences, Plovdiv, Bulgaria Department of Analytical Chemistry and Physic Chemistry, University of Food Technologies, Plovdiv, Bulgaria Anna Pawełczak Laboratory of New Herbal Products, Department of Vegetable and Medicinal Plants, Warsaw University of Life Sciences – SGGW, Warsaw, Poland

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Contributors

Naivy Pérez-Alonso Instituto de Biotecnología de las Plantas (IBP), Universidad Central Marta Abreu de Las Villas, Santa Clara, Cuba Ewelina Piątczak Department of Biology and Pharmaceutical Botany, Medical University of Lodz, Lodz, Poland Agnieszka Pietrosiuk Department of Pharmaceutical Biology and Medicinal Plant Biotechnology, Faculty of Pharmacy, Medical University of Warsaw, Warsaw, Poland Ewelina Pióro-Jabrucka Laboratory of New Herbal Products, Department of Vegetable and Medicinal Plants, Warsaw University of Life Sciences – SGGW, Warsaw, Poland Carmen Pop Department of Drug Industry and Pharmaceutical Biotechnology, Faculty of Pharmacy, University of Medicine and Pharmacy “Iuliu Haţieganu” Cluj-Napoca, Cluj-Napoca, România Pravin Prakash Molecular Biology and Biotechnology Division, Council of Scientific and Industrial Research-National Botanical Research Institute (CSIR-NBRI), Lucknow, Uttar Pradesh, India Jarosław L. Przybył Laboratory of New Herbal Products, Department of Vegetable and Medicinal Plants, Warsaw University of Life Sciences – SGGW, Warsaw, Poland Laiq ur Rahman Plant Biotechnology Division, Central Institute of Medicinal and Aromatic Plants (CSIR-CIMAP), Lucknow, India Sanjay Kumar Rai Department of Horticulture, Dr. Rajendra Prasad Central Agricultural University, Pusa, Samastipur, India Kishan Gopal Ramawat Department of Botany, University College of Science, M. L. Sukhadia University, Udaipur, Rajasthan, India Sullivan Renouard Laboratoire de Biologie des Ligneux et des Grandes Cultures, INRA USC1328, Orleans University, Orléans Cedex 2, France Institut de Chimie et de Biologie des Membranes et des Nano-objets, CNRS UMR 5248, Université de Bordeaux, Pessac, France Luis Alberto Ruiz-Ramirez Tecnológico Nacional de México/Instituto Tecnológico de Tuxtla Gutiérrez, Tuxtla Gutiérrez, Chiapas, México Fatiha Sahli Institut National de Recherche Forestière (INRF), Baïnem, Alger, Algeria Natesan Selvaraj Department of Botany, Periyar E. V. R College (Autonomous), Tiruchirappalli, Tamil Nadu, India Ganeshan Sivanandhan Molecular Genetics and Genomics Laboratory, Department of Horticulture, College of Agriculture and Life Sciences, Chungnam National University, Daejeon, South Korea

Contributors

xxi

Ashok Kumar Srivastava Department of Biochemical Engineering and Biotechnology, Indian Institute of Technology Delhi, New Delhi, India Rakesh Srivastava Molecular Biology and Biotechnology Division, Council of Scientific and Industrial Research-National Botanical Research Institute (CSIRNBRI), Lucknow, Uttar Pradesh, India Anna Stojakowska Maj Institute of Pharmacology, Polish Academy of Sciences, Krakow, Poland Dragana Stojičić Department of Biology and Ecology, Faculty of Sciences and Mathematics, University of Niš, Niš, Serbia Grażyna Sygitowicz Department of Clinical Chemistry and Laboratory Diagnostics, Faculty of Pharmacy, Medical University of Warsaw, Warsaw, Poland Katarzyna Sykłowska-Baranek Department of Pharmaceutical Biology and Medicinal Plant Biotechnology, Faculty of Pharmacy, Medical University of Warsaw, Warsaw, Poland Agnieszka Szopa Department of Pharmaceutical Botany, Faculty of Pharmacy, Jagiellonian University, Medical College, Kraków, Poland Wojciech J. Szypuła Department of Pharmaceutical Biology and Medicinal Plants Biotechnology, Faculty of Pharmacy, The Medical University of Warsaw, Warsaw, Poland Dhara Thakore Department of Biochemical Engineering and Biotechnology, Indian Institute of Technology Delhi, New Delhi, India Barbara Thiem Department of Pharmaceutical Botany and Plant Biotechnology, Poznan University of Medicinal Sciences, Poznań, Poland Anupam Tiwari Department of Botany, School of Bioengineering and Biosciences, Lovely Professional University, Phagwara, Punjab, India Duangjai Tungmunnithum Laboratoire de Biologie des Ligneux et des Grandes Cultures, INRA USC1328, Orleans University, Orléans Cedex 2, France Department of Pharmaceutical Botany, Faculty of Pharmacy, Mahidol University, Bangkok, Thailand Branka Uzelac Department of Plant Physiology, Institute for Biological Research “Siniša Stanković” - National Institute of Republic of Serbia, University of Belgrade, Belgrade, Serbia Praveen Chandra Verma Molecular Biology and Biotechnology Division, Council of Scientific and Industrial Research-National Botanical Research Institute (CSIR-NBRI), Lucknow, Uttar Pradesh, India Academy of Scientific and Innovative Research (AcSIR), Council of Scientific and Industrial Research, Ghaziabad, Uttar Pradesh, India

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Contributors

Radka Vrancheva Department of Analytical Chemistry and Physic Chemistry, University of Food Technologies, Plovdiv, Bulgaria Zenon Węglarz Laboratory of New Herbal Products, Department of Vegetable and Medicinal Plants, Warsaw University of Life Sciences – SGGW, Warsaw, Poland Meng Zhao Shanxi Normal University, Linfen, People’s Republic of China

1

An Introduction to the Process of Cell, Tissue, and Organ Differentiation, and Production of Secondary Metabolites Kishan Gopal Ramawat

Contents 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Historical Developments in Secondary Metabolites Production . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Plant Growth Regulators Used in Plant Tissue Culture . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1 Role of Plant Growth Regulators . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2 Plant Growth Regulators Used for Differentiation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 Cell and Tissue Differentiation and Production of Secondary Metabolites . . . . . . . . . . . . . . . . . 5 Organized Culture for the Production of Secondary Metabolites . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.1 Somatic Embryogenesis and Metabolic Studies . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.2 Shoot Culture and Secondary Metabolites . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.3 Roots Culture and Secondary Metabolites . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.4 Bioreactor Culture . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6 Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

2 6 7 7 9 10 12 12 14 15 17 17 18

Abstract

Plant cell and tissue cultures are indispensable tools to understand the process of differentiation that has become a powerful technology for commercial level production of micropropagated plants. Various technological innovations in growing and analyzing cell cultures have led to the production of useful primary and secondary metabolites from plant cell cultures using bioreactors up to a 75,000 L capacity. The processes of differentiation as well as production of secondary metabolites are complex, both involving a coordinated expression of several genes. Differentiation of cells and tissues causes qualitative and quantitative changes in the production of secondary metabolites. Metabolomics and transcriptome analysis of cells and tissues will provide more insight into genes K. G. Ramawat (*) Department of Botany, University College of Science, M. L. Sukhadia University, Udaipur, Rajasthan, India e-mail: [email protected]; [email protected] © Springer Nature Switzerland AG 2021 K. G. Ramawat et al. (eds.), Plant Cell and Tissue Differentiation and Secondary Metabolites, Reference Series in Phytochemistry, https://doi.org/10.1007/978-3-030-30185-9_35

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involved in biosynthesis of secondary metabolites. In this brief overview, process of differentiation of cells, tissues, and organs and production of secondary metabolites as affected by plant cell and tissue differentiation are presented. The importance of unorganized and organized cultures as production system is also discussed. Keywords

Cell culture · Secondary metabolites · Bioreactor · Differentiation · Hairy roots · Plant growth regulators

1

Introduction

Plant cell produces two types of metabolites: primary metabolites involved directly in growth and metabolism, viz., carbohydrates, lipids, and proteins, and secondary metabolites considered as end products of primary metabolism and are in general not involved in metabolic activity, viz., alkaloids, phenolics, and terpenes. Primary metabolites are produced as a result of photosynthesis, and these products are further involved in the cell component synthesis and stored as reserve food. Primary metabolites obtained from higher plants for commercial use are high volume low value bulk chemicals. They are primarily used as industrial raw materials, foods, or food additives, for example, vegetable oils, carbohydrates (sucrose, starch, pectin, and cellulose), and proteins (Fig. 1). Secondary metabolites are compounds biosynthetically derived from the primary metabolites but are of limited occurrence in the plant kingdom, may be restricted to a

Carbohydrate: starch in potato

Protein: various pulses

Triglyceride: vegetable oils like sunflower oil

Fig. 1 Three primary metabolites commonly present in storage organs of plants

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An Introduction to the Process of Cell, Tissue, and Organ Differentiation. . .

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particular taxonomic group (genus, species or family) and not required for the growth of plant. Medicinal plants are rich in secondary metabolites that is why these plants are termed as “medicinal” or “officinal” plants. Still 15% of flora has been explored phytochemically, and thus the scope of work is enormous [1, 2]. The required carbon skeleton is derived from carbohydrates synthesized from photosynthesis. The other major primary products involved in various classes of secondary metabolites are amino acids. Acetyl-CoA and mevalonic acid play a key role in the synthesis of various terpenoids, while skikimic acid pathway is involved in the synthesis of lignins and indole alkaloids. These secondary metabolites are synthesized by coordinated expression of several genes producing enzymes required for step by step biosynthesis. Genes responsible for synthesis of such secondary metabolites are highly expressive in some organs where these compounds are accumulated. However, there are examples such as nicotine (pyrrolidine alkaloid of Nicotiana species), which is mostly biosynthesized in roots but transported and accumulated in leaves. Therefore, specific mechanisms that evolved for such functions are also energy expensive involving ATP binding transporters [3, 4]. Out of several thousand genes present in plants, it is not clear how some of these genes are expressed in a coordinated manner. Interest in medicinal plants as a source of bioactive natural molecules has gained again in spite of intrinsic difficulties as combinatorial chemistry [comprising chemical synthetic methods that make it possible to prepare a large number (tens to thousands or even millions) of compounds in a single process] failed to deliver sufficient number of new drugs (only 36% out of 1073 molecules) in the recent past [5]. From 1981 to 2014 [6], 136 drugs were recorded effective against cancer in which 17% were synthetic while 83% were of natural origin. Hence medicinal plants are a source of several important medicines, and interest in discovery of new molecules is ever lasting. Some of the plants derived compounds approved by FDA (Food and Drug Administration, USA) in the last three decades as drug, and others which are under clinical trials are presented in Table 1. It is evident that these drugs have complex structures and are used for a wide range of diseases. Complexity of these secondary metabolites prevents their chemical synthesis, and nonavailability of uniform material for extraction is difficult in developing technology toward drug approval. Plant cell and tissue cultures provide an excellent source of continuous and uniform supply of raw material throughout the year without seasonal or geopolitical interferences. Therefore, even in the absence of a major breakthrough in the production of secondary metabolites from plant cell cultures, interest, experimentation, and exploration are continuous [9]. Well-known examples of plant-derived natural compounds that have attained significant reputation in therapy are paclitaxel and its derivatives from yew (Taxus brevifolia), vincristine and vinblastine from Madagascar periwinkle (Catharanthus roseus) and camptothecin and its analogues initially discovered in the Chinese tree Camptotheca acuminata, and cholinesterase inhibitor galanthamine (for the treatment of Alzheimer’s disease) and antimalarial and potential anticancer agent artemisinin derived from the traditional Chinese herb Artemisia annua [5, 10, 11]. About 100 natural compounds and their derivatives are under clinical trials especially for various types of cancers [7, 12]. Secondary metabolites

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Table 1 Selected examples of plant derived natural products entered in drug market as approved therapeutic or under clinical trials in last three decades [5, 7, 8] Generic name and chemical structure Plant species A. Approved therapeutic Artemisia annua Artemisinin

Trade name (year of introduction)

Disease (mechanism of action)

Artemisin (1987)

Malaria treatment (radical formation)

Capsaicin

Capsicum annum

Qutenza (2010)

Postherpetic neuralgia (TRPV1 activator)

Colchicine

Colchicum spp.

Colcrys (2009)

Gout (tubulin binding)

Dronabinol/ Cannabidol Dronabinol

Cannabis sativa

Sativex (2005)

Chronic neuropathic pain (CB1 and CB2 receptor activation)

Galanthamine

Galanthus caucasicus

Razadyne (2001)

Ingenol mebutate

Euphorbia peplus

Picato (2012)

Dementia associated with Alzheimer’s disease (ligand of human nicotinic acetylcholine receptors (nAChRs)) Actinic keratosis (inducer of cell death)

Irinotecan (structure), Topotecan (camptothecin derivatives)

Camptotheca acuminata

Camptosar (2004), Hycamtin (2007)

Solid tumors of colon, lung, and ovarian cancers (topoisomerase I inhibitors)

Omacetaxine mepesuccinate (Homoharringtonine)

Cephalotaxus harringtonia

Synribo (2012)

Oncology (protein translation inhibitor)

(continued)

An Introduction to the Process of Cell, Tissue, and Organ Differentiation. . .

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Table 1 (continued) Generic name and chemical structure Paclitaxel

Sphingomyelin/ cholesterol liposomal vincristine

Plant species Taxus brevifolia

Catharanthus roseus

Trade name (year of introduction) Taxol (Taxotere ® 1993), Abraxanec (2005), Nanoxelc (2007) Cabazitaxel (Jevtana ®) (2010) (Marqibo ®) 2012

Disease (mechanism of action) Cancer chemotherapy (mitotic inhibitor)

Relapsed acute lymphoblastic leukemia (cell-cycle-specific)

OH N

CH2CH3

N COOCH3

• H2SO4

H N

H

CH3O

CH2CH3 H COOCH3 OCOCH3 COOCH3

N H CHO

OH

B. Under clinical trials Betulinic acid Betula spp., Diospyros spp., Syzygium spp., Ziziphus spp., Paeonia spp., Sarracenia flava, Anemone raddeana Lycopodium cernuum Curcuma longa Curcumin (Turmeric) Epigallocatechin-3O-gallate

Camellia sinensis (Green tea)

Genistein

Genista tinctoria

Picropodophyllotoxin

Podophyllum hexandrum (syn. Sinopodophyllum hexandrum)

Various cancers, e.g., colorectal lung, colon, breast, prostate, hepatocellular, bladder, head and neck, stomach, pancreatic, ovarian and cervical carcinoma, glioblastoma, chronic myeloid leukemia cells (induction of apoptosis in mitochondria) 26 different trials: cognitive impairment, different types of cancers, cardiovascular disease etc. (NF-κB inhibition) etc. 14 different trials: different types of cancer, Epstein-Barr virus reactivation, Alzheimer’s disease, cystic fibrosis, diabetic nephropathy, obesity, influenza infection etc. (cell growth arrest and apoptosis induction) 5 different trials: various intestinal and lung cancers, Alzheimer’s disease, osteopenia, osteoporosis (protein-tyrosine kinase inhibitor, antioxidant) Trial for various cancer cells (tubulin binding/IGF-1R Inhibitor)

(continued)

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Table 1 (continued) Generic name and chemical structure Resveratrol

Plant species Vitis vinifera

Trade name (year of Disease (mechanism of introduction) action) 22 different trials: Prediabetes, vascular system injuries, lipid metabolism disorders, gestational diabetes, cardiovascular disease, type 2 diabetes mellitus, inflammation, coronary artery disease, memory impairment, various hypertension and heart problems, Alzheimer’s disease (NF-κB inhibition)

or their derivatives are not only used as medicines, but also in several other preparations such as cosmetics, insecticides, dyes, fragrances and flavors, and several other products. Though secondary metabolites are considered as waste products, their production requires energy; these metabolites are known to involve in various biological processes including plant defense. Therefore, their production is influenced by external abiotic and biotic factors [13]. In brief, following are important functions of secondary metabolites [10, 14–16]: 1. 2. 3. 4. 5. 6. 7. 8. 9.

As plant defense molecule against microbes, fungi and insects Role in plant and also insect reproduction Provide protection to insects against parasites As nitrogen storage molecules Mitotic inhibitors and modulation of microtubule structures Inhibition of DNA and protein synthesis As germination inhibitor (as allelochemical) Involve in plant immunity and programmed cell death Signaling molecules in angiosperm parasites

In this brief overview, importance of medicinal plants and their exploitation using plant cell and tissue culture technology is presented. This chapter discusses how differentiation occurs in cell and tissue cultures and how this differentiation affects the secondary metabolites.

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Historical Developments in Secondary Metabolites Production

Plant cell and tissues are grown aseptically in liquid or on gelled nutrient medium under controlled environmental conditions of light, temperature, and humidity. Since its inception, plant cell and tissue cultures have become an indispensable tool for micropropagation of ornamental and endangered plant species, haploids production

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using anther or pollen culture, development of transgenic plants, production of virusfree plants using meristems, and the study and production of useful metabolites. Plant callus and cell cultures developed with the development of analytical tools and discovery of auxin and cytokinin in plants and consequently their role in plant cell growth modulation and in differentiation [17, 18]. The subsequent landmark works were single cell cultures, plating techniques, and development of MS medium by Murashige and Skoog in 1962 [19]. The details of historical developments in plant biotechnology are described in many recent reviews [20, 21]. Study of secondary metabolites took a momentum with the availability of more sensitive tools of phytochemistry like GLC, TLC with densitometry, and HPLC as tissue mass produced in cultures is very small on dry weight basis. Currently, metabolomics and associated genomics are applied to unravel the biosynthetic pathways, gene involved, and control mechanism of secondary metabolites production [22]. Initial reports were concern about the occurrence of secondary metabolites in culture, followed by development of production medium and attempted production system with two stage culture system. Use of various classical approaches and techniques of genetic engineering for the production and improvement of secondary metabolites are presented in various reviews [20, 23]. The production of secondary metabolites was associated with the development of bioreactor technology followed by genetic transformation using Agrobacterium strains for rooty and shooty characters [24–26]. Growth of tobacco cells in 10 L glass carboy bioreactor in 1959 to 20,000 L industrial level production system in 1977 took only 18 years, whereas the first successful use of bioreactor technology for industrial or commercial production of secondary metabolites is for shikonin from Lithospermum erythrorhizon in 1983. Other successful examples of growth and production of useful metabolites up to 50,000 L bioreactor are scopolamine by Duboisia sp., podophyllotoxin by Podophyllum sp., protoberberine by Coptis japonica, rosmarinic acid by Coleus blumei, echinacea polysaccharides by Echinacea purpurea, arbutin by Catharanthus roseus, vanillin by Vanilla planifolia, and betacyanins by Beta vulgaris [20, 27]. Phyton Biotech (Ahrensburg, Germany) operates the world’s largest 75,000 L bioreactor facility for the production of secondary metabolites like taxol [28]. Several other techniques utilizing unorganized or organized cultures with production medium were feed batch cultures, elicitor, immobilization, and genetic transformation to optimally produce secondary metabolites [9, 29–31]. Employment of these different techniques for the production of secondary metabolites is given in different chapters in this book.

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Plant Growth Regulators Used in Plant Tissue Culture

3.1

Role of Plant Growth Regulators

Plant growth regulators, particularly auxin and cytokinin, play determining role in cell growth and differentiation, differentiation of meristems, and maintenance of apical dominance [32]. It is well established that auxin is synthesized in apical meristem and young leaves of shoots, whereas cytokinin is synthesized in root cap.

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Auxin moves basipetally and maintains a gradient from high to low, whereas cytokinin moves upward. In callus cultures, exogenous supply is given and gradients are not maintained. In plant postembryonic development, stem cells divide in meristem producing daughter cells, which differentiate to produce different organ structures. Cytokinins are known as crucial signaling molecules controlling meristem activity [33, 34]. During this process, the number of meristem cells remained constant, and new cells are produced in such a way that differentiation is achieved without disturbing the number of meristem cells. The actual mechanism controlling this activity is unclear, but during the process of differentiation several pathways are involved including those for lignifications, cellular meristematic activity, and formation of differentiating patterns. Auxin and cytokinin act antagonistically or synergistically to achieve the desired balance, and their concentration is responsible for a different morphological manifestation. It has been shown that auxin and cytokinin both modulate the synthesis and concentration of each other by controlling the signaling pathways (controlled at multiple levels, such as biosynthesis, transport, perception, and signaling) involving PIN family auxin efflux carrier proteins located at cell membrane [33, 35–37]. It is also well established that internal concentrations of auxin are maintained by synthesis, catabolism, and transport, which is influenced by exogenous and endogenous stimuli. Although most of the native auxin indole-3-acetic acid (IAA) is transported into cells by diffusion, the importance of AUX1/LAX carriers (proteins) could be clearly demonstrated in many developmental processes (controlled by AUX/LAX genes). They are involved in embryogenesis [38], hypocotyl apical hook development [39], root gravitropism [40], lateral root development [36, 41, 42], root hair development [43], phloem loading and unloading [44], and phyllotaxis [45]. Hairy root cultures grow on hormone free medium, and such a situation results in high production of secondary metabolites in spite of fairly high growth values. Callus and cell cultures deprived of auxin/hormone by growing on hormone free medium were attempted in Aegle marmelos, Catharanthus roseus, Ochrosia elliptica (in our laboratory), and Salvia miltiorrhiza, which resulted in increased production of their secondary metabolites [46]. Auxin polar transport is responsible for auxin flow and gradient which are critical for proper differentiation patterns of organs [32]. Polar transport of auxin provides essential directional and positional information for the vital plant cell developmental processes such as vascular differentiation, apical dominance, patterning, organ polarity, embryogenesis, organogenesis, phyllotaxis, and tropisms [47]. Auxin efflux inhibitors such as 2,3,5-triiodobenzoic acid (TIBA) or 1-naphthylphthalamic acid (NPA) have provided information about auxin efflux transporters and their involvement in the control of physiological and developmental processes in plants. Compounds that inhibit auxin efflux include synthetic phytotropins [NPA, 2-(1-pyrenoyl) benzoic acid (PBA)], cyclopropyl propane dione (CPD) and TIBA, and also natural flavonoids (e.g., quercetin) [48]. Plant hormones are master regulators of plant growth and development. Basic fundamental knowledge of their transport and metabolism on the cellular and subcellular is therefore crucial to a better understanding of developmental processes in plants [49]. Details of auxin-cytokinin regulation of growth and differentiation are

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beyond the scope of this brief chapter. Therefore, by using appropriate combination and concentration of auxin and cytokinin, shoot, roots, and embryos are produced from undifferentiated mass of cells called callus, which was demonstrated for the first time in tobacco callus cultures [18].

3.2

Plant Growth Regulators Used for Differentiation

Many plant growth regulators have been tried in plant cell and tissue cultures to obtain desirable or novel results including (a) classical plant hormones (auxins, cytokinins, gibberellins, abscisic acid, and ethylene), (b) others synthetic substances (morphactins, dikegulac salt, paclobutrazol, and prohexadione calcium), and (c) those having phytohormone-like regulatory roles (polyamines, oligosaccharins, salicylates, jasmonates, brassinosteroids, dehydrodiconiferyl alcohol glucosides, turgorins, systemin, and some inhibitors). Differential distribution of the plant growth regulatory substance auxin is known to mediate many fundamental processes in plant development, such as the formation of the embryogenic apical–basal axis, pattern formation, tropisms, and organogenesis (reviewed in [50]). Synthetic auxins that differ in the mechanisms of their transport across the plasma membrane have been used together with polar auxin transport inhibitors in many studies to understand their role in plant development. However, the exact mechanism of action of auxin efflux and influx inhibitors has not been fully elucidated [48]. Functions of a few commonly used plant growth regulators in plant cell and callus cultures are presented in (Table 2). It is beyond the scope of this brief chapter to discuss the role

Table 2 Effects of various growth modulating substances on growth and differentiation in unorganized and organized cultures [36, 37, 49] Plant growth regulators Auxin and similar products (IAA, NAA, IBA, 2,4-D, 2,4,5-T, Picloram, Dicamba)

Cytokinin (Kinetin, BA, Zeatin, 2iP, Thidiazuron) Ethylene Ethylene inhibitors (Silver nitrate) Abscisic acid (ABA) Synthetic substances: (Paclobutrazol, Morphactin, Dikegulac, Hexadione) Auxin transport inhibitor: Triiodobenzoic acid (TIBA) Auxin inhibitor (7-Azaindole)

Effects in plant tissue and cell cultures, depending upon concentration and interacting regulator Callus induction, growth and support of callus, induces vascular differentiation, rooting, inhibits differentiation at higher concentration, inhibits secondary metabolites production Cell division, vascular differentiation, induction of shoots, loss of shoot apical dominance, chlorophyll synthesis, and root apical dominance Affects lateral rooting, inhibits differentiation Promotes differentiation Somatic embryo maturation Too variable responses, somatic embryo promotion, suppression of callusing, branching, greening, shorter internodes Promotion of somatic embryos Promotion of somatic embryos

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and interactions of all growth regulators. However, functions are briefly described in the table, and optimal dose is generally achieved by empirical approaches and comparing allied species [29, 51–54].

4

Cell and Tissue Differentiation and Production of Secondary Metabolites

Plant cell and tissue cultures are extensively used to study cellular and tissue differentiation and also production of associated secondary metabolites. Most anthocyanins are glycosylated, accumulated in the vacuoles, and the most studied phenolic compound because they impart flower color. Plant cell culture is an excellent alternative for the production of anthocyanins of uniform quality as compared to in vivo materials like grape skins from wine industry, sweet potato, and red cabbage [55]. Isolated single cells in liquid medium are specifically interesting systems to study secondary wall formation by lignin synthesis using Zinnia elegans and a few other cultures, which is not possible using in vivo plant model [56]. Lignin precursor monolignols are synthesized from phenylalanine through shikimate pathway involving several steps. At low phenylalanine levels, this amino acid is used for protein synthesis [57]. Cytokinin and IAA are key plant growth hormones that regulate root development, its vascular differentiation, and root gravitropism; these two hormones, together with ethylene, regulate lateral root initiation [36]. Pathway for biosynthesis of some compounds (consequently many enzymes or genes) leading to cellular differentiation is same and expressed sequentially in a coordinated manner like a harmonium; keys are same but produce different rhythm when (ex)pressed in different sequences. Secondary metabolites are present in most of the cells of plant in its cytoplasm and vacuole, sometimes in higher amounts in specialized cells called glands, glandular cells, and/or trichomes. The structures of these secretory tissues may vary considerably from single cells (e.g., idioblasts or laticifer cells) to many cells and complex structures (trichomes, colleters, nectaries, osmophores, secretory cavities, and ducts). A laticifer is a single cell or a row of specialized cells that contain latex. Latex, which is cytoplasm of the cell, contains several compounds, minerals, proteins, and specialized compounds such as alkaloids or rubber (isoprene). As compared to laticifers, resin ducts are more complex organized tissues, having epithelium lining and lumen containing resin. This is a phylogenetic conserved trait to have laticifers or resin ducts [58]. Plant secondary metabolites are generally accumulated at high levels in specific tissues or cell types of plants of a genus (e.g., Urticaria) or family (e.g., Rutaceae, Burseraceae). Economically important examples of tissuespecific metabolites include alkaloids (e.g., morphine and codeine) and latex in laticifer cells in poppy, papaya (latex contain papain), and rubber trees (isoprene in latex), terpenes and saponins in epidermal cells of many plant families, and resins in Pinus trees [59]. Special metabolites are synthesized and accumulated in laticifers, resin canals, and specialized tissues. Details of these special structures are not presented here, but this state of tissue differentiation is a prerequisite for the

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production of compounds present in these structures. Therefore, this is a major hurdle in producing such tissue specific secondary metabolites in cell and callus cultures, and production of such compounds remains low, irrespective of various permutation and combinations of the medium or effectors, e.g., in Commiphora wightii [51, 60–63]. Various types of cellular and tissue differentiations obtained in cell and tissue culture are presented in Fig. 2. Lignin is an important structural and defensive component of plant secondary cell walls, and the second most abundant biopolymer on earth. Lignin is composed primarily of three hydroxycinnamyl alcohol monomers, referred to as monolignols. These monolignols are variedly polymerized to form complex structure of lignin. These monolignols are also involved in the formation of suberin with lipids, bark, and cork formation [59]. Lignin formation is an integral part of the process of differentiation, and tracheid-like structures are formed in callus and cell cultures, which ultimately form meristemoids and shoot buds. This complex process involves several precursor substrates and enzymes depending upon the cell type [64]. It is

b

a

c x

g

d f e Fig. 2 Various cellular and tissue differentiation in callus cultures initiated from explant (x). Symbolic photos represent: (a) Unorganized callus; (b) red callus due to presence of anthocyanin; (c) anthocyanin containing cells under light microscope; (d) resin canal showing epithelium lining; (e) L.S. resin canal; (f) secondary wall (sw) formation by lignification under TEM; (g) meristemoid formation as seen under light microscope. (All photos by the author)

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well established that carbon source in the medium has a profound effect on vascularization (secondary wall formation) as basic carbon skeleton is provided by sugars [65]. Terpenoids are the most structurally divergent secondary metabolites. About 25,000 compounds have been isolated, and their structures are elucidated [66]. Accumulation of terpenes is known in many plants, e.g., monoterpenoids in Lamiaceae plants are biosynthesized in secretory cells and accumulate in the epicuticular cavity of glandular trichomes, osmophores, conical-papillate cells, ducts, and cavities [67, 68], while terpenoids of woody plants are secreted into the resin duct. For volatile mono- and sesquiterpenenoids, their emission from flowers of several plants (Caesalpinia pulcherrima, Parkia pendula, Bauhinia rufa, Mucuna urens, Rosa x hybrid) and leaves of conifer and other woody plants is well known [69, 70]. Terpenoids act as defense molecules when herbivores attack the plants and defense mechanism is activated to produce volatiles. It is evident that this mechanism is activated in response to insect attack by gene expression [16]. Morphine, a major isoquinoline alkaloid in the latex of opium poppy, is accumulated in the large membranous vesicles of such latex. Immunofluorescence analyses using antibodies specific for five enzymes of alkaloid formation in opium poppy. Weid and coworkers [71] showed that two O-methyltransferases and an O-acetyltransferase were found predominantly in parenchyma cells within the vascular bundle of capsule and stem, while codeinone reductase was localized to laticifers. Another group reported that three of those biosynthetic enzymes of morphine were localized in sieve elements of this plant [72]. Therefore, it is evident that the transport of the intermediate from specific cell-type of vascular tissue to laticifer was involved with ABC transporters (located on cell membranes).

5

Organized Culture for the Production of Secondary Metabolites

Various explants are used for the initiation of in vitro cultures. Depending upon auxin-cytokinin concentration in the medium, most explants produce callus or organized structures. A schematic development of somatic embryos, morphogenesis/organogenesis leading to shoots formation, multiple adventitious shoot cultures, formation of adventitious roots, or callus mass is presented in Fig. 3. Details of organ formation in different species are briefly presented in the following paragraphs and in the chapters in the book.

5.1

Somatic Embryogenesis and Metabolic Studies

Somatic embryogenesis in cultures follows the same morphogenetic pathway as in vivo development forming globular, heart, torpedo stages, and lastly germination (Fig. 4). This shows that genetic makeup of somatic cells is totipotent. These embryos can be produced directly on explants (direct embryogenesis) or through

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Explant Balanced ratio of Auxin/cytokinin

Changed ratio of Auxin/Cytokinin

Callus

Morphogenesis

Adventitious Roots

High Cytokinin

Somatic embryogenesis

Adventitious Shoots Progress of differentiation Fig. 3 Initiation of various types of organized cultures from an explant. (All photos by the author)

callus formation (indirect somatic embryogenesis). The presence of auxin in medium supports the growth of nonembryonic cells and suppresses induction of somatic embryos but may be required for preserving embryonic characters of cells. Besides propagation, somatic embryogenesis is a valuable model system for investigating the structural, physiological, and molecular events occurring during embryo development (as compared to nonembryonic cells) and genetic transformation, e.g.,

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Germinated SE

Globular SE

Cell cultures

Plantlets

Globular

Torpedo

Fig. 4 Process of somatic embryogenesis as observed in carrot cell cultures plated in Petri dishes after treatment with 2,4-D. (All photos by the author)

differences in cytological characteristics, 1H-NMR analysis of intracellular metabolites, and glycolytic enzyme activities grape ([73], and references therein) and sugarcane cell cultures [74]. However, still somatic embryogenesis and conversion of somatic embryos to plantlets remain a challenging problem in some woody trees, legumes, and some conifers [75]. Therefore, development of high metabolites producing varieties and metabolites associated with differentiation are difficult in such species, e.g., resin producing conifers and plants of Burseraceae [61, 62].

5.2

Shoot Culture and Secondary Metabolites

Organized cultures like shoots grow slowly, and such cultures are likely to accumulate more secondary metabolites as compared to fast growing callus and cell cultures. Shoot cultures are expected to produce secondary metabolites profile similar to that of in vivo plants. These cultures may also show qualitative changes in secondary metabolites profile. Development of shoot buds from unorganized callus mass is a complex phenomenon controlled by cytokinins (in combination with an auxin) and thus affecting several genes related to cell, tissue (conducting tissues), and meristem differentiation thus influencing other pathways and metabolites production. There are several reports of shoot cultures and secondary metabolites content of in vitro grown shoots, e.g., tylophorine in Tylophora indica [76], rosmarinic acid in Melissa officinalis [77], and several others (Glycyrrhiza glabra, Spathiphyllum cannifolium); but detailed information about qualitative and quantitative changes occurring during the process of differentiation is scanty. Some selected examples of secondary

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Table 3 Selected examples of secondary metabolites produced in undifferentiated and differentiated cultures [78–80] Undifferentiated cell and callus cultures Ajmalicine (Catharanthus roseus) Anthocyanin (Vitis vinifera, Commiphora wightii, Nicotiana tabaccum) Capsaicin (Capsicum annuum) Crocin (Crocus sativus) Guggulsterones (Commiphora wightii) Naphthoquinones (Lithospermum erythrorhizon) Nicotine (Nicotiana tabaccum) Resveratrol, viniferin (Vitis vinifera) Stevioside (Stevia reboudiana) Taxol (Taxus brevifolia) Thaumatin (Thaumatococcus danielli) Vanillin (Vanilla planifolia)

Differentiated tissue cultures Artemisinin (Artemisia annua) Bacosides (Bacopa monnieri) Betalaines (Beta bulgaris) Eleutheroside E (Eleutherococcus koreanum) Genistin, puerarin (Pueraria species) Ginsenosides (Panax ginseng) Hypericins (Hypericum species) Platydesminium (Ruta graveolens) Rosmarinic acid (Melissa officinalis) Silymarin (Silybum marianum) Tylophorine (Tylophora indica) Withanolides (Withania somnifera)

metabolites reported from differentiated and undifferentiated cultures are presented in Table 3. Individual secondary metabolites are not discussed here and presented in the chapters in this book. Metabolomic profile of differentiated shoot and undifferentiated cultures can provide more insight about secondary metabolites and associated genomics about gene expression as elucidated in the case of benzoisoquinoline and monoterpenoid indole alkaloids, cannabinoids, caffeine, ginsenosides, withanolides, artemisinin, and taxol [22, 81, 82]. Ease of sequencing has revived the interest in mining genes related to biosynthetic pathways and enzymes involved.

5.3

Roots Culture and Secondary Metabolites

Root cultures offer a unique system to study metabolites associated with roots. Particularly hairy roots offer an advantageous feature that these transformed and organized cultures grow indefinitely without exogenous auxin supply (Fig. 5). Adventitious root cultures require auxin supply in the medium and can be maintained for a fairly long time (several years) with proper subcultures. Because of the use of auxin in the medium, callus formation is another problem associated with these cultures. Presence of auxin is a known suppress secondary metabolites biosynthesis, and hence secondary metabolite content may deplete with time. Hairy roots cultures are generally fairly productive of respective secondary metabolites, e.g., Anisodus luridus, Talinum paniculatum (Javanese ginseng), Silybum marianum,and Withania somnifera [83–86]. There are some exceptions in the case of Hypericum species as they contain hypericin, but hairy roots were devoid of

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Fig. 5 Normal roots (a) and hairy roots (b) in liquid medium. (Photo a by the author, b courtesy Prof. Sumita Jha, Kolkata)

hypericin. However, transformed plants regenerated from hairy roots contain hypericin [80]. Therefore, this unique organized culture system is not only used as a production platform, but also for new compounds, for example, glycoside conjugates of flavonoids in Scutellaria baicalensis and sarpagine alkaloids from Rauwolfia serpentina [87]. Hairy root cultures have been reported in several species such as artemisinin (Artemisia annua), astragalosides (Astragalus membranaceus), acteoside (Rehmannia glutinosa), centellosides (Centella asiatica), resveratrol (Arachis hypogaea), camptothecin (Camptotheca acuminata), vinblastine, vincristine (Catharanthus roseus), and kutkin iridoid glycosides (Picrorhiza kurroa) [88]. From the ongoing account, it is clear that growth and metabolism of organized and unorganized cultures are different, but sometimes they remain same in productivity as in Panax ginseng cell and root cultures [89]. Panax ginseng hairy root cultures are the most extensively studied root cultures for biomass production and developing scale-up technology for hairy roots grown in various types of bioreactors [90]. As mentioned above, metabolomics and genomics tools can provide more insight into genes expressed or affected due to transformation for hairy roots induction using Agrobacterium rhizogenes [91]. However, metabolomics and transcriptome sequence of a limited hairy root cultures have been reported viz., C. roseus, P. ginseng, A. membranaceus, R. glutinosa, C. asiatica, etc. [88]. In 2011, a consortium of US research organizations started Transcriptome Characterization, Sequencing, and Assembly of Medicinal Plants. The database [92] contains transcriptome data related to 31 medicinal plant species such as Campthotheca acuminata, Cephalotaxus harringtonia, Cinchona pubescens (the quinine tree), Colchicum autumnale, Coleus forskohli, Datura stramonium, Galanthus spp. Gloriosa superba, Papaver somniferum, Podophyllum spp., Taxus x media, and Tripterygium regelii [93]. A new database (www.phytometasyn.ca) contains information about additional 20 medicinal plants including California poppy (Eschscholzia californica) [94]. Therefore, all types of organized and unorganized cultures are used for the production of secondary metabolites. The production of

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secondary metabolites and use of a differentiating system depend upon the species and chemical nature of the secondary metabolites.

5.4

Bioreactor Culture

Bioreactors of various designs and capacity have been extensively used for growing cell cultures of different species but growth of organized cultures though attempted in many species albeit in a smaller capacity bioreactor. It is necessary to optimize the growth of the organized cultures in shake flasks before the use of any type of bioreactor. Both roots and shoot cultures have been grown in static and liquid cultures. Adventitious shoot culture and micropropagation have become commercial level technology. Biomass of several medicinal plants is generated through tissue culture for downstream processing. Hairy roots are developed and grown in many more plant species as production system for secondary metabolites, e.g., Artemisia annua, Atropa belladonna, Azadirachta indica, Beta vulgaris, Catharanthus roseus, Echinacea purpurea, Glycyrrhiza glabra, Hyoscyamus muticus, Lithospermum erythrorhizon, Panax ginseng, Picrorhiza kurroa, and Tagetes patula ([27, 95] and references therein). Shoot cultures grow well on static medium but in liquid medium (submerged, temporary immersion, bioreactor) pose problems of aeration, hyperhydracity (vitrification), and sometimes low biomass production. The major problem with developing scale-up technology for organized cultures is distribution of nutrients and aeration of cultures. The other issues related to the growth of shoots/root cultures in large bioreactor are high shear pressure, providing light to the cultures, and sealing and stability of shaft [27, 95]. However, in spite of these difficulties, shoot cultures have been attempted in Ananas comosus (10 L airlift bioreactor), Hypericum perforatum (2 L stirred tank bioreactor) Lavandula officinalis (5 L bubble column bioreactor), and several others ([96] and references therein). These problems of large-scale organized/unorganized cultures are discussed in detail elsewhere [27, 88, 95] and in some individual chapters in this book.

6

Conclusions

During the last five decades, plant tissue and cell cultures have become technology from a mere technique of growing tissues aseptically, and various branches developed to a commercially viable industrial level production system such as micropropagation, production of useful metabolites, and a platform to produce new proteins. Several high value primary and secondary metabolites are produced through cell culture technology in spite of unresolved constraints. There is no triggering mechanism for the production of secondary metabolites from cells and their release into the medium so that production can be achieved without sacrifice of the productive cultures. With the development and availability of new tools and techniques, focus has been shifted to study metabolomics and genomics. However, efforts are continuous to unravel the biosynthetic pathways, isolating and

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transferring the genes involved in biosynthesis and knowing the gene expression by metabolomics and genomics [82]. The ground has become set for combining different approaches to unravel the mystery surrounded around triggering and development of high level of production system. Acknowledgments Author is thankful to Prof S. Jha, Calcutta University, Kolkata, for providing photographs of hairy root cultures.

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82. Scossa F, Benina M, Alseekh S et al (2018) The integration of metabolomics and nextgeneration sequencing data to elucidate the pathways of natural product metabolism in medicinal plants. Planta Med. https://doi.org/10.1055/a-0630-1899 83. Sivanandhan G, Kapil Dev G, Jeyaraj M et al (2013) Increased production of withanolide A, withanone, and withaferin A in hairy root cultures of Withania somnifera (L.) Dunal elicited with methyl jasmonate and salicylic acid. Plant Cell Tissue Organ Cult 114(1):121–129 84. Qin B, Ma L, Wang Y, Chen M, Lan X, Wu N, Liao Z (2014) Effects of acetylsalicylic acid and UV-B on gene expression and tropane alkaloid biosynthesis in hairy root cultures of Anisodus luridus. Plant Cell Tissue Organ Cult 117(3):483–490 85. Gabr AMM, Ghareeb H, El Shabrawi HM, Smetanska I, Bekheet SA (2016) Enhancement of silymarin and phenolic compound accumulation in tissue culture of Milk thistle using elicitor feeding and hairy root cultures. J Genet Eng Biotechnol 14(2):327–333 86. Faisal A, Sari AV (2019) Enhancement of saponin accumulation in adventitious root culture of Javanese ginseng (Talinum paniculatum Gaertn.) through methyl jasmonate and salicylic acid elicitation. Afr J Biotechnol 18(6):130–135. https://doi.org/10.5897/AJB2018.16736 87. Sheludko Y, Gerasymenko I (2013) Biosynthetic potential of hairy roots for production of new natural products. In: Chandra S et al (eds) Biotechnology for medicinal plants. Springer, Berlin/ Heidelberg. https://doi.org/10.1007/978-3-642-29974-2_10 88. Ganjewala D, Kaur G, Verma PC (2018) An update on transcriptome sequencing of hairy root cultures of medicinally important plants. In: Srivastava V, Mehrotra S, Mishra S (eds) Hairy roots. Springer, Singapore, pp 295–310 89. Le KC, Jeong CS, Lee H et al (2019) Ginsenoside accumulation profiles in long- and short-term cell suspension and adventitious root cultures in Panax ginseng. Hortic Environ Biotechnol 60(1):125–134 90. Jeong GT, Park DH (2017) Mass production of transformed hairy root for secondary metabolites: a case study of Panax ginseng hairy roots. In: Malik S (ed) Production of plant derived natural compounds through hairy root culture. Springer, Cham, pp 183–201 91. Chahel AA, Zeng S, Yousaf Z et al (2019) Plant-specific transcription factor LrTCP4 enhances secondary metabolite biosynthesis in Lycium ruthenicum hairy roots. Plant Cell Tissue Organ Cult 136:323. https://doi.org/10.1007/s11240-018-1518-2 92. Transcriptome Characterization, Sequencing, and Assembly of Medicinal Plants Relevant to Human Health. https://apps.pharmacy.uic.edu/depts/pcrps/MedTranscriptomePlants/. Accessed 6 Apr 2019 93. Lau W, Sattely ES (2015) Six enzymes from mayapple that complete the biosynthetic pathway to the etoposide aglycone. Science 349:1224–1228 94. Xiao M, Zhang Y, Chen X et al (2013) Transcriptome analysis based on next-generation sequencing of non-model plants producing specialized metabolites of biotechnological interest. J Biotechnol 166:122–134 95. Mehrotra S, Mishra S, Srivastva V (2018) Bioreactor technology for hairy roots cultivation. In: Pavlov A, Bley T (eds) Bioprocessing of plant in vitro systems. Reference series in phytochemistry. Springer, Cham. https://doi.org/10.1007/978-3-319-32004-5_10-1 96. Ozlem Y-C, Aynur G, Fazilet V-S (2010) Large scale cultivation of plant cell and tissue culture in bioreactors. Transworld Research Network,37/661 (2),Trivandrum-695 023 Kerala, India,1–54

Part I Cell and Tissue Differentiation and Secondary Metabolites

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Glandular Trichomes on the Leaves of Nicotiana tabacum: Morphology, Developmental Ultrastructure, and Secondary Metabolites Branka Uzelac, Dragana Stojičić, and Snežana Budimir

Contents 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 General Aspects of Plant Trichomes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Leaf Glandular Trichomes of Nicotiana tabacum . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1 Types, Functions, and Distribution of N. tabacum Leaf Glandular Trichomes . . . . . . . 3.2 Morphology of N. tabacum Glandular Trichomes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.3 Development of N. tabacum Glandular Trichomes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.4 Ultrastructure of the Secretory Process in Tall Glandular Trichomes . . . . . . . . . . . . . . . . . 3.5 Cell Compartments Involved in the Process of Secretion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.6 Ultrastructure of the Secretory Process in Short Glandular Trichomes . . . . . . . . . . . . . . . 4 Secretion of Secondary Metabolites of Nicotiana tabacum . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1 Secondary Metabolites of N. tabacum . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2 Histochemical Characterization of Secretions of N. tabacum Tall Glandular Trichomes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.3 Secretion of Secondary Metabolites In Vitro . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5 Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Abstract

Glandular trichomes found on the surface of many higher plants contain specialized cells that produce and secrete copious amounts of particular secretory B. Uzelac (*) · S. Budimir Department of Plant Physiology, Institute for Biological Research “Siniša Stanković” - National Institute of Republic of Serbia, University of Belgrade, Belgrade, Serbia e-mail: [email protected]; [email protected] D. Stojičić Department of Biology and Ecology, Faculty of Sciences and Mathematics, University of Niš, Niš, Serbia e-mail: [email protected] © Springer Nature Switzerland AG 2021 K. G. Ramawat et al. (eds.), Plant Cell and Tissue Differentiation and Secondary Metabolites, Reference Series in Phytochemistry, https://doi.org/10.1007/978-3-030-30185-9_1

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products. Leaf glandular trichomes of the non-model plant species Nicotiana tabacum represent a biologically active and stress-responsive tissue that contributes to plant defense response against biotic and abiotic stress and also influences leaf aroma and smoke flavor. Two morphologically different types of tobacco capitate trichomes, long- and short-stalked, with distinct functions, display ultrastructural features that are common to terpene-secreting glands, but only the secretory cells of the tall glandular trichomes are considered to be the site of biosynthesis of certain exudate compounds, including diterpenes and sucrose esters. Ultrastructural and histochemical characterization of tall glandular trichomes is described in an attempt to understand the contribution of these glands to the total secretion produced. Possible roles of distinct cellular compartments involved in the secretory process and secondary metabolite secretion under in vitro conditions are discussed. Keywords

Glandular trichomes · Histochemistry · Morphology · Secretory process · Tobacco · Trichome development · Ultrastructure · In vitro

1

Introduction

Secretion refers to the complex phenomena of separation of secreted substances from the protoplast and their removal either to the plant surface or into internal spaces or their accumulation in some compartment of the cell [1]. The secreted substances are produced by the secretory tissues of diverse structure and topographic position that occur in most vascular plants. Glandular trichomes are most recently evolved secretory structures found on the surface of about 30% of all vascular plants. They vary in their structure, in the chemical composition of the substances they produce and the mode of their secretion, and in their function. Glandular trichomes are the specific sites for the biosynthesis and excretion of secondary metabolites [2–6] and antipathogenic proteins [7] in many plants. Secondary metabolites are low-molecular-weight natural products with a restricted taxonomic distribution that are usually not essential for normal growth, development, or reproduction but play a role in the plant interaction with its environment, ensuring the survival of the organism in its ecosystem. In order to solve ecological problems, plants must often use their chemistry to manipulate and interact with their environment, by producing specialized metabolites whose functions include defense against predation by herbivores and infection by microbial pathogens, UV protection, attraction of pollinators via color and scent, fertility, and signaling [8–10]. The secondary metabolism pathways are restricted to an individual species or genus and might only be activated during particular stages of growth and development, usually with highest levels of specialized metabolite production occurring during the transition from active growth to differentiation [11].

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Glandular Trichomes on the Leaves of Nicotiana tabacum: Morphology. . .

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Glandular trichomes secrete a wide range of secondary metabolites such as terpenoids, acyl sugars, phenylpropanoids, and fatty acid derivatives and thus contribute significantly to plant’s defense against diverse biotic and environmental challenges. Glandular trichomes secreting lipophilic substances occur in many families, e.g., Lamiaceae, Asteraceae, Geraniaceae, Solanaceae, and Cannabinaceae, and generally consist of a group of glandular cells at the apex of a stalk of one or more cells in length. The storage compartment of glandular trichomes is part of the glandular cell, or cells, which are metabolically active. Two major types of glandular trichomes, peltate and capitate, have been discerned. Peltate trichomes have common characteristics in structure and morphology across genera and mostly produce and store biogenic volatile or semi-volatile organic compounds related to plant abiotic or biotic stress responses [4, 12, 13]. There are various types of capitate trichomes, whose primary function is thought to be the repelling of the pests. Capitate trichomes are specialized to produce and store a large amount of diterpenes and a wide array of nonvolatile or poorly volatile compounds such as acyl sugars or defensive proteins that are directly exuded onto trichome surface [7, 14, 15]. The accumulation of glandular trichome exudates at the surface allows their direct contact with insects, microbes, and herbivores, thus providing the first line of defense at the surface of the plant as well as time for the activation of induced defenses [5]. Species of Nicotiana and closely related genera such as Solanum, Petunia, Physalis, and Lycopersicum have long been known to possess both glandular and nonglandular trichomes, but the use that these plants made of their own glandular trichome secretions remained quite unknown for a long time. The common tobacco (Nicotiana tabacum L.) is an allotetraploid species generated by interspecific hybridization between the ancestors of Nicotiana tomentosiformis and Nicotiana sylvestris approximately 200,000 years before present. The evidence that the characteristic aroma of tobacco is traceable primarily to the resinous exudate from glandular trichomes was presented as recently as 1944 [16], when a knowledge of glandular trichomes became of enhanced interest primarily because of their most prized attributes – aromas of oriental tobacco. In tobacco plants, the exudates of glandular trichomes, which seep directly out of the head cells and produce a resin-like liquid on the leaves, constitute as much as 16% of the dry leaf weight [17]. Presently, cultivated tobacco is among the most chemically and biologically studied species in the plant kingdom, with more than 2500 characterized metabolites that are constantly updated by continuing research [18]. Plant cell, tissue, and organ cultures have emerged as potential sources of secondary metabolites that are used as pharmaceuticals, agrochemicals, flavors, fragrances, coloring agents, food additives, and biopesticides. Through the application of various in vitro approaches and strategies, plant cell, tissue, and organ culture techniques permit manipulation of growth and production of the secondary metabolites in the microenvironment of in vitro cultures, independent of geographical or seasonal variation. From the clonal propagation, biomass, and secondary metabolite production in the in vitro cultures of medicinally important plants, the efforts to scale

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up the production have shifted toward optimizing culture conditions for secondary metabolite production through the application of cell line selection, elicitation, precursor feeding, two-phase co-culture, and other approaches, in order to meet increasing industrial demands and conserve natural sources of these molecules [19]. This study aimed at providing data about ontogenetical development and the localization of the main classes of secreted material and to characterize the cellular changes associated with gland development and secretory process in the leaf glandular trichomes of tobacco plants grown in vitro.

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General Aspects of Plant Trichomes

Plant trichomes are specialized unicellular or multicellular structures derived from the epidermal cell layer that cover most surfaces of most plants. Trichome morphology varies greatly, depending on the tissue and the species, which has often been used in plant classification [20]. In Lamiaceae, studies of trichome type, size, and density have shown that they can be variable within genera, within species (especially in size and density), and between different organs of the same plant but that most differences are species specific in nature and can often be of taxonomic significance even at the subspecific level [21]. Since the morphological and mechanical features (size, shape, density, orientation) of trichomes influence many aspects of plant physiology and ecology, their physiological functions are diverse [5]. Trichomes play an important role in mechanical defense against biotic and abiotic stresses, as well as in the chemical defense of plants by producing a variety of secondary metabolites. There are two main types of trichomes that can be discerned: nonglandular trichomes, mainly differing in their morphology, and glandular (secreting) trichomes, which typically differ in the substances that they secrete. Nonglandular trichomes play a role in mechanical defense against biotic and abiotic stresses and are generally not able to produce or secrete phytochemicals. They can serve to increase herbivore resistance by interfering mechanically with herbivore movement and/or feeding [12]. In addition, they affect water loss, leaf temperature, and photosynthesis through increased light reflectance. By increasing surface reflectance, they contribute to reduced interception of solar radiation and enhanced resistance to low water availability and photoinhibition stress [22, 23]. The degree of pubescence developing on the leaves in desert species changes seasonally, causing dramatic changes in leaf spectral characteristics [23], which alters both leaf temperature and photosynthesis, and is affected by air temperature, leaf water potential, and previous history of the apical meristem during the current growing season [22]. The higher density of nonglandular trichomes on the abaxial leaf surface suggests that they prevent water loss by shielding and trapping air over the stomata [20]. The nonglandular trichomes are diverse in morphology, anatomy, and microstructure. They are abundant in the early phases of the ontogenetic cycle, but their density decreases with leaf maturity [12, 21, 24, 25].

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Glandular Trichomes on the Leaves of Nicotiana tabacum: Morphology. . .

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The principal function of the glandular trichome type is the production of pest- or pollinator-interactive phytochemicals that are stored or volatilized at the plant surface and may also contribute significantly to defense against diverse environmental challenges. Glandular trichomes contain cells that are highly specialized for the biosynthesis and secretion of copious amounts of particular secretory products, such as nectar, mucilage, acyl lipids, digestive enzymes, or protective secondary metabolites [26]. Glandular trichome exudates, which in certain plants reach up to 10–30% of the leaf dry weight, usually contain lipophilic components that are not easily stored in large amounts within the cell due to their phytotoxicity and hence must be amassed outside of gland cells [20]. Glandular trichomes are regarded as the most recently evolved secretory structures. They appear to be absent from conifers, except for the juvenile leaves of some Pinus species, and are rare in cycads but are characteristic feature of a vast number of angiosperms [26–28]. Glandular trichomes of a number of plant species have been suggested to have developed phylogenetically from nonglandular trichomes [28]. Both types of trichomes are initiated and develop similarly up to the stage of a threecelled primordium, after which the differences between the two types begin to appear [29]. Glandular trichomes are constant features of many plant species and develop over the aerial vegetative and reproductive plant organs without external stimuli [28]. The morphology of glandular trichomes and the metabolites they produce vary greatly across plant species. From a functional point of view, based on the mode and timing of secretion, glandular trichomes may be classified into two types: short-term glandular trichomes, which start and end their secretion rapidly, and long-term glandular trichomes, which gradually accumulate their secretion in a subcuticular space [30]. Glandular trichomes secreting lipophilic substances occur in many families, e.g., Lamiaceae, Asteraceae, Geraniaceae, Solanaceae, and Cannabinaceae. Two major types of glandular trichomes are distinguished, peltate and capitate. Peltate trichomes comprise a basal cell, a short stalk cell, and a broad head consisting of many secretory cells arranged in one layer. Their basic morphology does not vary considerably, but they can substantially differ in their sizes [4, 12, 13, 31–34]. Capitate trichomes consist of a basal cell, a one- to several-celled stalk, and a head of usually one or two cells. There are various types of capitate trichomes, which differ significantly in both structure and size [24, 35–38]. Peltate glandular trichomes mostly produce and store biogenic volatile or semivolatile organic compounds related to plant abiotic or biotic stress-induced responses [5, 12]. Glandular trichomes that produce volatile compounds have extracellular storage spaces in which volatiles are secreted and stored [4, 38, 39]. Capitate trichomes typically produce a wide array of nonvolatile or poorly volatile compounds, such as diterpenoids or acyl sugars, which are directly exuded onto the surface of the trichome tip [3, 14, 15, 40, 41]. Their primary function is thought to be the repelling of the pests [40]. Both peltate and capitate glandular trichomes play a critical role in reducing leaf ozone uptake across a number of species and constitute a major factor in reducing ozone toxicity, functioning as a chemical barrier that neutralizes the ozone before it enters the leaf [42].

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3

Leaf Glandular Trichomes of Nicotiana tabacum

3.1

Types, Functions, and Distribution of N. tabacum Leaf Glandular Trichomes

Leaf trichomes of the non-model plant species N. tabacum contribute to plant defense response against biotic and abiotic stress and also influence leaf aroma and smoke flavor. Three morphologically different types of trichomes, covering entire leaf surface throughout development, are found on tobacco leaves: very long nonglandular trichomes and two types of capitate glandular trichomes. Tobacco leaf glandular trichomes represent a biologically active and stress-responsive tissue, as evidenced by gene expression analysis [43]. Tobacco leaf glandular trichomes were classified by Tanaka [44] into two groups: tall glandular trichomes with spindle-shaped glands and short glandular trichomes with very short stalk and spherical multicellular glands. Capitate trichomes differ in their morphological characteristics in the way that reflects different secretory processes within trichomes and probably distinct functions [25]. In Nicotiana species, one of the two capitate trichomes secretes resinous material, while the other secretes aqueous droplets in which nicotine is dissolved [45]. Tall N. tabacum glandular trichomes are distinguished by their large size, high density, and superior secretion ability. Besides providing the physiological consequences of a substantial protuberance, tall glandular trichomes can produce exudate amounting to 17% of leaf dry weight [46], as well as accumulate large calcium oxalate crystals in their gland cells [5]. They secrete sticky clear resinous material, which has been shown to contain cembratrienediol [47]. Meyberg et al. [45] compared fine trichome structure in three different Nicotiana species, in the light of diterpene secretion, and suggested that diterpenes found in the leaf surface gum of N. tabacum and N. sylvestris derive from the tall glandular trichomes but not from short ones. This is further supported by the fact that the leaves of Nicotiana rustica, which bear considerably fewer tall glandular trichomes per surface area but have equally numerous short glandular trichomes in comparison with N. tabacum and N. sylvestris, do not contain detectable quantities of diterpenes within their surface gum [47]. Proteomic analysis of tobacco leaf trichomes identified 1373 protein spots in total, out of which 680 were unique identifications [48]. Using LC MALDI-MS/ MS, 858 proteins were identified, many of which were enzymes involved in secondary metabolism (including enzymes involved in the synthesis of terpenoid precursors and in acyl sugar production) and some involved in secondary metabolite transport. This study also identified various (a)biotic stress response proteins, supporting the role of trichomes in plant defense [48]. Recently, Jud et al. [49] showed that various diterpenoids exuded by the glandular trichomes of N. tabacum act as an efficient chemical protection shield against stomatal ozone uptake by reacting with and thereby depleting ozone at the leaf surface. Short glandular trichomes are numerous on tobacco leaves, and their function is still uncertain. They do not exhibit resinous material on their surfaces. The absence of lipophilic secretion around the short glandular trichomes suggested that this

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Glandular Trichomes on the Leaves of Nicotiana tabacum: Morphology. . .

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trichome type does not actively contribute terpenoid components to the gum. Instead, they are assumed to function as hydathodes (stalked or trichome hydathodes), extruding aqueous droplets under conditions of high atmospheric humidity [45]. Short glandular trichomes additionally secrete nicotine, which is dissolved in the water droplets, most probably by an eccrine mechanism of secretion, since short glandular trichomes do not have markedly developed Golgi apparatus; hence granulocrine water secretion was not detected in the short glandular trichomes [45]. Short glandular trichomes appear to have a more direct connection to the vascular system, as evidenced from studies in Arabidopsis, where intense production of auxin in developing leaf hydathodes was correlated with differentiation of midveins and secondary vascular bundles [50]. Choi et al. [51] suggested a new function of short tobacco glandular trichomes as the apparatus for heavy metal excretion through crystallization. Nonchlorophyllous cells of short glandular trichomes multicellular glands were shown to accumulate within and excrete to the gland surface toxic metals, including Cd and Zn that form Ca-crystal precipitates [51, 52]. Moreover, Shepherd et al. [7] demonstrated a unique biosynthetic capability of tobacco short glandular trichomes, regarding them as specialized biosynthetic structures akin to tall glandular trichomes. The authors provided evidence for the synthesis of tobacco leaf surface proteins, collectively termed T-phylloplanins, in short glandular trichome head cells, and postulated that produced T-phylloplanins are secreted to the gland extracellular spaces and then transferred outside of the glands through constrictions at the termini of intracellular spaces, which are presumed to be secretory pores of unknown structure. Tobacco glandular trichomes are not uniformly distributed over the leaf surface. Rather, their distribution appears to follow the pattern of leaf expansion, as described by Avery [53] in studies of growth of tobacco leaves. The density of trichome population tends to be the greatest in areas of least expansion. Bentley and Wolf [54] analyzed 29 varieties and strains of field-grown oriental tobacco and 4 varieties that were grown under glass and showed varietal differences in trichome density. Trichome population of a given leaf becomes fixed by the time the given leaf has attained from one-fifth to one-fourth its mature size, with the number of trichomes on the upper leaf surface being approximately 20–25% greater than on the lower leaf surface. The number of trichomes per leaf in greenhouse-grown plants is strikingly lower than that of plants of the same varieties that were grown in the field. Leaves on plants grown in the field were 20–60% larger, had a greater trichome population per unit area, and may have 70–140% more trichomes per leaf, than those grown under glass. The total trichome population of each leaf on any field-grown plant, regardless of its position on the stem, is quite the same, except for the few of the uppermost leaves. Bentley and Wolf [54] found that in plants grown under glass, cotyledons entirely lacked trichomes, the first true leaf had only a few trichomes that were, for the most part, restricted to the principal veins and leaf margins, whereas a considerably greater number of trichomes occurred on the second leaf than upon the first one. Trichome population tends to increase rapidly and progressively with increase in the leaf position on the stem until approximately the eighth leaf [54]. Throughout the region

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extending from 8th to 28th leaf, the trend of increasing trichome population per leaf continues but is quite gradual, with only slight difference in number of trichomes on leaves nearest each other. A decline in trichome population with further progression along the uppermost region may be related causally to nutrition [54]. On the basis of these differences in hairiness, Bentley and Wolf [54] assumed hypothesized that hairiness is conditioned by two fundamental factors, one hereditary and one environmental, presumably light. Under the hypothesis that there is a correlation between the morphology of glandular trichomes and their chemical composition, Capdesuñer et al. [55] studied the possible connections between the morphological features of the tobacco leaf and the production of six secondary metabolites in different tobacco lines and found the relationship between the morphology and distribution of different types of trichomes and the contrasting profiles (chemical composition of diterpenes and sugar esters as major secondary metabolites) of the leaf exudates, as well as the terpene concentration necessary to achieve antimicrobial and antioxidant activity [55]. High density of trichomes was related to high concentrations of terpenes in foliar exudates. A greater number of large and branched tall glandular trichomes in most of the accessions analyzed were related to large amounts of cembratrienediols and more complex diterpene profiles. No direct relationship was found between a greater number of short glandular trichomes on the upper leaf side and the presence of any specific diterpene.

3.2

Morphology of N. tabacum Glandular Trichomes

Developing tobacco leaves are covered with single uniseriate nonglandular trichomes and two types of glandular trichomes that exhibit a variety of structural complexity [54, 56–58]. Tall glandular trichomes found on leaves of tobacco plants have multicellular stalks atop a single basal cell and unicellular or multicellular glands and can significantly differ in their size and gland morphology (Fig. 1a). The basal cell is conical in shape (Fig. 1b) and differs from surrounding epidermal cells by the presence of a few plastids (Fig. 1c). Basal trichome cell is connected to the epidermis by numerous plasmodesmata, through which active unidirectional transport of molecules occurs: from the basal cell apically into distal trichome cells but not into the subtending epidermal cell [59]. Christensen et al. [59] suggested that the identified symplastic and apoplastic barrier is generated specifically at the epidermis/trichome interface. The stalk length varies considerably, depending on the number of stalk cells and their length. Younger glandular trichomes usually have two to four stalk cells (Fig. 1d), whereas mature ones have at least six stalk cells that are cylindrical in shape and elongated. Gradation in the length of the stalk cells along the trichome axis is evident (Fig. 2a). The shortest one is the terminal stalk cell, atop which the glandular heads are positioned.

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Fig. 1 Leaf indumentum of in vitro grown Nicotiana tabacum. (a) Tall glandular trichomes of different size and gland morphology. (b) Upper view of abaxial epidermis (e) showing conically shaped basal cell (b) of tall glandular trichome, subtending the lowermost stalk cell (s). Note dark blue stained inner walls of stomata (arrowheads) guard cells, surrounding stomatal pore, after staining with Nadi reagent. (c) Side view of abaxial leaf surface, showing tall glandular trichome basal cells (b) differing from surrounding epidermal cells (e) by the presence of a few plastids. Note centrally positioned nucleus and peripherally positioned chloroplasts in the lowermost stalk cell (s). (d) Upper view of adaxial epidermis showing short-stalked (arrows) and young tall (arrowheads) glandular trichomes, after staining with Sudan Red 7B/hematoxylin

Glands can be unicellular, with spherical or bulb-shaped secretory head (Fig. 2b), or multicellular (Fig. 2c–e). Glandular head may become multicellular by the formation of periclinal walls only (Fig. 2c, d) or by the formation of both periclinal and anticlinal cell walls (Fig. 2e). Although tall glandular trichomes are mostly uniseriate, occasionally bifurcate trichomes, having a gland at each tip, are observed (Fig. 2f). Unlike tall glandular trichomes, the short glandular trichomes display rather uniform morphology. They appear to be more abundant on the adaxial side and consist of a short, single-celled stalk atop the basal cell and a round multicellular head (Fig. 2g). Stalk cell is commonly curved so that the short trichomes do not stand erect but at an angle to the leaf surface, with tendency to lean toward the leaf margins.

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Fig. 2 Morphology of glandular trichomes on the leaves of in vitro grown Nicotiana tabacum. (a) Variously sized tall glandular trichomes showing gradation in the length of the stalk cells (s) along the trichome axis. (b) Tip of the tall glandular trichome showing the uppermost stalk cell and a single-celled head, stained bright pink with Ruthenium Red. (c) Tip of the tall glandular trichome with two-celled head, stained with Phloroglucinol. (d) Tip of the tall glandular trichome with threecelled head. Note numerous bright green chloroplasts in the glandular cells. (e) Tip of the tall glandular trichome with four-celled head, where two uppermost head cells are divided by periclinal wall and two bottom glandular cells arise after an anticlinal division of the lowermost head cell. (f) Bifurcate trichome with an unicellular gland at each tip. The cell walls of stalk and head cells, as well as the secretion, are stained dark blue to black with Sudan Black B. (g) Short-stalked glandular trichome with multicellular head, stained with Nadi reagent. Note curvature within the stalk cell. b basal cell, s stalk cell, h glandular head

3.3

Development of N. tabacum Glandular Trichomes

Trichome production and maturation is limited to short periods early in leaf development [60]. In developing leaf primordia, protodermal cells destined to become trichomes are the first cells to differentiate and cease to divide, while surrounding cells continue normal division [61]. Genetic dissection of trichome development in Arabidopsis thaliana, a genetic model system for the analysis of pattern formation and cell differentiation, enabled us to discern several distinct, genetically controlled steps, from trichome initiation to maturation [62]. Trichomes are initiated in a field of dividing epidermal cells, with a characteristic spacing pattern, based on a mechanism where initially equivalent epidermal cells compete with each other via cell-cell interactions and where networks of transcription factors appear to act together as activators or inhibitors of trichome cell fate [63].

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Fig. 3 Developing glandular trichomes on leaf primordia of different ages, seen in longitudinal sections (a) and transverse sections (b) of Nicotiana tabacum shoot apices

Based on the ultrastructural changes observed, three stages in glandular trichome development can be distinguished: presecretory, secretory, and post-secretory [4, 24 ]. The presecretory stage starts with trichome inception from a single protodermal cell and comprises all developmental phases during which cell divisions and growth occur, until a fully developed trichome reaches its characteristic structure. The surface of a young, developing tobacco leaf is covered with glandular trichomes at different developmental stages, where young trichomes at early developmental stages are found next to mature ones, suggesting a greatly asynchronous trichome development. Glandular trichomes are initiated very early in tobacco leaf development, where they arise from a single expanding protodermal cell undergoing a periclinal division. These differentiating protodermal cells that form protuberances prior to cell division, as well as the resulting two-celled structures, are present already on the first pair of leaf primordia (measured ca. 0.2–0.8 mm in length, Fig. 3). Short and tall glandular trichomes of tobacco cannot be easily distinguished at their inception, both being first discernible as protruding protodermal cells with an asymmetrical cytoplasmic distribution (Fig. 4a). The trichome initial has a large nucleus with prominent nucleolus, dense cytoplasm, and small vacuoles (Fig. 4b). After enlarging and extending above from the leaf surface, the expanded protodermal cell is partitioned by a periclinal, asymmetrical cell division, resulting in the two-celled stage trichome comprised of an apical, more meristematic cell and a basal cell (Fig. 4c–e). Slightly older trichomes are further partitioned by periclinal cell divisions, separating an apical initial and a stalk cell, atop a vacuolated basal cell (Fig. 4f, g).

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Fig. 4 Glandular trichome initiation on leaves of Nicotiana tabacum grown in vitro. (a) Protruding protodermal cell (arrow) on a juvenile leaf. (b) The trichome initial with an asymmetrical cytoplasmic distribution. (c) Early two-celled stage trichome, after the first periclinal division. (d, e) Glandular trichome initials consisting of a single cytoplasmically dense apical cell with large nucleus and a single, more vacuolated basal cell. (f, g) Three-celled stage trichomes, formed after an asymmetrical periclinal division of apical cell. By the appearance of the newly formed daughter cells, it may be possible at this stage to discern between developing short-stalked trichome (f) and tall glandular trichome (g), in which large daughter cell will undergo several rounds of cell divisions to form a multicellular stalk

The apical initial of the short glandular trichome undergoes periclinal cell division (Fig. 5a). Stalk cell of the short glandular trichome elongates without undergoing further divisions (Fig. 5b), whereas that of the tall glandular trichomes divides periclinally (Fig. 5c). Following sequential periclinal and anticlinal cell divisions of the apical cell, the head of the short glandular trichome comprises a number of partially vacuolated glandular cells (Fig. 6). Fully developed short glandular trichome at completion of cell divisions consists of one basal cell, one elongated stalk cell, and multicellular head. A gradient of maturation can be observed for short glandular trichomes along the leaf blade mediolateral axis, from younger trichomes close to the midrib, to fully developed trichomes, toward the leaf margin. In tall glandular trichomes, the apical initial may remain unicellular or give rise to multicellular head as a result of periclinal (Fig. 7) or rarely anticlinal cell divisions

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Fig. 5 Early development of glandular trichomes in Nicotiana tabacum grown in vitro. (a) The late-telophase apical initial (arrow) of the short glandular trichome undergoing periclinal cell division. (b) Young short glandular trichome, with partially vacuolated glandular head cells (arrowhead) and an elongated stalk (arrow) that remains unicellular. (c) Young tall glandular trichome after the periclinal division of the stalk cell, subtending an unicellular apical initial (asterisk)

(Fig. 2e). Periclinal divisions of the stalk cell eventually give rise to a multicellular stalk of variable length, made up of highly vacuolated cells with centrally positioned nuclei. Both short and tall glandular trichomes develop rapidly on leaf primordia, and fully developed forms can be observed already on young, not fully differentiated tobacco leaves [58, 64].

3.4

Ultrastructure of the Secretory Process in Tall Glandular Trichomes

Head cells of the tall glandular trichomes at the presecretory stage ultrastructurally resemble meristematic cells, having large nuclei with prominent nucleoli in a dense cytoplasm with little vacuolation. Occasional mitochondria and a few short cisternae

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Fig. 6 Sequential stages in short glandular trichome development. (a) Anticlinal and periclinal cell divisions in the glandular head. (b) Short glandular trichome at completion of cell divisions, comprising one basal cell (b), an elongated stalk cell (s), and glandular head (h) Fig. 7 Tips of tall glandular trichomes at later developmental stages, with multicellular heads resulting from sequential periclinal cell divisions of the apical cell. Note densely stained secretory cells (sc) containing numerous small vacuoles and highly vacuolated terminal stalk cells (ts) with centrally positioned nuclei

of endoplasmic reticulum (ER) are also observed (Fig. 8a). The plastids are small and contain scarce tubular membrane elements, a few small starch grains, and large globular electron-dense intraplastidial bodies bounded by a single membrane (Fig. 8b). The observed chloroplast inclusions could be thylakoid bodies involved in the formation of the lamellae in developing chloroplasts [65]. Similar electron-

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Fig. 8 Transmission electron micrographs of tall glandular trichomes of in vitro grown Nicotiana tabacum. (a) Trichome head cell in the presecretory stage. (b) Higher magnification of presecretorystage head cell showing plastids with poorly developed lamellar system and large electron-dense globular inclusions (asterisks). (c–g) Trichome head cells in the secretory stage. (c) Elaborate network of endoplasmic reticulum, mitochondria, and large globular electron-dense inclusions dispersed in the cytoplasm. (d) Chloroplast with well-developed lamellar system and a few small starch grains (asterisk), in close association with endoplasmic reticulum (arrow). (e) Extremely osmiophilic granular content within vacuole. Note short cytoplasmic protrusion (arrow) into the vacuolar space. (f) Membranous vesicles filled with the cytoplasmic matrix at the periphery of the vacuole and in the periplasmic space (asterisk). (g) Electron-dense fibrous material released from the outer side of the cuticle. N nucleus, Nu nucleolus, P plastid, ER endoplasmic reticulum, M mitochondrion, V vacuole, C chloroplast, CW cell wall, Cu cuticle. Scale bars: 2 μm (a, c), 1 μm (b, d), 0.5 μm (f), 0.2 μm (g), 0.1 μm (e)

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dense inclusions found in certain chloroplasts in the head cells of Xanthi tobacco were suggested to be a deposit of insoluble lipid materials [56]. Such densely stained globular inclusions were also observed in some of the chloroplasts in trichome head cells of tobacco cultivar Tobacco Introduction (T.I.) 1068, but there was no evidence that these inclusions were related to the secretory product [57]. At the secretory stage, characterized by the marked proliferation of the endomembrane system, head cells appear to be more vacuolated (Fig. 8c–f). Numerous mitochondria, spherical or ellipsoidal in shape and with many well-developed cristae, are present in the cytoplasm that appears less electron dense (Fig. 8c). The chloroplasts of tobacco tall glandular trichome secretory cells, which sometimes contained small starch grains, have well-developed internal membrane system with numerous grana (Fig. 8d). The dominant component of the secretory cells at this stage is elaborate network of smooth ER composed of tubular and cisternal elements that are dispersed throughout the cytoplasm (Fig. 8c). In some regions, short cisternae of rough ER are present, in addition to ER profiles surrounding mitochondria and plastids (Fig. 8c, e). Cisternae of ER were also found appressed to the plasma membrane (Fig. 8g). Vacuoles of various sizes contain osmiophilic material occurring as net-like deposits and globular masses (Fig. 8d, e) and occasional membrane whorls (Fig. 8f). Vesicular structures and suspended polymorphous bodies are occasionally observed in the periplasmic space (Fig. 8f). The membranous vesicles of variable sizes are mostly round to ovoid in shape, but some appear to have polygonal contours, and their surface appears rough, resembling coated vesicles. Smaller vesicles are more tightly packed, and their content appears more electron dense (Fig. 8f). Membranous vesicles present in the periplasmic space greatly resemble secretory vesicles associated with abundant electron-dense secretions of secretory cells [4, 13, 37]. From the outer side of the cuticle, which is less electron dense than the inner part, osmiophilic fibrous material is released (Fig. 8g).

3.5

Cell Compartments Involved in the Process of Secretion

The cells involved with the process of secretion are usually characterized by dense cytoplasm containing numerous mitochondria and small vacuoles, while the frequency of other cell compartments and organelles varies according to the particular substance secreted [28]. The most common ultrastructural feature of cells secreting hydrophobic essential oils and resins is the presence of extensively developed smooth endoplasmic reticulum, much of which is spatially associated with plastids containing osmiophilic material. In addition to extended endoplasmic reticulum, ultrastructural features that are common in terpene-secreting gland cells are leucoplasts with poorly defined internal membranes, or normal or sometimes unusually shaped but otherwise normally appearing chloroplasts, an association of endoplasmic reticulum and plastids, and the relative absence of Golgi [9]. Plastids are, in addition to ER, most often considered to produce lipophilic substances [9, 56, 66]. However, the plastids may differ in degree and type of their differentiation, in the manner by which osmiophilic material accumulates in them,

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and in the manner by which it is secreted from them, as shown for different types of head cells of Inula viscosa L. [67] and Artemisia annua L. [68], suggesting different nature of secreted lipid substances in each cell type. The glandular trichome secretory cells often contain plastids that are nonpigmented, unusually shaped or otherwise different from the chloroplasts of neighboring chlorenchyma tissue. The presence of typical nonpigmented amoeboid leucoplasts devoid of thylakoids and ribosomes correlates strongly with the presence of significant quantities of monoterpenes in the secretion product, but no such correlation was found between the occurrence of leucoplasts in the secretory cells and the presence of sesquiterpene, phenylpropanoid, or aliphatic compounds in the secretion [69]. The secretory cells producing only small amounts or no monoterpenes contain plastids that display various structural characteristics (thylakoids, ribosomes, tubular networks). The glandular cells of some plants (members of Solanaceae, Compositae, Asteraceae) contain functional chloroplasts. Chloroplasts found in glandular trichome secretory cells of different N. tabacum cultivars were well-developed, with normal granal stacking and stromal development [45, 54, 56, 58]. Comparative ultrastructural study of secreting and nonsecreting glandular trichomes of two tobacco genotypes showed that the secreting glandular trichomes of T.I. 1068 had well-developed chloroplasts, while the nonsecreting glandular trichomes of T.I. 1406 lacked chloroplasts and had unusual inclusions in the cytoplasm that were apparently developed from the aggregation of the membranous material [57]. Nielsen et al. [57] hypothesized that the gene responsible for the lack of secreting capability interrupted normal chloroplast development and that chloroplasts are necessary for the synthesis of certain N. tabacum exudate compounds. Although head cells of capitate trichomes often contain chloroplasts without starch grains [56, 68, 70], starch accumulation was observed in plastids of soybean glandular trichome cells [71] and in chloroplasts of tobacco tall glandular trichome head cells [57, 58]. Starch accumulation in chloroplasts indicates that the production exceeds export. Duke and Paul [68] postulated that the glandular trichome chloroplasts are exporting photosynthates at a dramatically higher rate than mesophyll chloroplasts, which is consistent with the view that the chloroplasts of the gland cells are highly involved in converting photosynthates into terpenoids and exporting them to the cytoplasm. The presence of starch grains in the chloroplasts of the trichome head cells was also reported for T. I. 1068 [57], whereas Akers et al. [56] and Meyberg et al. [45] did not observe starch but reported electron-dense globular inclusions in the chloroplasts of the head cells of the tall trichomes. Fully developed chloroplasts in tall glandular trichome head cells of in vitro grown tobacco plantlets contained occasional starch grains and very few small electron-dense globules, whereas large electron-dense globules were observed only in the plastids with poorly developed lamellar system [58]. Although secretory cells of glandular trichomes in most plants lack photosynthetic capability [9, 72], Keene and Wagner [3] described the green appearance of the glandular trichomes of the N. tabacum genotype they studied and showed that the formation of certain diterpenes in their head cells was halted by a photosynthetic inhibitor. Bright green chloroplasts in secreting cells of certain Nicotiana species that are high level accumulators (~5 to 30% dry weight of leaves) suggest a possible

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relationship between their capacity to accumulate exudate and their photosynthetic capacity to fix carbon and/or to produce ATP and NADPH [3, 20]. Conversely to the assumption that the secondary metabolism is often the most predominant metabolic process in trichomes, Cui et al. [73] found tobacco trichomes to be mostly involved in primary metabolic and photosynthetic activities, thus supporting earlier comparative proteomics findings [74]. Genes for photosynthesis and primary metabolism were identified with a high score in trichome EST libraries [43], which is quite different from findings in other plant species, such as Mentha piperita, where photosynthesis-related genes were totally absent. This phenomenon is supported by morphological and structural observations that head cells of tobacco trichomes contain a number of developed chloroplasts, whereas peppermint trichomes do not contain chloroplasts but leucoplasts [4]. The structure of tobacco trichome chloroplasts and the intensity of their fluorescence were shown to be affected by leaf developmental stage [75] and also by environmental factors [76, 77], implying a very complex regulation of their biological activity. Endoplasmic reticulum, in addition to participating in the synthesis of lipophilic substances, may also be involved with intracellular transport of the lipophilic substances from their sites of synthesis to the plasma membrane [1]. Smooth endoplasmic reticulum is often a prominent feature of oil- and resin-secreting glandular cells at the secretory stage [4, 13, 24, 35, 38, 68]. Its abundance makes it possibly an important part of the secretory process. The marked proliferation of the endomembrane system, occurring before the onset of secretion, indicates the beginning of the secretory stage in glandular cell development [24, 35, 70]. In addition to participating in the synthesis of lipophilic material, ER may also take part in intracellular transport of the lipophilic substances from their sites of synthesis to the plasma membrane [1, 28]. Smooth ER is frequently observed in close association with plastids [4, 24, 35, 68], mitochondria [24, 70, 78], or plasma membrane [4, 24, 79]. Turner and Croteau [80] postulated that abundant ER-leucoplast membrane contacts found in peppermint glandular cells could facilitate transfer of limonene to the outer leaflet of ER membranes, while transient contacts between smooth ER and mitochondrial membranes might facilitate isopiperitenol movement. Close contact sites between membranes of different organelles, separated by a narrow layer of aqueous cytoplasm, act in non-vesicular traffic of lipids (the extraction and transfer of individual lipid molecules from a donor bilayer to a target bilayer, usually with the assistance of lipid transfer proteins). Membrane contact sites limit diffusion of trafficked substrates within the confined nanospace, thus reducing loss to the bulk cytoplasm and allowing much higher rates of targeted transfer [81]. Membrane contact sites between smooth endoplasmic reticulum and other organelles are common in plant cells that secrete terpenoids and seem likely to play role in intracellular transfer of terpenoids [26, 81]. To be secreted out of secretory cells, secretory products have to move from the site of biosynthesis through the cytosol, plasma membrane, and hydrophilic cell wall. The translocation of hydrophobic molecules out of the cell requires plasma membrane-localized transporters, such as ATP-binding cassette (ABC) and/or multidrug and toxic compound extrusion (MATE) transporters, to cross the lipophilic

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plasma membrane [82]. In trichomes, ABC transporters were shown to transport diterpenes across the plasma membrane [83]. NtPDR1, a plasma membrane ABC transporter from N. tabacum, is expressed in the tall glandular trichomes of the leaves, stem, and flowers and is involved in diterpene transport [83]. Subsequently, lipid transfer proteins (LTPs), capable of binding small hydrophobic molecules, are likely required to help cross the hydrophilic cell wall and to prevent lipophilic secretory material from repartitioning into the plasma membrane [81]. In tobacco, glandular-specific NtLTP1 protein is required for lipid secretion from glandular trichomes [84]. It is abundantly expressed in tall glandular trichomes but not in short glandular trichomes or epidermal cells [84]. Cuticular rupture has not been observed in capitate trichomes of a number of species. Gland cuticle, with its narrow channels filled with pectocellulosic wall material, is likely to allow the release of some secretion components [24]. Lipids produced by summit cells of I. viscosa trichomes were shown to be secreted through small pores in the cuticle, starting in the form of small drops that gradually grew in size until they coalesced into a drop covering the tip of the trichome [67]. Capitate trichomes of Salvia officinalis that predominantly produce lipophilic secretions possess cutinized unicellular heads with a small pore on its distal end, through which secretory material is presumably exuded [12]. In fully active capitate trichomes of Leonotis leonurus, the glandular head is also covered by a thick cuticle, penetrated by a random fibrillar network continuous with the cellulosic layer of the cell wall [24]. Slight detachment of the cuticle forms a small subcuticular space atop the secretory cells, where the secreted material accumulates temporarily, until its release through cuticular micropores [24, 31]. Subcuticular space has never been detected in either glandular trichome type of N. tabacum [45, 56–58, 64]. Subcuticular space had also never been found in terpene-producing long-stalked trichomes of Cucurbita pepo [36] or short-stalked capitate trichomes of Pogostemon auricularius [85], where the secretion process was indicated by lipid droplets on the apical head cell. The study of Caissard et al. [86] demonstrated that labdane diterpene sclareol accumulates in a crystalline epicuticular form on secretory trichomes of clary sage. N. tabacum leaf trichomes exhibit a typical bilayered cuticular membrane, composed of a thin outer cuticle proper and a thicker cuticular layer, where the cuticle proper consists of no more than three to four lamellae [87]. Exudates that escape cuticular containment via pores (striae) can migrate down from the glandular head and flow out onto the epidermal plane, along depressions between cells where fungal spores may collect [5].

3.6

Ultrastructure of the Secretory Process in Short Glandular Trichomes

Head cells of fully developed short glandular trichomes at the early secretory stage display characteristics typical of secretory tissues with high metabolic activity, as found in mature glandular trichomes of a number of species: the abundance of mitochondria in highly organized cytoplasm, large nuclei with prominent nucleoli,

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Fig. 9 Transmission electron micrographs of short glandular trichomes of in vitro grown Nicotiana tabacum. (a) Glandular cells of short trichomes at early secretory stage. (b) Detail of early secretory stage glandular cell, showing presence of extremely osmiophilic globule (asterisk) on the periphery of the vacuole. (c) Formation of extraplasmic space (asterisk), marking the onset of the secretory stage. Note long narrow cisterns of endoplasmic reticulum parallel to the plasma membrane and surrounding nucleus and mitochondria (arrowheads). (d) Advanced vacuolation of secretory stage glandular cells. Note densely stained cytoplasmic protrusions (asterisks) into vacuoles and conspicuous plasmodesmata (arrow). (e) Detail of parietal cytoplasm in secretory stage glandular cells, with long cisterns of endoplasmic reticulum surrounding plastids, and parallel to plasma membrane and nucleus. Note plastids with electron-dense globular inclusions (asterisks), in close proximity to the small vacuole in which osmiophilic material is deposited (arrow). (f) Glandular cells of short glandular trichomes at advanced secretory stage. Note densely stained droplets of secretory product in increased extraplasmic space and spherosomes (asterisks) in the peripheral cytoplasm. (g) Highly vacuolated glandular cells of short trichomes at late secretory stage, showing numerous droplets of secretory product in cytoplasm and extraplasmic space. N nucleus, P plastid, ER endoplasmic reticulum, ES extraplasmic space, M mitochondrion, V vacuole. Scale bars: 5 μm (a, g), 2 μm (b, f), 1 μm (c, d, e)

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and an elaborate network of endoplasmic reticulum (Fig. 9a) [4, 38, 56, 64, 88]. Variously shaped plastids have electron-dense stroma, sparse lamellar development, and occasional starch grains. A number of small vacuoles are present, many of which contain extremely osmiophilic globules at their periphery (Fig. 9a, b). Long, parallel cisterns of endoplasmic reticulum are present in the parietal cytoplasm and around nucleus (Fig. 9b, c). Plasmodesmata occur frequently between glandular head cells. The outer cell walls of the glandular head develop a thin layer of cutin. The onset of the secretory stage is marked by the formation of the extraplasmic space between the cell wall and the plasma membrane, as a result of the plasma membrane withdrawal from the cell wall (Fig. 9c) [56, 64]. Large central vacuole restricts the cytoplasm to the cell periphery, where long cisterns of ER, mitochondria, and plastids are found (Fig. 9d). Plastids often contain electron-dense globular inclusions in less electron opaque stroma and starch grains (Fig. 9e). As active secretion advances, an increase in extraplasmic space occurs, and lipid droplets and spherosomes appear in the peripheral cytoplasm (Fig. 9f). Numerous densely stained droplets of secretory product in the extraplasmic space are the characteristic feature of short glandular trichome secretory cells of tobacco plants. Akers et al. [56] described large accumulations of the secretory product in the extraplasmic space, between the cells or within the space formed by the juncture of the cell walls with the outer cutinized wall. Large accumulations of secretory product were never observed in short glandular trichomes of in vitro grown tobacco plants, where substantial amount of relatively small droplets was found in the extraplasmic and intercellular spaces [64]. Osmiophilic material was also detected in the extraplasmic space and between the cuticle and cell wall proper in short glandular trichome heads of N. sylvestris [45]. The onset of senescence of tobacco short glandular trichome secretory cells occurs concurrently with the accumulation of secretory product (Fig. 9g). Structural modifications that signal the onset of senescence of glandular cells include degradation of cellular organelles and reduction in the cytoplasm density, followed by structural changes of the tonoplast and other cell membranes [64]. The plastids appear swollen as the number of osmiophilic plastoglobules increases, while the cytoplasm appears fragmented and largely disorganized and contains suspended polymorphous vesicular bodies. The formation of membranous vesicles aggregated along the cell periphery is common in senescing glandular trichome cells [71], as well as membrane blebbing and vesiculation [89, 90]. Deterioration of the tonoplast precedes breakdown of the plasma membrane [91]. Secretory cells of the short glandular trichomes do not contain chloroplasts [45, 56, 64 ]. At the early secretory stage, their plastids have a uniformly staining matrix, with little lamellar development. As the secretion progresses, voluminous electrondense globular inclusions appear, suggesting the plastids as the source of secretory product [45, 56, 64]. However, the release of osmiophilic globules from the plastids into the cytoplasm has not been observed [56, 64]. As the synthesis of secretory product decreases, starch accumulation is observed, indicating the resultant synthesis of starch [56]. Proliferation of the endomembrane system marks the transition from the presecretory to the secretory stage, with endoplasmic reticulum often becoming the

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dominant component of glandular cells [24, 85]. In some glands, during the process of secretion, the appearance of ER elements has been reported to change from flat lamellar form present in the parietal cytoplasm to highly reticulated form, composed of tubular and cisternal elements and dispersed throughout the cytoplasm [24, 92]. No such ultrastructural changes during the process of secretion were found in tobacco short glandular trichomes, in which ER remains parallel to the nucleus and the plasma membrane, throughout the secretory stage [56, 64]. The individual elements of ER are often positioned extremely close to the plasma membrane and may form appositional contacts in the absence of membrane fusion [93]. Several lines of evidence suggest that appositional contacts between ER and membrane systems may mediate the rapid transfer of lipid molecules between the two systems [94]. Extremely osmiophilic deposits are found in the vacuoles of the short glandular trichome head cells at the onset of secretion. Similar, extremely osmiophilic granular content in the vacuoles was shown to represent condensed tannins, which are derived from chloroplasts and stored in vacuoles [95, 96]. Osmiophilic deposits also occur in the vacuoles of the peltate trichomes in Zeyheria montana [13] and Lavandula pinnata [92], in which osmiophilic substances are produced in plastids and then transferred from the plastids and rough ER to the vacuoles, where they are further processed. It is therefore presumed that vacuoles cannot produce but only process osmiophilic substances [13, 92]. In tobacco seedlings exposed to toxic levels of cadmium, crystals containing high amounts of cadmium and calcium are formed in amorphous osmiophilic deposits in vacuoles of head cells of both short and tall trichomes [51].

4

Secretion of Secondary Metabolites of Nicotiana tabacum

Plant secondary metabolites are an extremely diverse group of natural products and can be broadly classified into three major classes: terpenes (such as plant volatiles, carotenoids, sterols), phenolics (such as phenolic acids, coumarins, lignans, stilbenes, flavonoids, tannins, and lignins), and nitrogen- or sulfur-containing compounds (such as alkaloids and glucosinolates). Terpenoids and alkaloids are considered to be the major groups of more than 100,000 secondary metabolites that have been isolated from plants [97]. Nicotiana species produce diverse secondary metabolites including alkaloids, phenolic compounds, acyl sugars, and terpenoids. Terpenoids and phenolics are often secreted by glandular trichomes in a species- and cultivar-specific fashion, whereas alkaloids are not common in glandular trichome exudates [5].

4.1

Secondary Metabolites of N. tabacum

Terpenoids are the most common compound group found in glandular trichome exudates [2, 20]. Various classes of tobacco produced isoprenoids include

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tetraterpenes (carotenes), triterpenes, sterols, diterpenes (cembranoids and labdanoids), and sesquiterpenes (capsidiol, phytuberol, and phytuberin) [98]. A large number of tobacco isoprenoids are involved in the typical tobacco aroma, and some tobacco diterpenes were shown to possess plant growth regulator properties [99, 100]. Tobacco sesquiterpenes act as phytoalexins, low-molecular-weight antimicrobial substances, which cannot be detected in healthy plants but are produced and accumulated in a response to microbial infections or other kinds of stress [101]. Two types of diterpenes in exudates produced by glandular trichomes of Nicotiana spp., polycyclic labdanoids and macrocyclic cembranoids, are the major tobacco diterpenes and account for >50% of secretions [40]. Cembranoids and labdanoids are synthesized in the glandular heads of the trichomes on the surface of the leaves and flowers [98]. The results of Heeman et al. [47] indicated that the occurrence of diterpenoids, especially cembranoids, in the genus Nicotiana is restricted to only a few species: diterpenes could only be detected in the leaf surface gums of 2 of the 16 investigated Nicotiana species, other than tobacco (N. tomentosa and N. sylvestris). All the N. tabacum varieties investigated by Heeman et al. [47] contained diterpenoids of the cembranoid type, the main products in each case being α- and β-2,7,11-cembratriene-4,6-diol, but different varieties showed very marked variations in the content of both of these compounds. α- and β-4,8,13-Duvatriene-1,3-diol were isolated from glandular heads of trichomes of tobacco leaves, and it was suggested that only the heads of glandular leaf trichomes are the site of duvatrienediol biosynthesis [3, 14]. Future synthetic biology research is likely to be concerned with the metabolic engineering of cembranoids, given their antibacterial, antitumor, antifungal, and neuroprotective properties in humans and antihuman immunodeficiency virus activity [102]. Alkaloids are nitrogen-containing secondary metabolites, most of which display strong biological activities and presumably function as defense compounds in plants. The biosynthesis of some alkaloids occurs in multiple organelles within a cell and in multiple tissues within a source organ, from which some alkaloids are then translocated to sink organ. Therefore, the transport of alkaloids within the cell and between the cells and organs has been suggested to be important in alkaloid biology [103]. Nicotine and other tobacco alkaloids (nornicotine, anabasine, and minor alkaloids) are distributed in roots, stems, and leaves, where they have been reported to play various functions such as protection against insects and other herbivores, regulatory substances for growth, and detoxification [18]. There have been no reports on de novo synthesis of alkaloids in glandular trichomes [104]. Nicotine, the most abundant alkaloid in roots, stems, and phloem, is biosynthesized in roots and translocated to all aerial parts of the plant including glandular trichomes [105, 106]. Nicotine accumulates in leaf vacuoles at high concentrations and plays an important role in defense against insect pests and herbivores [107]. In the leaf, nicotine is converted to nornicotine, another major tobacco alkaloid, and both are found inside trichomes [105], but neither is appreciably secreted from the plant [104]. In trichomes, nornicotine is converted to N-acylnornicotines, which are rapidly secreted from the trichomes to the leaf surface and are hence found entirely in the trichome exudate but

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not inside the trichomes [105, 106]. In a number of Nicotiana species, nicotine biosynthesis is rapidly induced by herbivore attack, wounding, or exogenous applications of jasmonic acid or its methyl ester [108]. Laue et al. [106] were first to demonstrate that trichome-based defense, i.e., the production of N-acyl-nornicotine alkaloids, is also rapidly induced by wounding and methyl jasmonate. Flavonoids and anthocyanins are universal plant compounds, responsible for the color of flowers, fruits, and sometimes leaves. In higher plants, flavonoids occur in free (aglycones) and bound (glycosides) form and accumulate in cell vacuoles and chromoplasts in many plant species. Flavonoid aglycones are lipophilic and highly soluble in mixed exudates containing terpenoids [109]. Flavonoids are generally present as water-soluble glycosides in cell vacuoles and chromoplasts, but when present in the epicuticular wax on the leaf surface, flavonoids are nonglycosylated and very often O-methylated [110]. Several studies have shown that tobacco trichomes excrete flavonoids, with 3-O-methylquercetin and other methylated quercetin derivatives being the main flavonoid constituents in Nicotiana exudates [111]. In exudates of several species of Nicotiana and Solanum and further members of the Solanaceae, most of the aglycones are widespread flavonols, flavones occur throughout the family, but flavanones are rare [111]. Plant acyl sugars, typically glucose or sucrose esters that often contain branchedchain fatty acids, are nonvolatile metabolites which are produced and stored in glandular trichomes of many Solanaceae, including Solanum, Nicotiana, Datura, and Petunia species [112]. A significant fraction of these compounds are exuded onto the surface of aerial organs, where they exhibit important protective role against insects. In addition to being directly toxic to herbivores, acyl sugars are excellent surfactants and emulsifiers that may easily entrap insects by sticking to arthropod cuticles, thereby immobilizing or suffocating them [5, 40]. These highly viscous lipids constitute a significant proportion of leaf biomass in the Solanaceae, up to 20% of the leaf dry weight in the wild tomato species Solanum pennellii [113], and are present in lesser amounts in different tobacco species. Glandular heads alone have proven sufficient for acyl sugar synthesis in N. tabacum, whereas epidermal strips from which trichomes were removed failed to synthesize them [14]. Kroumova and Wagner [114] demonstrated that detached trichomes of Nicotiana benthamiana were capable of acyl sugar synthesis. Major trichome form in S. pennellii, classified as type IV [115], appears to be largely responsible for acyl sugar secretion and also more productive in this respect than the corresponding trichome type of N. benthamiana, where acyl sugar accumulation was found to increase with leaf age [41]. In both species, the expression of genes involved in the production and breakdown of branched chain amino acids is higher in trichomes than in underlying leaf tissue. The relatively low expression in trichomes of genes associated with photosynthesis suggests that the trichomes are relying to a significant degree on imported carbon for acyl sugar production [41]. Acyl sugars of tobacco tall glandular trichomes are known to aid in solubilization of lipophilic diterpenoids [40]. Sucrose esters and, less markedly, duvatrienediols, especially highly exuded in N. tabacum, were found to migrate from the glandular head down the trichome stalk to the epidermal cells below, apparently according to

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their relative polarity, particularly under high relative humidity, and both were shown to be chemically stable, at least until senescence [40].

4.2

Histochemical Characterization of Secretions of N. tabacum Tall Glandular Trichomes

Histochemical studies allow a quick and inexpensive preliminary identification of the classes of bioactive compounds in tissues and cell compartments and are often deployed to localize in situ the main classes of secondary metabolites present in plant secretions. Most studies of glandular trichomes from the histochemical point of view mainly concerned Lamiaceae species [12, 24, 32, 116–118] but are rare in Solanaceae [58, 119]. Together with morphological and structural investigations, histochemical studies provide data on secretory modes and secretions, which allow a functional role in plant interactions with its abiotic and biotic environment to be postulated for different glandular trichome types. The main classes of metabolites secreted by the tall glandular trichomes of N. tabacum grown in vitro were characterized by the histochemical tests in plant exudates in situ (Fig. 10). The secretions of tall glandular trichomes stained positively for lipophilic compounds, as revealed by Sudan Black B, Sudan IV, Sudan Red 7B/hematoxylin, and OsO4 staining (Fig. 10a–d). The positive reactions in nonspecific tests using Sudan Black B and OsO4 can be attributed to long chain lipids, aliphatic hydrocarbons with ester or ketone functions, triglycerides, steroids, and free fatty acids [120]. Both neutral and acidic lipids were present within the head cells, as demonstrated by Nile Blue A staining (Fig. 10e). Purple droplets observed within head cells after Nadi reaction evidenced the presence of terpenoids, which were also present in the halo around chloroplasts (Fig. 10f). The use of Nadi reagent to demonstrate different terpenoids in plant secretions was first reported for Pinus pinaster resin duct cells [121]. Since then, the presence of terpenoids (resiniferous acids and essential oils) using Nadi reagent was revealed in the trichome glandular cells or in their secretion in a number of plant species belonging to Compositae [122], Lamiaceae [12, 25, 123, 124], Asteraceae [125], Solanaceae [58, 119, 126], Orobanchaceae [127], Cucurbitaceae [36], Bignoniaceae [13], Boraginaceae [128], Plantaginaceae [129], and others. Diterpenes found in the leaf surface gum of N. tabacum are produced and secreted by the tall glandular trichomes but not by the short ones that do not exhibit resinous material on their surfaces [3, 45]. Tall glandular trichomes stained positively for water-insoluble neutral polysaccharides, as revealed by periodic acid-Schiff’s (PAS) reaction (Fig. 10g), and for noncellulosic acidic polysaccharides (mucilage, pectins) as shown by the Ruthenium Red test (Fig. 10h), indicating copious amounts of mucilaginous polysaccharides secreted by the head cells of tall trichomes. A positive reaction after staining with Ruthenium Red, observed also in the secreted material of capitate trichomes in Salvia aurea [116] and S. officinalis [12], is indicative of the massive presence of viscous, adhesive polysaccharides, which represent a good mechanical defense

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Fig. 10 Histochemical characterization of secretory products of tall glandular trichomes on leaves of in vitro grown Nicotiana tabacum. (a) Cell walls of both stalk and head cells and the secretion are stained dark blue to black with Sudan Black B. (b) Lipophilic secretion stained dark red to orange

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against aphids [12]. Droplets of secreted material were seen on the head cells of type I and type II capitate trichomes of S. aurea, but scanning electron microscopy observations did not reveal cuticular rupture, nor pore formation [116]. Since no discharge mechanism was observed in either capitate trichome type, it was hypothesized that the secretory product is released by cuticular exudation and that the material may possibly be discharged on the collapse of the cells. The presence of alkaloids in the secretion and in the glandular cells, as shown by intense yellow-brown color of the head cell and particularly its exudate, was demonstrated after staining with iodine/potassium iodide and Wagner reagent (Fig. 10i, j). Similar results were reported in Artemisia campestris (ssp. maritima), where the occurrence of alkaloids in oleoresin was confirmed after the alkaloid extraction by tartaric acid [122]. The trichome head cells gave positive reaction for phenolic compounds (Fig. 10k). In addition, trichome head cell vacuoles stained positively for tannins (polyphenols), as evidenced by Toluidine Blue O staining (Fig. 10l). No lignification was observed after staining with phloroglucinol-HCl (not shown).

4.3

Secretion of Secondary Metabolites In Vitro

Reports on the histochemical detection of different secondary metabolites carried out in plantlets grown in vitro are scarce [34, 124, 130]. Plants grown in vitro are continuously exposed to a defined microenvironment, providing nearly optimal conditions for plant proliferation, in contrast to marked variations of the growth conditions of field-grown plants. It is therefore not surprising that there have been reports on the relatively higher yield of secondary metabolites in tissue culture plantlets, callus, and cell cultures than in in vivo plants [131]. Several factors ä Fig. 10 (continued) with Sudan IV. (c) Red-to-brownish staining of lipophilic secretory droplets and plastids in the head cells with Sudan Red 7B/hematoxylin. Note red-to-brown droplets in both apical and subapical gland cell. (d) Osmium tetroxide test showing brown-to-black staining of unsaturated lipids in secreted exudate on the outer surface of the head cell, as well as in plastids and droplets within the head cell. (e) Nile Blue A staining used to distinguish neutral lipids (essential oils) from acidic lipids (resiniferous acids), showing the presence of copious amounts of acidic lipids (stained intensely blue) in material dispersed within the head cell and on the nucleus surface, as well as in exudate droplets on the outer cell surface; neutral lipids (stained pink to red) are present in small amounts. (f) Nadi reagent staining for terpenoids, showing purple droplets of various sizes in the gland cell. (g) Bright pink staining of tall glandular trichome with periodic acid-Schiff’s (PAS) reagent for water-insoluble neutral polysaccharides, showing intensely stained secretory cell wall and distal peripheral cell cross walls. (h) Ruthenium Red test for acidic polysaccharides (mucilage, pectins) showing positively stained tall glandular trichome head cell. Note particularly intense staining of the cross cell wall between head cell and terminal stalk cell. (i, j) Tall glandular trichome head cells stained positively for alkaloids (yellow-brown color), using iodine/potassium iodide (i) and Wagner reagent (j). (k) Tall glandular trichome stained with ferric trichloride for phenolic compounds. Note weak green-to-brown staining in the head cell vacuole and more intensely stained secretory droplets which appear to have been exuded through the cuticular layer. (l) Bicellular trichome head cells showing faint blue-greenish color in the vacuolar space, indicative of polyphenols, after staining with Toluidine Blue O

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might contribute to increased secondary metabolite production in vitro, including the addition of plant growth regulators, sucrose, phenylalanine, macronutrients, elicitors, etc. [132–137]. Cell and organ cultures generally require an exogenous supply of plant growth regulators for cell and organ growth and proliferation of biomass and metabolite accumulation. The plant growth regulator type and concentration are crucial factors for the growth and metabolite accumulation [138–140]. Secondary metabolite production can be drastically altered in response to different plant growth regulators in the culture medium [139, 141–144]. Exogenous application of cytokinins has been shown to affect the essential oil production, often generating higher essential oil yield [124, 145, 146]. The availability of carbon, nitrogen, and sulfur resources, having a large impact on the production of specific classes of primary metabolites, consequently affects the levels and composition of secondary metabolites derived from these primary metabolites [97, 147]. Lower levels of nitrogen and a change in the molar ratio of NH4+/NO3 were also shown to increase secondary metabolite production [143]. Attempts to apply in vitro culture techniques for the large-scale production of highly valuable metabolites have been rather limited [148, 149] because highly differentiated glandular trichomes that produce these compounds are seldom obtained in vitro or their differentiation is inhibited by in vitro conditions [150–153]. Furthermore, the specific conditions of in vitro culture may lead to the occurrence of abnormalities in plant morphology, anatomy, and physiology [154]. In general, leaves of in vitro cultured plants have altered anatomy, including reduced width of cuticle and epicuticular wax [155]. Their thin, soft, photosynthetically hypoactive leaves have malfunctioned stomata and lower trichome diversity, number, and density, resulting in excessive transpiration [150, 151, 156]. In addition, the conditions of conventional in vitro culture may accelerate senescence process [157, 158]. These modifications may result from low irradiance, low concentration of CO2 during photoperiod, presence of sucrose in the culture medium, poor aeration, and high relative humidity in the culture vessels. Generally weak staining demonstrated by different histochemical tests could be accounted for by lower production of secondary metabolites observed under in vitro conditions [34, 148, 149]. In vitro conditions with higher relative humidity, lower light levels, and aseptic environments may account for the lower essential oil accumulation and synthesis of different amounts of monoterpenes compared to wild-growing plants [159]. Bakhtiar et al. [149] suggested that a lower quantity of pentacyclic triterpenoids in Thymus persicus plantlets regenerated in vitro compared to the wild-growing mother plants could be attributed to climatic fluctuations that may result in an inconsistent metabolite production. Despite these shortcomings, plant cell, tissue, and organ culture is a valuable method for the multiplication of selected genotypes and chemotypes, since it can assist in overcoming the individual variability derived from genetic and biochemical heterogeneity, which is the major difficulty in the large-scale production of specialized metabolites [160, 161]. In vitro propagation through axillary shoot formation reduces the possibility of the occurrence of abnormalities that exist with other methods [162]. Along with somatic embryogenesis and adventitious shoot induction, micropropagation can efficiently

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serve to propagate clonally plants under controlled conditions, regardless of the season, and to induce quantitative and qualitative modifications in the production of plant secondary metabolites by changing nutrient and hormonal composition of the growth medium or different physical factors [163]. The secondary metabolite production in cell and organ cultures depends on the type of culture employed. For example, nicotine content in tobacco cell suspension cultures ranges from 0.007% to 2.9% dry weight, while callus cultures produced 0.05–3.75% [97]. Miedzybrodzka and Yeoman [164] investigated duvatrienediols accumulation with reference to the roles of culture origin (tobacco shoot cultures originating from seedlings, callus, and cell suspension cultures), cytokinin, trichome type, and environmental effects. Duvatrienediols were not detected in callus or cell suspension cultures and were only detected in seedling-derived shoot cultures that developed trichomes, with the yield comparable to greenhouse-grown material under optimal culture conditions [164]. Shoot cultures of many medicinal and aromatic plants have been shown to accumulate secondary metabolites to a greater extent compared to natural plants [165, 166]. A number of authors have reported on qualiquantitative differences in the volatile profile between wild-growing plants and those grown in vitro [165, 166]. The variation in the composition of the essential oils could be the consequence of different ontological stage, where in vitro cultured plants are by definition considered as juvenile-stage plants [167].

5

Conclusions

Although a number of research groups have been focusing on developmental aspects of glandular trichome biology for several decades, our current understanding of the development of glandular trichomes is still quite fragmentary. Knowledge of the process of gland secretion comes largely through the study of ultrastructural changes associated with the development of secretory cells. The underlying mechanisms concerning transient subcellular peculiarities that are specific to the time of secondary metabolite production and secretion, particularly those involved with the interorganelle trafficking of molecules, need to be further elucidated. Acknowledgments This research was financially supported by the Ministry of Education, Science and Technological Development of the Republic of Serbia, Grant № 173015.

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The Structural Peculiarities of the Leaf Glandular Trichomes: A Review L. E. Muravnik

Contents 1 2 3 4 5 6

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Structure and Distribution of the Glandular Trichomes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Development of the Glandular Trichomes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Functions of the Glandular Trichomes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Chemical Content of Secretion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ultrastructure of the Glandular Trichome Producing Secondary Metabolites . . . . . . . . . . . . . . 6.1 Synthesis and Accumulation of Phenolic Substances . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.2 Synthesis and Accumulation of Monoterpenes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.3 Synthesis and Accumulation of Sesquiterpene Lactones . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.4 Synthesis and Accumulation of Cannabinoids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7 Mechanisms of Secretion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8 Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

64 65 67 69 71 75 78 82 84 86 87 88 89

Abstract

Leaf glandular trichomes are multicellular secretory structures that arise from protodermal cells. They are characterized by a diverse form, localization, density, and composition of secreted substances. Formation of the glandular trichomes is caused by the need to protect plants from various biotic and abiotic factors, including herbivorous insects, fungi, pathogens, extensive light, UVB radiation, or high temperature. The main feature of the glandular trichomes is an ability to synthesize and accumulate the various compounds of primary and secondary metabolism. Morphology, development, ultrastructural characteristics of the cells producing the secondary compounds, content of the secretory products, as well as mechanisms of secretion are discussed in this chapter. L. E. Muravnik (*) Laboratory of Plant Anatomy and Morphology, Komarov Botanical Institute of the Russian Academy of Sciences, St. Petersburg, Russia e-mail: [email protected]; [email protected] © Springer Nature Switzerland AG 2021 K. G. Ramawat et al. (eds.), Plant Cell and Tissue Differentiation and Secondary Metabolites, Reference Series in Phytochemistry, https://doi.org/10.1007/978-3-030-30185-9_3

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Keywords

Leaf · Glandular trichomes · Morphology · Anatomy · Histochemistry · Ultrastructure · Mechanisms of secretion Abbreviations

AAR ADS CHS CS CYP71AV1 ER GAO GAS GPPS IPD L6OH LS PAL PR RER SER STL THC

1

Aldehyde D11(13) reductase Amorpha-4,11-diene synthase Chalcone synthase Cadinene synthase Cytochrome P450 monooxygenase Endoplasmic reticulum Germacrene A oxydase Germacrene A synthases Geranyl diphosphate synthase Isopiperitenol dehydrogenase Limonene-6-hydroxylase Limonene synthase Phenylalanine ammonia-lyase Pulegone reductase Rough endoplasmic reticulum Smooth endoplasmic reticulum Sesquiterpene lactones Tetrahydrocannabinol

Introduction

Glandular trichomes are the secretory structures developed on the surface of all aerial organs, both vegetative and reproductive. They arise from the protodermal cells as a result of series of anticlinal and periclinal divisions [1]. Authors who have studied the secretory structures called them glands [2, 3], hairs [4, 6], glandular trichomes [7, 9], or extrafloral nectaries [10–12]. Glandular trichomes are found on the aerial surface of 20–30% of vascular plant species [9, 13]. Morphology of the glandular trichomes varies from unicellular to multicellular, from uniseriate to multiseriate; trichomes can have an evident head or a head in them is absent; they form a long or short stalk, and also may not have it at all; some sessile trichomes are sunken. There are a lot of types of the glandular trichomes according to their form. The main types among them are peltate, capitate, cylindrical, conical, scaly, and others. Trichome morphology is specific for the plants from different taxa. A monograph by Metcalfe and Chalk [14] gives a detailed review about the diversity of trichomes. Thus, the peltate trichomes with a disk-shaped head and a very short stalk and capitate trichomes with long stalk are common secretory structures in representatives

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of Lamiaceae family [15–20]. The biseriate structure is typical for the glandular trichomes of many Asteraceae plants [21–25]. Multiseriate and uniseriate glandular trichomes are formed on the aerial organs of Saxifragaceae species [26]. The most important feature of the secretory tissues is the ability to synthesize, accumulate, and release the chemical substances of primary and secondary metabolism. Fahn [3] divided all secretory tissues into two groups: (1) tissues eliminating unmodified or only slightly modified substances, which refer to hydathodes, salt glands, and nectaries; and (2) secretory tissues, in which cells are capable to synthesize and secrete hydrophilic and lipophilic compounds; they include mucilage secreting trichomes, protein secreting trichomes, lipid secreting trichomes, colleters, resin ducts, secretory cavities, and laticifers. According to this classification, the glandular trichomes belonging to the second group will be discussed in this chapter. The glandular trichomes of the plants from the different taxa have specific secondary metabolites. Information on the composition of secondary compounds in the glandular structures can be found in some reviews [27, 28]. For example, trichomes of Leonotis leonurus contain diterpenes [29], Helianthus annuus – sesquiterpenes [30], Artemisia – sesquiterpene lactones [31], Solanum tuberosum – phenols [32], Drosera peltata – naphthoquinones [33], Empetrum nigrum – bibenzyls, catechins, and flavanones [34], Nicotiana species – alkaloids [35]. To determine the chemical composition of accumulated substances, methods of histochemistry, immunocytochemistry, and analytical chemistry are usually used. A series of histochemical tests and fluorescent markers reveals the presence of lipids [36, 37], terpenes [38, 39], sesquiterpene lactones [40, 41], flavonoids [23, 42], polyphenols [25, 43], and alkaloids [18, 44]. The activity of the biosynthesis enzymes for secondary metabolites belonging to different classes, as well as the intracellular localization of these enzymes in the glandular trichomes, was demonstrated in studies based on immunofluorescence and immunogold labeling [45– 48]. Technological progress in the field of liquid chromatography combined with ultraviolet/visual light spectroscopy, liquid chromatography combined with mass spectrometry or with nuclear magnetic resonance spectrometry, gas chromatography combined with mass spectrometry, and capillary electrophoresis combined with mass spectroscopy made it possible to identify the chemical composition of individual compounds present in glandular trichomes [37, 49–52]. Functions of the glandular trichomes depend on their structure, organ on which they are situated, timing of activity, and chemical content of secretion. This chapter is dedicated to the leaf trichomes arising in the early stages of the vegetative shoot development and functioning throughout the growing season.

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Structure and Distribution of the Glandular Trichomes

Morphology of the glandular trichomes is very vivid. This diversity is one of the important features used by botanists to construct taxon-specific systems. Intratribal classification of the Lactuceae (Asteraceae) has been the focus of research carried out by Krak and Mráz [53]. According to study of stems and leaves in 135 plant species,

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trichomes were divided into eight types and subtypes based on their morphology and anatomy. On the leaves and sepals of 52 species, representing all sections of the Drosera genus (Droseraceae), 14 different types of glandular hairs were found [54]. In combination with simple morphological features, glandular hairs facilitate the identification of species even in the pharmaceutically important cut crude drug. Usefulness of the trichome characteristics was revealed repeatedly for classification at the level of the tribe, genus, or section [26, 55, 56]. In spite of the numerous taxon-specific features of glandular trichomes synthesizing lipophilic compounds, these glands have some general attributes. Primarily, trichomes are multicellular secretory structures. They may consist of as few as three to five cells (Mentha piperita – [3]; Satureja thymbra – [57]) or account tens cells (Triphyophyllum peltatum – [58]; Nepenthes – [59]). The upper cells of the glandular trichomes called secretory are arranged in one (Origanum dictamnus – [15]; Ocimum basilicum – [17]; Mentha piperita – [19]; Pterocarya rhoifolia – [60]), two (Dionaeae muscipula, Drosophyllum lusitanicum, Brocchinia reducta – [61], Betula nana – [62]), or three layers (Nepenthes khasiana – [61]; Tussilago farfara – [63]). Secretory cells can form a head that, in some species, is sunken in the epidermal tissue (Heliamphora folliculata – [64] or rise above the level of the leaf surface due to a stalk consisting of one to several cells (Leonotis leonurus – [18]; Salvia officinalis – [44]; Sigesbeckia jorullensis [23]; Withania somnifera – [65]). Usually, the secretory cells have a dense cytoplasm. The cells of a stalk include large vacuoles and are considered as reservoir cells [66]. In some families, including Lamiaceae, Droseraceae, Lentibulariaceae, or Bignoniaceae, one or some specialized cells are between the head and stalk. They are the barrier or neck cells. The main feature of the barrier cells is a cutinized lateral cell wall forming an equivalent of a Casparian strip. Owing to the cutinized lateral walls, the backflow of secreted substances through the wall is prevented and only controlled transport of substances through plasmodesmata occurs [3, 16, 61, 67]. In addition, such cutinization contributes to the mechanical support of the multicellular head, which develops after completion of cell divisions [19]. In glandular trichomes that do not have a large disc-shaped head, barrier cells are absent, as it was found in Asteraceae family plants [22, 23, 41]. At the base of the entire multicellular trichome structure, one or more basal cells are located. The glands on the different organs of one plant can vary in a shape. For example, on leaves and flowers of Plectranthus ornatus (Lamiaceae), there are glandular trichomes of five morphological types [68]. First of all, peltate trichomes are confined to the abaxial leaf surface, whereas two types of capitate trichomes as well as digitiform ones are uniformly distributed on both leaf sides. In contrast to the leaves, the calyx and corolla contain unusual conoidal glandular trichomes. On the vegetative and reproductive organs of Salvia officinalis, also belonging to Lamiaceae, the distinct types of glandular hairs (one peltate and four capitate) with different location and modes of secretion were identified and a functional role of each type postulated [44]. The capitate trichomes of the first type had only hydrophilic secretion. In peltate and capitate trichomes of the second type, hydrophilic secretion prevailed. In the remaining types, lipophilic secretion dominated. According to Giuliani and Maleci [69], in many species belonging to subfamily Nepetoidae of Lamiaceae family, the peltate and small capitate hairs are found on the leaves and bracts, while the large

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capitate hairs present on the calyx. Formation of the diverse glandular trichomes on the surface of the different organs was shown also in Asteraceae plants. In Inula helenium, the short glands are situated on all aerial parts, whereas the stalk hairs are present only on leaves and stems [70]; in two Doronicum species, the cylindrical trichomes without a distinct head are met on the leaves, whereas trichomes with obvious head are formed on the peduncle and phyllaries [71]; in Helianthus annuus, the biseriate capitate trichomes are only on surface of the anther tips and leaves, while uniseriate linear trichomes appear on the all organs except anthers [72]. Trichomes of several types can present on the same organ. Two types of the glandular trichomes are situated on the leaves of Salvia sclarea; they function in a different time and by different ways [73]. According to Bosabalidis [57], the leaves of the aromatic plant Satureja thymbra have numerous glandular trichomes of two distinct types: hairs and scales. Morphometrical analysis showed that leaf surface occupied by glandular scales are about 17-fold larger than leaf surface occupied by glandular hairs. Theoretical calculations estimated the essential oil yield from glandular scales as 3.5% per 100 g of dry leaves. Two types of trichomes occur on the leaves of some species of Nicotiana tabacum [74]. Authors suggested that the long trichomes produce and secrete a resin containing diterpene cembratrienediol, whereas the short trichomes probably give off hydrophilic nicotine. Three types of the glandular trichomes are described on the leaves of Salvia blepharophylla [37]. They are characterized by differences in the composition of the secretion. While peltate trichomes contain terpenes and O-dihydroxy phenols, only terpenes are detected in the capitate trichomes of two types. Three types of glandular trichomes are located on the pericarp of four Juglans species (Juglandaceae) [75]. Trichomes of the investigated types differ from one another in dimensions, chemical content of secretion, as well as in the mechanism of secretion. In some species of Arnica (Asteraceae), there are three types of the glandular trichomes on the leaves, peduncle, and phyllaries [76]. In addition to morphological differences, they demonstrate the diverse reactions to histochemical dyes and different mechanisms of secretion. Differences in the distribution and dimensions of the trichomes are between the two sides of leaves and bracts. For example, capitate and peltate trichomes are found on each side of the leaf of Leonotis leonurus; their number is predominant on the abaxial surface [77]. On the stem leaves of three Doronicum species, two types of the glandular trichomes [16] are formed [41]. While in D. pardalianches glandular trichomes are distributed equally on the both sides, in D. orientale trichomes of one type are rare and in D. macrophyllum they are more common on the abaxial surface compared with adaxial one; glandular trichomes of the other type are formed on the abaxial side only.

3

Development of the Glandular Trichomes

The functioning of trichomes is noted at the early stages of shoot development, as well as throughout the growing season. Trichomes are the first differentiating cells of the epidermal surface on leaf primordia [78]. In Ocimum basilicum, glandular hairs not only form in the early stages of development, but have a high rate of their

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differentiation [17]. In Helianthus annuus, within the first 40 h of seed germination, the glandular trichomes emerge on primordia of the first true leaves. During the following 20–30 h, trichomes develop from trichoblasts to fully differentiated hairs [52]. Different developmental stages of the glandular trichomes were observed in the same leaf of Vernonia galamensis because new glands continue to form until leaf expansion is complete [79]. The uniseriate digestive glands of Pinguicula vulgaris (Lentibilariaceae) are initiated from one protodermal cell of the leaf primordia near the shoot apex and mature fully while still enclosed in the bud. The differentiation of the gland proceeds acropetally in a result of two periclinal divisions, after that a series of anticlinal divisions happens in a terminal cell [80]. The terminal cell and its derivatives retain the ultrastructural characteristics of the meristematic cells up to the completion of their divisions. They are poorly vacuolated, have electron dense cytosol, contain many free ribosomes, the endoplasmic reticulum is sparse, and it has the form of short granular cisternae. The same sequences of events from one protodermal cell through two periclinal divisions and series of anticlinal divisions of the terminal cells leading to the formation of a disk-shaped head takes place in Origanum dictamnus [15], Thymus vulgaris [16], and Mentha x piperita [19] belonging to Lamiaceae family. In biseriate glandular trichomes of Artemisia chritmifolia (Asteraceae), the first division is anticlinal and then several periclinal divisions occur. The glandular trichomes are formed in developing leaves up to the ninth pair. Older leaves are covered with fully developed trichomes [81]. Unlike Artemisia, the ontogenetic development of Helichrysum aureonitens starts from the first periclinal division of the initial cell [82]. When the stalked trichomes are formed, anticlinal divisions can occur in terminal cells, giving a multicellular head [23, 63, 83]. Cytoplasm of the head cell is dense and rich with organelles. In trichomes of Asteraceae plants, divisions are not synchronized between two columns of the cells but occur independently in each column. It was shown for Inula viscosa [84], Stevia rebaudiana [85], Sigesbeckia jorullensis [23], Vernonia galamensis [79], Tussilago farfara [63], three species of Doronicum [41], and others. The development of the glandular trichomes can be divided into three phases: pre-secretory, secretory, and post-secretory [86, 87]. In the trichomes of Cannabis sativa and Leonotis leonurus, the pre-secretory stage starts with the differentiation of the trichome from an initial cell and ends when the trichome reaches a fully developed structure. Secretory cells have a dense cytoplasm, large nuclei with prominent nucleoli, and small vacuoles. Occasional Golgi bodies and a few short cisternae of rough endoplasmic reticulum, generally restricted to the cell periphery, are also observed. The plastids contain scarce tubular membrane elements, a few starch grains, and large globular electron-opaque intraplastidial bodies bounded by a single membrane. At the secretory stage, proliferation of the endomembrane system occurs. Endoplasmic reticulum consists of tubular and cisternal elements, which could be seen throughout the cytoplasm. An increase in the number of Golgi bodies happens. The arrangement of the cisternae in the Golgi body frequently shows a clear polarity. The trans-cisternae are associated with numerous vesicles, which

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could be divided into two categories: large vesicles with smooth membrane and small, coated vesicles. The smooth endoplasmic reticulum of the secretory cells forms associations both with the leucoplasts and the plasma membrane; it often contains densely staining material, and may be involved in the transport of the secretory products. Secretory activity has nearly stopped in older trichomes. The number of Golgi bodies, vesicles, cisterna and tubules of endoplasmic reticulum, and mitochondria are reduced. The glandular cells undergo vacuolization, and the cytoplasm becomes very dense and begins to degenerate.

4

Functions of the Glandular Trichomes

Glandular trichomes are formed to protect plants from various biotic and abiotic factors, including herbivorous insects, fungi, pathogens, extensive light, UVB radiation, or high temperature. Plant protection is ensured by both changing of the glandular trichome density and a synthesis of the various chemical agents [9]. Trichome formation is an important component of resistance against herbivorous insects [88]. For example, the perennial herb Arabidopsis lyrata has a genetically based polymorphism in trichome production and occurs in glabrous and trichome-producing forms. It has been shown that glabrous plants are more damaged by herbivorous than trichome-producing plants [89]. Because plants cannot change the density of trichomes on already-existing leaves, an induced increase in trichome production can only be expressed in leaves developing during or subsequent to attack [90]. Usually the magnitude of the increase in trichome density is between 25% and 100%, but in some cases as large as 500– 1000% [91]. In addition to protection against herbivorous insects, the leaf trichomes can protect mesophyll cells from damage caused by UV-radiation [92–94]. Ultraviolet B results in a considerable reduction of photosystem II photochemical efficiency in not pubescent leaves of Quercus ilex [92]. At the same time, methanol extracts of the leaf trichomes show significant absorbance in UV. The trichomes in this plant contain flavonoids including kaempferol glycosides. Some studies suggest that flavonoids in the trichomes may function as the effective filters against the harmful ultraviolet B radiation. The capacity of leaf hairs to protect underlying tissues against UVB was assessed during leaf development in Cydonia oblonga, Eriobotrya japonica, and Olea europaea [93]. In three species, the trichome density and relative quantities of phenolic constituents in the trichomes were considerably higher in young leaves. Moreover, in de-haired leaves of Olea europaea the underlying photosynthetic tissues were more easily damaged by UVB. The same peculiarities were observed in Quercus ilex, Q. coccifera, and Verbascum speciosum during the development of their leaves [94]. Flavonoids, especially flavonoid aglycones, are accumulated in the trichomes of plants from arid or semiarid habitats, where high insolation exists. Trichome frequency is markedly greater in leaves of Phillyrea latifolia exposed to full solar

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radiation than in leaves growing on the shady side [42]. The concentration of leaf phenylpropanoids varies significantly between sun and shade leaves, with a marked increase in sun leaves. In winter leaves of Thymus sibthorpii, glandular hairs are numerous and secrete a higher amount of essential oil, compared to summer leaves. The major components of the essential oils in leaf extracts are linalool and p-cymene. Linalool dominated in the winter leaves and p-cymene in the summer leaves. The high content in linalool is ascribed to the low environmental temperatures. Due to the antioxidant properties of the essential oils, the plant can tolerate low temperatures in winter, which stimulate oxidative stress [95]. Among biologically active substances, which are produced by the glandular trichomes, there are large amounts of primary and secondary compounds. The sticky substance secreted by the trichomes of Inula viscosa contains both lipids and polysaccharides that give the leaf a shiny appearance [84]. Such secretion may be useful in light reflection and thereby in reduction of leaf temperature. The presence of viscous, adhesive substances in secretion of the trichomes in Salvia officinalis offers good mechanical defense. This is demonstrated by the fact that aphids, which invade the plant to feed on sap, stuck in the viscous substances released when they trigger rupture of the cuticle [44]. Various phenolics, monoterpenes, sesquiterpenes, and alkaloids are contained in the glandular trichome secretion. They are involved in chemical defense: terpenoids are olfactory deterrents for insects [39, 87, 96]; flavonoids and tannins are the strongest contact deterrents [97]; bitter sesquiterpene lactones make the plant inedible for herbivorous insects [23, 30], and alkaloids are poisonous [98]. Many of the above-mentioned substances have bactericidal and fungicidal activity [9, 99, 100], as well as possess allelopathy [44]. In those cases, when the glandular trichomes are damaged by insects, the phenolic compounds stored within them can be released. Once released, the phenolic compounds are oxidized to quinines by polyphenol oxidase and “glue” insects to the leaf surface so that they are unable to feed [1, 101]. The antioxidant properties of these compounds also protect plants against bacterial or viral attack by counteracting excessive reactive oxygen production [9]. The glandular trichomes located on the modified leaves, or traps, of carnivorous plants are able to attract, capture, kill, and digest prey, and absorb the resulting mineralized products. The secretory tissues of these plants have multiple roles. For example, the digestive glands of Nepenthes produce not only enzymes but also antibacterial and antifungal substances [102]. Visual and olfactory cues are also among the mechanisms of prey attraction [103]. The heads of the glandular trichomes of many Droseraceae carnivorous plants are colored bright red owing to anthocyanins [61] or naphthoquinones [104]. These pigments act as a visual attractants, so far as many Drosera species and Drosophyllum lusitanicum catch aerial prey. On the other hand, presence of the naphthoquinones is a peculiar defense against microbial organisms during enzymatic degradation of insect biomass on the leaf surface.

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Peristome and lid of the pitchers in Nepenthes species form extrafloral nectaries, in which synthesis of the sweet nectar as well as emission of volatile compounds takes place [103]. The production of aromatic substances in pitcher traps of Heliamphora, Darlingtonia, and Sarracenia species is associated with nectar too [61]. The main sources of honey-like scent in Heliamphora folliculate traps are nectar-spoons at the top of the pitcher [64]. The leaves of few Drosera species produce a lemon-like or sweet honey scent. It turned out, Drosera species that have scented leaves have scentless flowers, whereas species that do not have fragrant leaves have scented flowers [105]. Depending on the mode and timing of secretion, Werker [96] classified the glandular hairs into two types: (a) short-term glandular hairs, which start and end secretion rapidly and serve for the protection of young organs; and (b) long-term glandular hairs, in which secretory materials accumulate gradually under an elevated cuticle and which serve for the protection of mature organs.

5

Chemical Content of Secretion

Glandular trichomes are the site in which diverse lipophilic compounds are synthesized. Moreover, many secondary metabolites accumulate only in the trichomes. A study on Nicotiana tabacum leaves demonstrated that removal of glandular heads from midrib reduced or eliminated duvatrienediol biosynthetic capacity [106]. Isolated glandular trichomes of peppermint synthesize monoterpenes, and secretory cells represent the site of biosynthesis of monoterpenes in plants [107]. Sesquiterpene lactone artemisinin was identified in Artemisia annua in leaf extracts of a biotype containing trichomes, while it was absent in leaf extracts of a mutant biotype lacking trichomes [108]. A number of histochemical procedures are often used to identify and localize the main classes of chemical substances in the trichome cells. As a rule, they are performed on fresh hand sections. The following chemical reagents are applied to localize phenols: vanillin/HCl gives red coloration of tannins [109]; ferric chloride provides dark blue color of polyphenols [110] (Fig. 1a); fast blue gives red to dark brown color of polyphenols [110]; potassium dichromate demonstrates brown coloration of condensed tannins [110]; 4-nitrosophenol/H2SO4 identifies a cherry red color of tannins and catechins [112] (Fig. 1b); and toluidine blue detects turquoise color of polyphenols [111] (Fig. 1c). In addition to histochemical tests, autofluorescence in UV [113] (Fig. 1d) as well as the use of fluorescent markers can help identify phenolic compounds. So, flavonoids are visualized by induction of fluorescence with aluminum chloride [114], Wilson reagent [115], or natural reagent [42] (Fig. 1e). Using a series of histochemical reactions, the researchers repeatedly revealed in glandular trichomes the various phenolic compounds, including polyphenols (in Origanum vulgare [6], Salvia aurea [38], Lippia scaberrima [43]), flavonoids (in Helichrysum aureonitens [82], Plectranthus ornatus [68], Calceolaria adscendens [39], Phillyrea latifolia [42], Sigesbeckia jorullensis [23], Cucurbita pepo [36], Matricaria chamomilla [116] and Santolina ligustica [117]), and tannins (in Salvia officinalis [44] or Pterocarya rhoifolia [60]).

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Fig. 1 Histochemical tests to identify the secondary metabolites in the glandular trichomes. (a) Fresh section of the trichome in Inula helenium stained with ferric chloride for phenols. Phenols present in the upper cells of the trichome, subcuticular cavity (SC) is colorless. (b) Fresh section of the trichome in Empetrum nigrum stained with nitrous acid for phenols, which accumulate in secretory cells as well as in neck cells. (c) Fresh section of the trichome in Tussilago farfara stained with toluidine blue for phenols. They are found in the head cells. (d) Induction of green autofluorescence of phenylpropanoids in UV light on the fresh section of the trichome in I. helenium. Phenylpropanoids are seen in the two upper layers; red autofluorescence of chloroplasts is seen in the lower cells. (e) Yellow–green fluorescence of phenylpropanoids in UV light on the fresh section of the trichome in E. nigrum stained with Natural Product reagent. Fluorescence is seen in vacuoles and the cell wall of head and neck cells; it is absent in other cells of the stalk. (f, g, h) Fresh sections of the trichomes in I. helenium (f), Arnica foliosa (g), and Doronicum orientale (h) stained with NADI reagent for terpenes. Terpenes are localized in the secretory cells and subcuticular cavity. (i, j) Fresh sections of the trichomes in I. helenium (i) and Arnica montana (j) stained with concentrate hydrochloric acid for sesquiterpene lactones. Sesquiterpene lactones are seen in the subcuticular cavity. Scale bar – 25 μm

For visualization of terpenes, NADI test has developed and is most often used [118]. In result of this reaction, terpenes give dark blue or violet staining of cytoplasm and subcuticular cavity in the head cells (Fig. 1f–h). Positive reaction was shown in the leaf trichomes of Artemisia campestris [81], Leonotis leonurus [18], Salvia aurea [38], S. officinalis [44], Plectranthus ornatus [68], Stevia rebaudiana [119], Santolina ligustica [117], Cucurbita pepo [36], Rosa х damascena [120], two Chrysolaena species [25], Rubus idaeus [121], three Doronicum species [41], and many other studied plants. A unique acid-staining technique has been developed to identify sesquiterpene lactones (STL) that dye cells of the glandular trichomes in orange to red [40] (Fig. 1i–j). STL are revealed both in cytoplasm of the cells and in subcuticular cavity above the trichome head. STL is present in the glandular trichomes of many Asteraceae species (in Artemisia nova [31], A. umbelliformis [122], Arnica species [123]), Lamiaceae (in Salvia officinalis [44]), and other families (for instance, Scrophulariaceae – in Calceolaria adscendens [39]).

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A number of histochemical tests were suggested to establish accumulation of alkaloids in glandular cells. They include reactions with Wagner, Dittmar, Ellram, and Dragendorff reagents [124]. If secretory tissues contain alkaloids, Wagner and Dittmar reagents give red brown coloration of the cells, Ellram reagent produces a densely red cellular content, while Dragendorff reagent provides brown color. Based on the histochemical methods, accumulation of alkaloids was established in the glandular trichomes of Cannabis sativa [124], Salvia officinalis [44], and Zeyheria montana [125]. In interpreting histochemical results, caution must be taken since histochemical techniques give information about the chemical class but are not specific for the determination of the concrete compounds. Histochemical tests can be verified using biochemical analytical methods. The last ones include gas chromatography-mass spectrometry, high-pressure liquid chromatography-UV-diode array detection, highperformance liquid chromatography–electrospray ionization-mass spectrometry, high-performance thin-layer chromatography, as well as capillary electrophoresis. Using analytical methods, it was possible to establish the presence of individual compounds in glandular trichomes in many plant species. So, secretory trichomes producing phenolic compounds are found in Cucurbita pepo, Helianthus annuus, Santolina ligustica, Sigesbeckia jorullensis, Stevia rebaudiana, Leonotis leonurus, Origanum vulgare, Plectranthus ornatus, Salvia aurea, S. blepharophylla, Rosmarinus officinalis, Thymus vulgaris, Phillyrea latifolia, Orobanche ramose, Lippia scaberrima, and others belonging to Cucurbitaceae, Asteraceae, Lamiaceae, Oleaceae, Orobanchaceae, and Verbenaceae families. The main classes of phenolics found in the secreted material of the glandular trichomes are flavonoids and tannins [23, 42]. In addition, flavonoid aglycones are identified in the secretion of few species [37, 87]. Phenolic components eugenol, cis-3-hexenyl benzoate, cis-jasmone, transnerolidol, and benzyl alcohol were identified in the essential oil of Salvia blepharophylla by GC–MS analysis [37]. Flavonoids luteolin 7-O-glucoside and quercetin 3-O-rutinoside present exclusively in the secretory products of glandular trichomes of Phillyrea latifolia leaves exposed to high level of light [42]. The identification of phenylpropanoids was made by interfacing the HPLC-DAD with an HP1100 MSDI API and electrospray mass spectrometer operating in the negativeion ionization mode. Analysis of the extracts from leaf glandular trichomes of Lychnophora ericoides showed the presence of five acetylated flavonoids as a result of employing negative ion “chip-based” nanospray tandem mass spectrometry [126]. To unequivocally confirm the identity of the flavonoids found, other glandular trichomes were submitted to the analytical methodology developed for the analysis of the secondary metabolites in crude extracts by HPLC/DAD-MS. This analysis confirmed the availability of chrysin, pinocembrin, pinostrobin, pinobanksin, and 3-O-acetylpinobanksin in the glandular trichomes of L. ericoides. The composition of the secretory products may include the various terpenes, among which monoterpenes, diterpenes, sesquiterpenes, and sesquiterpene lactones are detected. So, GC–MS analysis of volatile compounds emitted by the trichomes

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that are located on the leaves, sepals, and petals of Rosa x damascena demonstrated different content of monoterpenes in each extract [120]. Chemical analysis was performed on R. damascena “Quatre Saisons Blanc Mousseux” and its parent R. damascena “bifera” and on R. centifolia “muscosa” and its parent R. centifolia. Comparison of the volatile compounds emitted by the trichomes allowed the researchers to speculate on the independent occurrence of mutation in two lines of Rosa x damascena. Qualitative and quantitative GC–MS analysis of the essential oil from glandular trichomes of Salvia aurea revealed camphor as the main constituent [38]. Two terpenes, camphor and cineole, are emitted into the surrounding atmosphere and are adsorbed by soil. Authors suggested that these substances led to the inhibition of germination and growth of grassland herbs, by an allelopathic effect. Accumulation of the diterpene duvatrienediol has been observed by HPLC in detached, intact heads of Nicotiana tabacum glandular trichomes [106]. None was found in trichome stalk, epidermal or subepidermal tissue extracts. The essential oils isolated from leaves of Plectranthus madagascariensis contained a diterpene 6,7-dehydroroyleanone [50]. Comparative thin-layer chromatography of the extracts from isolated glandular trichomes of Salvia divinorum and the leaves from which they had been removed revealed the presence of high concentrations of diterpene salvinorin in the glandular trichomes [127]. In Sigesbeckia jorullensis glandular hairs, sesquiterpenes and other terpenes of higher molecular mass are the main components of the secretion. In addition to the 27 sesquiterpenes, 18 compounds of the essential oil were found to be oxygenated diterpenes [23]. A relationship between sesquiterpene lactones and peltate glands has been previously shown in many publications [31, 122, 128]. Synthesis and accumulation of the sesquiterpene lactones are characteristic of many species of Asteraceae [129]. The glandular trichomes often produce mixed secretions, with lipophilic and hydrophilic components simultaneously in the same trichomes, but more often with one predominating over the other. The glandular trichomes of insectivorous plants of Droseraceae refer to such secretory structures. For instance, in Drosophyllum lusitanicum the glands are able to synthesize polysaccharide mucilage and release a very strong honey scent as well [61]. Pectins are present in the secretion of the glandular trichomes of not only carnivorous plants, but also of other species [18, 38, 69, 84, 85, 123, 130]. The digestive and slime glands of D. lusitanicum, tentacles of Drosera, and digestive glands of Dionaea muscipula have red color. Coloration of the glandular trichomes is provided by anthocyanins cyanidinglycoside and pelargonidin-glycoside, whereas volatile substances may be terpenoids and phenylpropenes [131]. Composition of secretion in the subcuticular cavity can be established in several ways: removal of the material from the glands with isolation of the glands from the plant and solvent extraction. To identify the contents of the glandular trichomes and to compare it with the contents of surrounding cells, some researchers use glass capillaries, which can penetrate into the subcuticular cavity and collect the secretion inside it. Voirin [132] used a capillary to examine the oil from heads of Mentha  piperita glands. Authors showed that the free lipophilic flavonoid aglycones together

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with monoterpenes are located in the peltate glands. After using glass microcapillaries and following analyses by GC–MS or HPLC, an accumulation of monoterpenoid nepetalactones was found in the subcuticular cavity of the peltate glandular trichomes in Nepeta racemosa [49], as well as sesquiterpene lactones and flavonoids were detected in the capitate glandular trichomes of various Scalesia species [133].

6

Ultrastructure of the Glandular Trichome Producing Secondary Metabolites

One of the specific structural features of most lipid-secreting glandular trichomes is an ability to form a cavity derived from separating of a cuticle and outer wall of the secretory cells where secretion accumulates. Secretory products are thrown out when a cuticle is broken off. As shown in some studies of Lamiaceae plants, a subcuticular cavity contains usually essential oils, among which there are monoterpenes, sesquiterpenes, as well as phenylpropenes [107, 131, 134]. Subcuticular cavity filled with toxic substances is a first line of defense at the surface of the plant. Two modes of development of subcuticular cavity are described in the literature. According to the first, a large space develops between the cuticle proper and underlying cell wall [125, 135]. The second mode includes enzymatic break down of a middle layer of cell wall rich in pectin and following division of cell wall into two unequal parts. In this case, a thin outermost pectin layer of the wall together with the cuticle proper rises up, while a thick lower part remains pressed to plasma membrane. Ultrastructural details of subcuticular cavity formation are given below. The initial stage is an enzymatic loosening of the fibrillary matrix of the outer cell wall. Additional deposition of cutin globules leads to the formation of the cutinized layer in it (Fig. 2a). The gaps between the fibrils arise as small and elongated electron-translucent areas, often in a tandem arrangement (Fig. 2b), indicating their aggregation and imparting to the outer cell wall the appearance of a loose fibrillary mesh. As a result of the wall loosening and of secretion accumulation, a subcuticular space is formed by the detachment of the cuticle together with outermost pectin layer of cell wall (Fig. 2c). The secretion consists of finely granular hydrophilic material mixed with large electron-light oil droplets and vesicle-like structures of irregular surface (Fig. 2d). The vesicles with an electron-light content are delimited by an electron-dense layer and apparently increased in size by fusion. The dense layer around the vesicles may represent an interphase between lipophilic and hydrophilic compounds. The different density of vesicle contents and emulsionlike appearance of secretion probably reflects the different chemical phases in them [39, 69]. Several studies suggested that the cuticle remains relatively thick during expansion or even becomes thicker with age. Kim [136] assumed that in Cannabis glands, vesicle-like structures within the subcuticular space carry deposits of cutin to the expanding cuticle. No evidence to support either of the complex assumptions was obtained with peppermint glands [19]. Rather, it was shown that during expansion of

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Fig. 2 Ultrastructure of cell wall and formation of subcuticular space in the glandular trichomes, synthesizing the secondary metabolites. (a) Deposition of cutin in the cutinized layer (CCW) of upper cell wall (CW) in the trichome of Doronicum orientale, synthesizing phenols and terpenoids; C cuticle. (b)Appearance of the elongated light areas (LA) in cutinized layer of the cell wall in D. macrophyllum. (c) Increase of the light areas in the number and size and fusion of them one with others in trichomes of D. pardalianches; SC subcuticular cavity. (d) Fully formed subcuticular

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the subcuticular space, the gland cuticle becomes thinner along the lateral sides while the central region of the dome remains thick. Similar processes of subcuticular cavity formation were shown in the glandular trichomes of Chrysanthemum morifolium [21], Inula viscosa [84], Thymus vulgaris [16], Cannabis sativa [8], Artemisia annua [22], Ocimum basilicum [17], Nepeta racemosa [137], Leonotis leonurus [18], Stevia rebaudiana [85], Sigesbeckia jorullensis [23], Vernonia galamensis [79], Pterocarya rhoifolia [60], and Doronicum species [41]. It was concluded that wall reinforcement along the cuticle may give resistance to the secretory cavity when large amounts of secretion are stored. In Asteraceae plants, subcuticular space expansion began with the rising of the cuticle above one of the most terminal cells and continued until the cuticles of the two terminal cells merged into one. In contrast to the large subcuticular cavity, the fundamentally different place for accumulation of secretion is the periplasmic space and loose cell wall. In the glandular trichomes with such a place of storage, the upper cell wall is thick. Under thin layer of cuticle proper, there is a thickened cutinized layer with dense network of dendritic structures and inter-fibrillary lamellae (Fig. 2e). The network of dendrites is oriented perpendicular to the plasma membrane, while inter-fibrillary lamellae resemble a chain of transparent crystals, which lie parallel to it. Kim and Mahlberg [8] named these structures “hyaline areas.” A subcuticular cavity in these glandular trichomes is small or quite absent. For instance, in the capitate trichomes of Leonotis leonurus [87], Betula species [62], Tussilago farfara [63], or in the peltate trichomes of Pterocarya rhoifolia [60], the cell wall has above-mentioned ultrastructural features. Probably, differences between the two modes of accumulation of secretion in the glandular trichomes depend on the dissimilarity of the chemical content of the secretion and are characteristic for the trichomes located on the different organs. So, unlike the peltate glandular trichomes in Leonotis leonurus, which produce lipid secretion, the secretory pull of the capitate trichomes contains predominantly the protein-polysaccharide components. Another example relates to the glandular trichomes of Doronicum species localized on the vegetative and reproductive organs. The presence of polysaccharides, phenols, and terpenoids were found in the leaf trichomes, which are able to form the subcuticular cavity [41]. At the same time, in the glandular trichomes situated on the phyllaries, polysaccharides are found in small quantity; however, secondary metabolites prevail.

ä Fig. 2 (continued) cavity filled with complex secretion in the trichome of A. chamissonis, synthesizing sesquiterpene lactones. (e) The cell wall with a thick cutinized layer in the trichome of the phyllary in D. macrophyllum, in which a subcuticular cavity is absent. (f) Formation of the numerous micro channels (MC) in cutinized layer in the trichome of D. macrophyllum. G Exocytosis of the multivesicular body (MvB) to periplasmic space in the trichome of Empetrum nigrum, synthesizing bibenzyls and chalcones. (h) Content of the periplasmic space (PS) in the anticlinal cell wall in the trichome of E. nigrum. (i) The periplasmic space with osmiophylic globules of secretion (S) in the trichome of Arnica chamissonis. Scale bar – 0.5 μm (a–c, e–i); 5 μm (d)

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These trichomes have a thick outer cell wall, and a subcuticular cavity in them is absent (Fig. 2e, unpublished data). After overcoming the plasma membrane, the secretion is temporarily stored in the periplasmic space and subsequently passes through the wall. In the absence of cuticle rupture, the release of secretion possibly occurs through cuticle micro channels (Fig. 2f) that will be discussed in the section 7. The volatile components also may pass through minute pores of the intact cuticle [138]. In the periplasmic space, there are both hydrophilic and lipophilic components of secretion (Fig. 2g–i). This fact determines the formation of a complex of ultrastructural features that contributes to the synthesis of the primary and secondary metabolites. Betula glandular trichomes form a periplasmic space filled by black secretory material [62]. Small osmiophilic globules arise in a periplasmic space of Inula viscosa [84], Artemisia annua [22], Achillea millefolium [139], or Tussilago farfara [63]. In the following sections, ultrastructural features that are characteristic of trichomes synthesizing secondary metabolites of the different compositions will be demonstrated.

6.1

Synthesis and Accumulation of Phenolic Substances

The crucial role in synthesis and transport of phenolic compounds belongs to the smooth endoplasmic reticulum. According to the molecular biological studies, there are multienzyme complexes, or metabolons, catalyzing sequential reactions of flavonoid and phenylpropanoid metabolism that are situated on the membranes of the endoplasmic reticulum [45, 140]. It was shown that L-phenylalanine ammonialyase, cinnamate 4-hydroxylase, and later enzymes of multienzyme complexes are on the ER in Arabidopsis thaliana [141] and Nicotiana tabacum [142]. Leaf trichomes of Betula species were studied as example of the glandular trichomes synthesizing phenolic compounds [62]. They are characterized by many traits indicative of high metabolic activity, e.g., the occurrence of RER, numerous plastids, osmiophilic vacuoles, multivesicular bodies, and lipid droplets. The smooth reticulum is an obligatory organelle of the peltate glandular trichomes of Pterocarya rhoifolia [60] synthesizing catechin and quercetin, of the capitate glandular trichomes of Empetrum nigrum [34] in which bibenzyls and chalcones are the main secondary metabolites, of Arnica chamissonis participating in biosynthesis of sesquiterpene lactones (Fig. 3a, unpublished data), and also of insectivorous plant Drosophyllum lusitanicum containing anthocyanins (Fig. 3b). The degree of SER development varies in the cells of the glandular trichomes of the different localization. In the trichomes of E. nigrum, a more abundant network of SER is formed in the middle and lower cells of a head compared with the upper cells. Besides SER, secretory cells of the glandular trichomes synthesizing phenolic substances contain leucoplasts of various shapes (Fig. 4). A typical feature of the plastids in these trichomes is a high ratio of surface area to volume of one organelle; often leucoplasts are so elongated that just a narrow layer of stroma is inside an envelope (Fig. 4f). Plastids with cup-shaped invaginations were also found in the

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Fig. 3 Endoplasmic reticulum in the secretory cells of the glandular trichomes synthesizing secondary metabolites. (a) An abundant smooth endoplasmic reticulum (SER) in the trichomes of Arnica chamissonis, synthesizing sesquiterpene lactones. (b) Elements of the smooth and rough endoplasmic reticulum (RER) in the trichomes of Drosophyllum lusitanicum, synthesizing flavonoids; GA Golgi apparatus. (c) Tubules of the smooth endoplasmic reticulum with the grey content in the trichomes of Tussilago farfara, synthesizing phenols and terpenoids; Mb microbody. (d) Cisterns of the rough endoplasmic reticulum with the black content in the trichomes of Doronicum macrophyllum, synthesizing phenols and terpenoids. The same content there is in the periplasmic space. Scale bar – 0.5 μm

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Fig. 4 Leucoplasts in the secretory cells of the glandular trichomes synthesizing secondary metabolites. (a) Trichomes of Arnica longifolia, synthesizing sesquiterpene lactones. Endoplasmic reticulum forms a sheath around plastid envelope (arrow); M mitochondrion. (b) Trichomes of Doronicum pardalianches, synthesizing phenols and terpenoids. (c) Trichomes of Empetrum nigrum, synthesizing bibenzyls and chalcones; MvB multivesicular body (d) Trichomes of Drosophyllum lusitanicum, synthesizing flavonoids. (e) Trichomes of D. rotundifolia, synthesizing naphthoquinones. (f) Trichomes of Tussilago farfara, synthesizing phenols and terpenoids; OI osmiophilic inclusion. Scale bar – 0.5 μm

trichomes in Juglans species [75], synthesizing flavonoids, polyphenols, and tannins. Cheniclet and Carde [143] suggested that increasing surface area indicates structural prerequisites for intensive exchange by metabolites between the plastids and surrounding cytoplasm. In secretory cells of the trichomes, there is a close association of a plastid envelope with a reticular sheath in order to transport metabolites from leucoplasts into cisternae of RER [34] (Fig. 4a, b, e, arrows). This characteristic is typical for various glandular structures producing phenolic substances [60, 62]. Before, it has been shown that plastids in some way take part in the synthesis of different phenolic compounds (flavonoids, chlorogenic, and p-coumaric acids, esters of hydroxycinnamic acids, and others [144, 145]). There are two possible paths for the synthesis of above-mentioned substances: either the leucoplasts of the secretory cells contain the same enzymes as the multienzyme complexes on the membranes of the ER or the precursors of the phenols are synthesized in the plastids and then transported to cisternae of endoplasmic reticulum associated with the leucoplast envelope for subsequent modification.

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In glandular trichomes of some plants, for example, E. nigrum or Drosera rotundifolia, there is an accumulation of lipophilic material in the plastids in the form of the plastoglobules (Fig. 4c, e). Moreover, homogeneous electron-dense droplets are also found in cytosol of secretory (Fig. 4f) and especially stalk cells. The same lipid droplets were in the secretory cells of the glandular trichomes in some Betula species producing flavonoids [62]. Prevention of the toxic effect of phenolic compounds on the cytoplasm of the cells is achieved by their isolation inside vacuoles as water-soluble glycosides and sulphates [146, 147] or in the cell wall as aglycones. Flavonoid aglycones are revealed in secretion of some glandular trichomes [42, 132]. It was demonstrated that ca. 30% of the whole petal flavonols are located in the cell wall [148]. Before entering the cell wall, phenols are secreted from the cytoplasm into the periplasmic space. In the glandular trichomes, vacuoles can be transparent and contain loose sediment [79] (Fig. 5a), large osmiophilic globules [149] (Fig. 5b), or osmiophilic layer on tonoplast [62, 150] (Fig. 5c). If flavonoids are in vacuoles, they may be representing in the form of water-soluble glycosides [97]. Usually dark inclusions in vacuoles are regarded as tannins or other phenolic compounds forming a deposit during preparation of the plant material for transmission electron microscopy [39, 151–153]. Moreover, phenols are capable of producing some complexes with carbohydrates and proteins, which are visible as intravacuolar bodies [154, 155]. Spatial separation of transparent vacuoles and vacuoles with a black precipitate was demonstrated in the glandular hairs of Drosera rotundifolia, synthesizing polysaccharide mucilage as well as flavonoids and naphthoquinones [149], and in peltate trichomes of Betula species, synthesizing polysaccharides and flavonoids [62]. There is a correlation between the presence of osmiophilic deposition in the vacuoles of the middle and lower cells of the head in Empetrum nigrum and a higher degree of SER development. Having established the presence of bibenzyls, chalcones, and flavons in methanol extracts of leaves, Muravnik and Shavarda [34] concluded that phenolic substances synthesized on the SER are accumulated in the vacuoles of the glandular trichomes. To study the location of biosynthetic pathways of flavonoids at the cellular and subcellular level, immunohistochemical techniques were used [45]. Schopker and coworkers [45] demonstrated in situ localization by immunofluorescence and immunogold labeling of the two central enzymes of flavonoid biosynthesis, phenylalanine ammonia-lyase (PAL) and chalcone synthase (CHS), in glands of Primula kewensis. Both enzymes were exclusively found in the head cells of the glands. PAL is located in the regions of the dense tubular endoplasmic reticulum and occasionally found in more or less spherical organelles that have not yet been identified. Furthermore, an appreciable proportion of the enzyme protein was associated with the plasma membrane and the cell wall of the head cells. In contrast, the occurrence of CHS was restricted to the spherical, unidentified cell compartments. These findings indicate that the gland cells take part in flavonoid biosynthesis.

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Fig. 5 Vacuoles in the secretory cells of the glandular trichomes synthesizing phenols. (a) Vacuoles (V ) with fibrils in the trichomes of Empetrum nigrum, synthesizing bibenzyls and chalcones; L leucoplast. (b) Vacuoles with large osmiophilic globules in the trichomes of Drosera aliciae, synthesizing anthocyanins. Scale bar – 2 μm. (c) Vacuoles with osmiophilic deposition in the trichomes of Aldrovanda vesiculosa, synthesizing naphthoquinones. Scale bar – 1 μm (a); 2 μm (b, c)

6.2

Synthesis and Accumulation of Monoterpenes

Terpenoids are the most common metabolites secreted by glandular trichomes of taxonomically distant plants [156, 157]. At the same time, monoterpenes are more often found in plants belonging to the family Lamiaceae. The glandular trichomes, in which biosynthesis of monoterpenes occurs have the following principal ultrastructural features: amoeboid leucoplasts, abundant endoplasmic reticulum, and numerous mitochondria. The secretory cells of the mature glandular trichomes contain leucoplasts of various forms, in which surface/volume ratio is often increased in the same way as in trichomes synthesizing phenolic compounds (Fig. 3). Elongated leucoplasts are found in the trichomes of Artemisia annua [22], Nepeta cataria [158], some Juglans species [75], and many other plants. Occasionally, leucoplasts have cup-shaped invaginations (Fig. 4f). Expanded surface of leucoplasts indicates that there is intensive metabolic exchange between the plastids and other organelles of cytoplasm during an active secretion stage [143]. Leucoplasts of the glandular trichomes are often situated in close association with endoplasmic reticulum elements; probably ER-plastid complexes facilitate transfer

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of terpene precursors from plastids to reticular tubules or cisternae. This feature is typical for various glandular structures producing terpenes [139, 153, 159, 160]. Plastids of the secretory cells can have numerous lamellae filled with the osmiophil inclusions (Fig. 4b), for example, in the secretory hairs of Inula viscosa [84], Origanum dictamnus [15]; Achillea millefolium [139] Helichrysum aureonitens [82], or Tussilago farfara [63]. Analytical and structural studies performed on 45 species of higher plants showed a very close correlation between the presence of leucoplasts in the secretory cells and the amount of monoterpenes in the extracts [143]. Moreover, key enzymes of monoterpene biosynthesis have been found in leucoplasts [161]. Formation of numerous elements of smooth endoplasmic reticulum is a common characteristic of the secretory cells of the glandular trichomes synthesizing terpenes [22, 158, 160]. It consists of branched tubules localized predominantly at the periphery of the cytoplasm that promotes transport of the metabolites through plasma membrane. Sometimes the tubules of smooth endoplasmic reticulum have an opaque content [18, 63, 160] (Fig. 3c, d). In the glandular trichomes synthesizing and accumulating monoterpenes, activity of Golgi apparatus relates not only with the synthesis of the carbohydrate components of secretion, but also with the synthesis and transport of hydrolytic enzymes. Hydrolases carried through the plasma membrane participate in loosening of the apical cell wall and thereby facilitate transport of the secretory products [162]. Endoglycanases can play a main role in this process [18]. As a rule, the secretory vesicles with a smooth membrane are rare in the lipid glandular trichomes; however the coated vesicles are separated from trans-Golgi reticulum [18, 60]. Just with coated vesicles, the transport of acid hydrolases through the plasma membrane is usually associated [163, 164]. In the head cells, there are a great number of mitochondria that locate at the peripheral cytoplasm. They illustrate the higher metabolism of the secretory cells in relation to other tissues. On the advanced stages of secretion, lipid and numerous osmiophilic droplets in the cytoplasm of the head cells have also been found in trichomes producing monoterpenes [15, 87, 138]. A detailed ultrastructural study of the peltate glandular trichomes in Mentha x piperita in relation to monoterpene production and the mechanism of essential oil secretion were completed by Turner et al. [19]. On the presecretory stage, terminal cells resemble meristematic cells because they contain proplastids and small vacuoles. The secretory stage coincides with the maturation of the secretory cell leucoplasts in which monoterpene biosynthesis is known to be initiated [165], and the formation of extensive smooth endoplasmic reticulum occurs. The smooth endoplasmic reticulum appears to form associations with both the leucoplasts and the plasma membrane; it often contains densely staining material and may be involved in the transport of the monoterpene-rich secretion product. Ultrastructural transformations include the formation of a large subcuticular secretion storage space above the trichome head cells. The glandular stalk cells also undergo intriguing modifications correlated with secretory activity. These alterations include the

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development of large plastids, numerous microbodies, and abundant mitochondria. The presence of micro bodies suggests active oxidative metabolism. In order to prove the participation of some organelles of the glandular trichome cells in the biosynthesis of monoterpenes, immunocytochemical localization of the enzyme activity was studied. The enzymes, geranyl diphosphate synthase (GPPS) and limonene synthase (LS), involved in monoterpene biosynthesis in mint, have been localized by immunogold labeling in the leucoplasts of peppermint gland secretory cells [166]. This was the only cell type within the glandular trichomes that was labeled by the antilimonene synthase polyclonal antibodies. The cytochrome P-450-dependent monooxygenase limonene 6-hydroxylase (L6OH) was associated with the endomembrane system, specifically with the endoplasmic reticulum [47]. Trans-isopiperitenol dehydrogenase (IPD) label was found in the mitochondria and pulegone reductase (PR) label in the cytoplasm. The localization of these five enzymes of monoterpene synthesis verifies that formation of this class of compounds is indeed restricted to the secretory cells of peltate glandular trichomes in mint. Thus, multiple subcellular compartments are necessary for monoterpene biosynthesis.

6.3

Synthesis and Accumulation of Sesquiterpene Lactones

STL are a subclass of sesquiterpenoids. Since STL are very toxic to the cytoplasm, they accumulate in the subcuticular cavity of the glandular trichomes. The tissue localization of the antimalarial sesquiterpenoid compound artemisinin in Artemisia annua was determined by differential extraction of glanded and glandless biotypes [101]. Results indicated that artemisinin as a product of secretion is entirely localized in the subcuticular space of the capitate glands of A. annua. The main ultrastructural features of the glandular trichome cells involved in the biosynthesis and removal of sesquiterpene lactones are presented below. In all cells of the upper layers of the glandular trichomes, SER is the main organelle. The black inclusions are revealed, for instance, within the reticular tubules in the glandular trichomes of Sigesbeckia jorullensis [167], Tussilago farfara [63], and Doronicum species [41]. Besides tubules of SER, RER cisterns are found. The activity of the Golgi apparatus is predominantly directed not at the synthesis of the carbohydrate component of the secret, but at the production of the hydrolytic enzymes that is confirmed by the appearance of the trans-Golgi reticulum with coated vesicles. The hydrolytic enzymes are necessary for loosening of the apical cell wall to facilitate transfer of the secretory products. In the glandular trichomes of Asteraceae family, there are two populations of plastids, namely leucoplasts and chloroplasts. This is due to different functions, which plastids carry out. If simple sugars are formed in chloroplasts as a result of photosynthesis, after that terpene precursors appear in leucoplasts [22]. For instance, the apical cells in Artemisia annua, Achillea millefolium, Vernonia galamensis, or Tussilago farfara glandular trichomes have amoeboid leucoplasts [22, 63, 79, 139]. The subapical cells possess both leucoplasts and large

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chloroplasts without starch grains. The stalk cells contain only chloroplasts. The basal cell pair has leucoplasts with osmiophilic globules attached to their outer envelope. Beginning of secretion is accompanied by the appearance of an osmiophilic product in the stacked thylakoids, plastid envelope, and smooth endoplasmic reticulum. So far, as the peripheral cytoplasm of the apical cells is enriched in SER tubules, osmiophilic globules are seen in the periplasmic space too [22, 81]. Gradually, a small globular deposit increases in dimension [25]. The precipitate appears not only in the apical cell layer, but also behind the plasma membrane of the cells of several underlying layers. Numerous osmiophilic inclusions in cytosol resembling lipid drops were shown in the glandular trichomes of Arctium lappa [168], as well as of Artemisia annua [22] and in the cytoplasm of the cell suspension culture of Achillea millefolium [139]. These structures are formed, by all appearances, as a result of aggregation of fatty acids, the quantity of which in glandular trichomes is rather high. The first committed steps in biosynthesis of all cyclic sesquiterpenes and sesquiterpene lactones are conducted by amorpha-4,11-diene synthase (ADS) [169] and amorpha-4,11-diene hydroxylase, a cytochrome P450 monooxygenase (CYP71AV1) [170]. Artemisinic aldehyde D11(13) reductase (AAR) is third enzyme of artemisinin biosynthesis [171]. In order to investigate the function of different cells of multicellular glandular trichomes, a method based on the laser microdissection pressure catapulting technique has been developed for isolation of different whole cells from A. annua glandular trichomes [172]. Experiments showed expression of the three abovementioned enzymes involved in artemisinin biosynthesis only in the apical cells. No detectable amplification was observed in the subapical cells. Therefore, it was concluded that artemisinin biosynthesis takes place in the two outer apical cells, while the two pairs of chloroplast-containing cells have other functions in the overall metabolism of the glandular trichomes. In the glandular trichomes of Helianthus annuus, the second model species of Asteraceae family, three enzymes were identified as two germacrene A synthases (GAS) and one δ-cadinene synthase [173]. All three sesquiterpene synthases were predominantly expressed in the stalk cells of capitate glandular trichomes at active secretory stage. Amrehn et al. [48] established a polyclonal antibody against germacrene A oxydase (GAO) and followed the expression of this enzyme in cells of different developmental stages of glandular trichomes by means of immune labeling with fluorescence microscopy. The immune label correlated with the appearance of SER in secretory and stalk cells. Stalk cells and secretory cells differed in form, size, and types of plastids, but both had structures necessary for secretion. No GAO-specific immunoreaction was found in sunflower leaf tissue outside of glandular trichomes or in developing glandular trichomes before the secretory phase had started. The results indicated that not only secretory cells but also nearly all cells of the glandular trichomes were involved in the biosynthesis of STL and that this process was not linked to the presence or absence of a specific type of plastids.

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Synthesis and Accumulation of Cannabinoids

Glandular trichomes in Cannabis sativa (Cannabaceae) are always of great interest due to their terpenophenolic compounds, the cannabinoids synthesizing ability. Cannabinoids include the marijuana hallucinogen, tetrahydrocannabinol (THC). The ultrastructure of the glandular trichomes possesses the general pattern for terpene secreting glands in having a dense cytoplasm, abundant endoplasmic reticulum, rare Golgi bodies with secretory vesicles, and numerous leucoplasts. However, C. sativa displays several distinctive ultrastructural and functional features that are not characteristic of other lipophilic glands. Foremost among these features is development and function of its highly modified plastids [86]. The plastids in pre-secretory stages resemble typical leucoplasts with few internal membranes, but with the onset of secretion undergo a rapid and extensive increase in number and develop the highly organized paracrystalline bodies. The paracrystalline bodies increase in size until they nearly fill the plastid, leaving only a small open stomal area. Formation of the secretory cavity begins as a swelling and loosening within the upper cell walls [8]. Initial stages reveal a fibrillary character of the wall, indicating on a hydrophilic compartment within the matrix. The hyaline flatted areas represent a second compartment containing electron-dense secretory products. They usually lie parallel to the plasma membrane. The same inclusions were found in the capitate glandular trichomes of Stachys heraclea [69] and in the cylindrical trichomes of Doronicum species [41], which was named as electron dense fibrillar material or interfibrillar lamellae. The lipophilic vesicles bounded by a unit membrane are associated with the secretory cavity. During development of the secretory cavity, its upper wall increases in thickness. Kim and Mahlberg [8] concluded that maintaining of a uniformly thick wall under conditions of the secretory cavity development is possible if the glandular cells introduce new components into the growing wall. Localization of delta 9-tetrahydrocannabinol in glandular trichomes of C. sativa bracts was shown with a monoclonal antibody–colloidal gold probe by electron microscopy [46]. Upon formation of the secretory cavity, the immunolabel detected THC in the upper cell wall facing the cavity as well as the subcuticular wall and cuticle throughout development of the secretory cavity. THC was detected in the fibrillary matrix associated with the disc cells and with matrix in the secretory cavity. The antibodies identified THC on the surface of secretory vesicles, but not within the secretory vesicles. Gold label also was localized in the anticlinal walls between adjacent disc cells and in the wall of epidermal and mesophyll cells of the bract. Grains were absent or detected only occasionally in the cytoplasm of secretory or other cells of the bract. No THC was detected in controls, which included sections treated with antibody alone, or treated with protein A-gold alone. These results indicate THC to be a natural product secreted particularly from disc cells and accumulated in the cell wall, the fibrillar matrix and surface feature of vesicles in the secretory cavity, the subcuticular wall, and the cuticle of the glandular trichomes. THC, among other chemicals, accumulated in the cuticle may serve as a plant recognition signal to other organisms in the environment.

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Repeatedly, it has been found that during maturation of the glandular trichomes the following changes take place in their metabolic activity. In the secretory cells of immature trichomes, the synthesis of polysaccharide and hydrolytic enzymes occurs, which is accompanied with Golgi apparatus activity and the proliferation of RER [50, 69, 73, 139]. In some plants, acid polysaccharides secreted by trichomes can protect the growing shoot against desiccation and mechanical damage during growth; in other plants, mucilage in the secretory drop can be attractive to pollinating insects. The hydrolytic enzymes contribute to loosening the cell wall making its structure permeable to lipids [18, 19, 162]. In the mature trichomes, synthesis of the secondary metabolites predominates; it involves participation of the dense network of SER and numerous leucoplasts, possessing an enhanced surface area, infrequent lamellae, and reticular sheaths. Phenols, terpenoids, and sesquiterpene lactones play a role of the antibacterial, fungicide, and insecticidal barriers. Simultaneous presence of phenolics and terpenoids in secretion is typical for trichomes of the vegetative and reproductive organs [23, 36, 37, 62, 68, 69, 133, 152, 174]. In all studied plants, quantities of the synthesized compounds belonging to every class of secondary metabolites are specific. The ultrastructural characteristic of cells illustrates the structural features of the organelles involved in the synthesis of chemically different secondary metabolites.

7

Mechanisms of Secretion

Two distinct modes of secretion are there in the glandular structures: eccrine and granulocrine [3]. In the eccrine mode, substances are transported directly through the plasma membrane, whereas in the granulocrine mode secretory products are enclosed in specific vesicles for exocytosis. Mechanisms of secretion of the primary substances are well known. So, the polysaccharides are carried out of the cytoplasm of the secretory cells in a result of exocytosis of large Golgi vesicles [17, 37, 44, 125]. Hydrolytic enzymes are transported by coated vesicles, which can merge with each other, creating multivesicular bodies, and enter the cell wall [164] or the vacuoles [175]. Question relative to the modes of secretion of the secondary metabolites from cytosol to the periplasm space remains disputable. As the processes of biosynthesis of phenols and terpenes involve different intracellular compartments, it is obvious that transfer of the recently synthesized substances should also be realized by the different ways. Thus, some phenolic substances (e.g., anthocyanins) are sequestered in vacuoles through a glutathione pump in the tonoplast membrane [151]; others (flavonol glycosides) pass through plasma membrane by active molecule transportation and are accumulated in the cell wall [148]. According to Turner et al. [19], transport of monoterpenes from one subcellular compartment to another and the eventual secretion against the concentration gradient require a number of specialized lipid carrier and transfer proteins. Point of view of elimination of terpenoids from endoplasmic reticulum through the transient fusion of the reticular membrane with the plasma membrane or through

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the establishment of the permeable contacts between these membranes was suggested by Vassilyev [160]. Another probable mechanism for elimination of the essential oil from the cell supposes extravasation of the oil vesicles with plasma membrane [20, 176, 177]. After overcoming the plasma membrane, the secretion moves along the cell wall and reaches the cuticle. Hydrophilic substances, particularly polysaccharies, move across the cuticle along the branched dendritic structures, called otherwise fibrillary network, reticulate cuticle, or microchannels [178]. Microchannels are formed to facilitate secretion of the hydrated ionic compounds. Ions are lipid insoluble and require an aqueous pathway across cuticles. Average pore radii ranged from 0.45 to 1.18 nm [162]. Presence of micro channels was shown in the nectary cells [179, 180] or in the glandular trichomes [41] (Fig. 2f). For lipophilic substances, there is a parallel diffusion pathway, including cutin and amorphous waxes [162, 181]. According to the Schönherr [162], the permeability of the cuticle for lipids is increased with increasing solubility of the hydrophobic substances in these polymers. Paiva [182] recently hypothesized that a mechanical action of the protoplast is necessary to ensure the cell wall crossing by some substances. The mechanical action of the protoplast, in the form of successive cycles of contraction and expansion, causes the material accumulated in the periplasmic space to cross the cell wall and the cuticle. This action is particularly relevant for the release of lipids, resins, and highly viscous hydrophilic secretions. In the glandular trichomes of some plants that do not form a subcuticular cavity, the secretion can cross the cuticle through true pores or canals. The presence of cuticular pores is typical for the glandular trichomes of Droseraceae carnivorous plants [61]. These pores were found to have wall-free opening in the 80 nm thick cutinized layer. Formation of cuticular gaps takes place at the final stage of maturation of the glands. The positive correlation between the cuticle thickness and the diameter of the cuticular pores was shown in 36 investigated Drosera species [183]. In addition to Droseraceae plants, true pores, or openings, were found in the capitate trichomes of Stachys germanica [69] belonging to Lamiaceae and glandular trichomes of Arnica montana belonging to Asteraceae (our unpublished data).

8

Conclusions

Formation of the glandular trichomes on the surface of the vegetative organs is an adaptive reaction of the plants to the influence of various environmental factors. In response to excessive UVB radiation, increased temperature, and damage caused by herbivorous insects or pathogens, plants form secretory structures which can synthesize various compounds of primary and secondary metabolism. As a rule, the secretion of glandular trichomes changes during the plant development and is characterized by a mixed composition. In this connection, the various cell organelles are involved in biosynthesis. Numerous studies have shown that the presence of smooth or rough endoplasmic reticulum, leucoplasts of the various

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shapes and internal structure, the formation of reticular sheathes near the outer plastid membrane, as well as mitochondria are needed for synthesis of the secondary compounds. All biochemical diversity of secretory products is provided by the difference in multienzyme complexes localized on the membranes of these organelles. The accumulation of secretion occurs in vacuoles or in the wall of the secretory cells. Secondary metabolites of the plant origin are used in the manufacture of medicines, aromatic compounds for the food and perfume industry, as well as crop protection products. The glandular trichomes are often the only place of biosynthesis of these substances. Since secretory structures are localized on the surface of vegetative organs and can be isolated, they represent a convenient model system for a detailed study of biosynthetic processes of biologically active compounds. The genetic control of the biosynthesis of secondary substances, the study of regulatory mechanisms of gene expression, and the development of biotechnological methods prove to be the promising areas of glandular trichomes research. Acknowledgments I appreciate the “Core Centre Cell and Molecular Technology in the Plant Science” at the Komarov Botanical Institute (St. Petersburg) for provision of equipment for light and electron microscopy. The present study was carried out within the framework of the research project (АААА-А18-118031690084-9) of the Komarov Botanical Institute of the Russian Academy of Sciences.

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Accumulation of Secondary Metabolites and Improved Size of Glandular Trichomes in Artemisia annua Neha Pandey, Anupam Tiwari, Sanjay Kumar Rai, and Shashi Pandey-Rai

Contents 1 2 3 4 5 6

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Trichome Structure and Function in Artemisia annua . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Molecular Regulation of Glandular Trichome Development in Artemisia annua . . . . . . . . . Biosynthesis of Artemisinin in Glandular Trichomes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Metabolites of Glandular Trichomes of Artemisia annua . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Correlation Between Secondary Metabolite Accumulation and Size of Glandular Trichomes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.1 Growth Regulators, Trichomes, and Artemisinin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.2 Abiotic/Biotic Stresses, Trichomes, and Artemisinin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7 Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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N. Pandey Laboratory of Morphogenesis, Centre of Advanced Study in Botany, Institute of Science, Banaras Hindu University, Varanasi, Uttar Pradesh, India Department of Botany, CMP Degree College (A Constituent PG College of University of Allahabad), Prayagraj, Uttar Pradesh, India e-mail: [email protected] A. Tiwari Department of Botany, School of Bioengineering and Biosciences, Lovely Professional University, Phagwara, Punjab, India e-mail: [email protected] S. K. Rai Department of Horticulture, Dr. Rajendra Prasad Central Agricultural University, Pusa, Samastipur, India e-mail: [email protected] S. Pandey-Rai (*) Laboratory of Morphogenesis, Centre of Advanced Study in Botany, Institute of Science, Banaras Hindu University, Varanasi, Uttar Pradesh, India e-mail: [email protected] © Springer Nature Switzerland AG 2021 K. G. Ramawat et al. (eds.), Plant Cell and Tissue Differentiation and Secondary Metabolites, Reference Series in Phytochemistry, https://doi.org/10.1007/978-3-030-30185-9_31

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Abstract

Glandular trichomes are multicellular epidermal outgrowths that have characteristic globular head made up of secretory cells and store large quantities of specialized secondary metabolites. Artemisia annua is known for its medicinally important secondary metabolite “artemisinin” which is synthesized and stored in glandular trichomes. However, our understanding of morphological and transcriptional control related to glandular trichome development and accumulation of secondary metabolites in A. annua is available in scattered form. This chapter deals with the trichome biology including developmental and functional aspects along with their correlation with secondary metabolite accumulation in response to various biotic and abiotic signals of the environment using A. annua as model. This chapter also emphasizes the molecular mechanisms behind trichome development in A. annua and provides a glimpse of molecular players involved in this process. There are many environmental as well as intrinsic factors which directly or indirectly affect secondary metabolite synthesis and as a result determine the size of glandular trichomes. The compiled information available for A. annua trichome biology can further be utilized for exploring trichome engineering in many medicinal or aromatic plants which are less explored. Keywords

Artemisia annua · Glandular trichomes · Secondary metabolites · Trichome bioengineering · Artemisinin

1

Introduction

According to the World Malaria Report [48] issued by the World Health Organization (WHO), around 219 million cases of malaria have been reported in the year 2017 with an estimated death of 43,5000 patients, which is approximately 8% lower than 2010. However, this figure of reported malaria cases worldwide is insignificantly lower than cases reported during 2015–2016. If we look at the percent of reduction in malaria occurrence worldwide, the situation seems to be alarming. Plasmodium falciparum alone was found to be responsible for 99.7% of malaria in Africa region during 2017 and is also prevalent in Southeast Asia. However, P. vivax is the causal organism of 74.1% of malaria cases in America. Malaria and artemisinin have a strong relationship since hundreds of years. Use of Artemisia annua plant extract for treating malaria-like symptoms such as fever and chills has been reported since more than 2000 years in ancient systems of herbal medicine in China. Later on with enormous efforts of Tu You You and co-workers, the active ingredients of A. annua extracts and further structure of artemisinin are identified, and its derivative compounds were elucidated [14, 46]. For her work Tu You You has been honored with the 2015 Nobel Prize in Medicine or Physiology.

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Today artemisinin-based combination therapies are among the first-line drugs for treatment of malaria. Not only malaria, artemisinin was further found to be effective against numerous other disorders including cancer. Before 2013, drug artemisinin was solely extracted from Artemisia annua. However, with the success of “The Semisynthetic Artemisinin Project” funded by The Bill and Melinda Gates Foundation in partnership with the University of California, Amyris Inc., and a nonprofit organization called PATH Drug Solutions, drug production has increased [32, 33]. Recent reports suggest that dried leaves of A. annua are a better performer than pure artemisinin for treating resistant malaria as well as showed slower evolution of malaria drug resistance [6, 9]. Therefore, pharmaceutical or semisynthetic artemisinin can’t bear the burden of artemisinin-resistant malaria treatment worldwide. People are still focusing on plant-based drugs, and therefore it is necessary to better understand the chemistry of artemisinin biosynthesis in plant as well as the possible ways for its enhanced production in planta. Artemisinin is chemically a sesquiterpene lactone with an endoperoxide bridge and is synthesized and stored in special structures called glandular secretory trichomes. Leaves and stem of A. annua plant are covered with these glandular trichomes [7, 8]. Plants use these trichomes as excellent defense tools against a number of environmental threats imposed by herbivores, pathogens, fungi, bacteria, harmful radiations, extreme temperature, drought, and many others. There are a number of different metabolites which are stored in glandular trichomes and are poisonous to many herbivores and other pathogens. The present chapter deals with the trichome biology including developmental aspects, functional aspects, and their correlation with secondary metabolite accumulation in response to various signals. This chapter also covers the molecular mechanisms involved in trichome development in A. annua and also provides a list of molecular players involved in the process. This will provide a compiled data for further use in designing experiments related to glandular trichome bioengineering. The secondary metabolite population of glandular trichomes is species specific, and therefore in order to improve artemisinin production, it is necessary to identify molecular players involved in glandular trichome development and associated events in A. annua.

2

Trichome Structure and Function in Artemisia annua

Plants generally bear small specialized structures of epidermal origin on the leaf and/ or stem surface called as trichomes which play important defensive roles [31, 39]. According to their morphology, plants possess two kinds of trichomes: glandular and nonglandular. A. annua family Asteraceae is rich in glandular trichomes [45]. However, in A. annua both glandular secretory trichomes and T-shaped nonglandular trichomes are found, and both are of epidermal origin [7]. Glandular secretory trichomes of A. annua are ten-celled biseriate structure, each side

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Fig. 1 Structure of glandular secretory trichomes of Artemisia annua. (a) A leaf. (b) A portion of leaf bearing glandular trichomes as observed under florescence microscope. (c) Scanning electron micrograph of a leaf showing glandular trichome of A. annua. (d) Transverse section of young stem showing glandular trichome on the epidermis. (e) Highly magnified image of a leaf showing cellular structure of glandular trichome of A. annua. (f) Cellular diagram of a glandular trichome of A. annua showing subapical and apical cells and subcuticular space. (Credit for 1(E) goes to Meg Stark, Technology Facility, Department of Biology, University of York, UK)

consisting of one basal cell, one stalk cell, two subapical cells, and one apical cell [7, 31] (Fig. 1). These glandular trichomes function as physical barrier for plant and save them from various biotic and abiotic stresses. Trichomes are the storage organ for various secondary metabolites which are mostly poisonous to many insects/flies. They also protect plants from harmful radiations such as UV-B [11, 39]. Glandular trichomes of A. annua are known to synthesize and store artemisinin and many other secondary metabolites belonging to terpenoid, alkaloid, and polyphenol group. Beside this, they also possess some polysaccharides and calcium oxalate [3, 50]. Artemisinin biosynthesis was thought to take place in apical cells

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of glandular trichomes; however, Olofsson and co-workers [30] performed laser microdissection and RNA amplification and found that both apical and subapical cells produce artemisinin which is stored in the subcuticular space of glandular trichomes. Glandular trichomes are generally regarded as the only types of structure in A. annua which have the capacity to synthesize artemisinin [44]. Any other cell type such as the epidermis, mesophyll, or even T-shaped trichomes is considered to be incapable of artemisinin biosynthesis, and this is probably one of the reasons behind low concentration of artemisinin in plants. Very recently with the help of genetic breeding, Judd and team [17] have demonstrated production of artemisinin and its derivatives such as artemisinic acid, arteannuin B, and dihydroartemisinic acid in nonglandular trichomes of A. annua. This has opened new possibilities of engineering plants for higher production of artemisinin. The development of glandular trichomes in A. annua was closely monitored by Duke and Paul [7] with the help of transmission and scanning electron microscopy. They observed that in youngest leaf primordia, the foliar cell starts differentiating into the gland cell and after differentiation a ten-celled biseriate structure is formed. They also observed variable plastid population in different cell types of glandular trichomes. Apical cells contain mostly leucoplast, whereas subapical cells were found to have more amorphous chloroplasts.

3

Molecular Regulation of Glandular Trichome Development in Artemisia annua

Our current understanding of the development of glandular trichomes is quite fragmentary. However, the information available till date suggests involvement of various molecular players in glandular trichome development and determination of their density on plant organ. Being a terpene-rich plant, A. annua trichome development and related events are becoming a major research focus for many research groups around the world. Jasmonic acid plays a crucial role in glandular trichome initiation and development in A. annua. Interaction of HOMEODOMAIN PROTEIN 1 (AaHD1) with this important phytohormone is the major regulatory pathway for trichome initiation and development in A. annua. Figure 2 gives a hypothetical model for the regulation of trichome initiation in the presence and absence of jasmonic acid. Recently many molecular players were identified which have various roles in trichome initiation, development, and determination of trichome density (Table 1). MYB transcription factors have an important role in trichome development. Recently a MIXTA-type transcription factor was found to have a key role in trichome development [41]. Further it was found that this MIXTA-type transcription factor makes a complex with HD-ZIP IV (a homeodomain leucine zipper IV) transcription factor and this HD-ZIP IV/MIXTA complex promotes initiation of glandular trichome formation. This complex is also known to regulate cuticle development in A. annua [51]. Additionally, initiation of trichome

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Fig. 2 A hypothetical model explaining events and molecular players involved in glandular trichome initiation and development in Artemisia annua. AaHD8 (a homeodomain leucine zipper IV transcription factor) interacts with MIXTA-like protein AaMIXTA1, forming a regulatory complex which directly promotes the expression of AaHD1 (HOMEODOMAIN PROTEIN 1). AaHD1 remains bounded with AaJAZ (jasmonate ZIM proteins) and attenuates function of AaHD1 as well as suppresses the expression of AaHD1 downstream genes. This results in repression of jasmonic acid (JA)-mediated trichome initiation. Presence of JA causes SCF-COI1-mediated ubiquitination of AaJAZ which in turn degrades AaJAZ via 26 S proteasome system and releases AaHD1. The free AaHD1 now activates the downstream pathway genes involved in glandular trichome initiation and development. Many transcription factors regulate the downstream pathway. Initial AaHD8-AaMIXTA1 complex also regulates leaf cuticle development which indirectly helps in trichome development

development was found to be regulated by AaMYB1 transcription factor [29]. An AP2 transcription factor from A. annua, TRICHOME AND ARTEMISININ REGULATOR 1 (TAR1), is known to play a significant role in trichome development and artemisinin biosynthesis [42]. Another important transcription factor, AaWRKY, also regulates trichome development as well as artemisinin biosynthesis to a great extent. GLANDULAR TRICHOME-SPECIFIC WRKY 1 (AaGSW1) is a glandular trichome-specific transcription factor and regulates artemisinin biosynthesis and trichome development in A. annua [5]. Many other transcription factors regulate trichome development and artemisinin biosynthesis in direct or indirect ways. A basic leucine zipper transcription factor AaHY5 is known to be a positive regulator of artemisinin biosynthesis and acts via AaGSW1, which is another positive regulator of artemisinin biosynthesis [12].

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Table 1 List of genes/transcription factors and their role in determination of trichome development and density Plant Artemisia annua Artemisia annua

Gene name TTG1

Function Regulation of trichome density

References Lui et al. [22]

AaMIXTA1

Shi et al. [41]

Artemisia annua

HD-ZIP IV/MIXTA complex

Artemisia annua Artemisia annua Artemisia annua

AaHD1 (homeodomain protein 1) GL3

Regulation of trichome development and artemisinin biosynthesis Initiation of glandular trichome development and cuticle formation Glandular trichome initiation

Artemisia annua Artemisia annua

TFAR1

Artemisia annua Artemisia annua Artemisia annua Artemisia annua

AaMYB1

TAR1 (TRICHOME AND ARTEMISININ REGULATOR 1) AaGSW1 (GLANDULAR TRICHOME-SPECIFIC WRKY 1) AaHY5 AabHLH1 AaORA

Regulation of trichome density Regulation of trichome initiation and artemisinin biosynthesis Regulation of trichome density and morphology Regulation of trichome development and artemisinin biosynthesis Regulation of trichome development and artemisinin biosynthesis Positive regulation of artemisinin biosynthesis Positive regulation of artemisinin biosynthesis Positive regulation of artemisinin biosynthesis

Yan et al. [51] Yan et al. [51] Lui et al. [22] MatiasHernandez et al. [29] Maes et al. [27] Tan et al. [42] Chen et al. [5] Hao et al. [12] Li et al. [20] Ma et al. [25]

Many miRNAs have been recently found to be involved in trichome development in Arabidopsis such as miRNA156 and miRNA390 which target genes involved in trichome development [34, 52]. However, in A. annua trichome development, no such role of miRNA has yet been elucidated. Identification of various transcription factors involved in glandular trichome biology has interestingly opened a platform for bioengineering genes specific to glandular trichomes in A. annua. This has led to more profitable production of artemisinin in plants.

4

Biosynthesis of Artemisinin in Glandular Trichomes

Artemisinin biosynthesis takes place in cells of glandular trichomes in A. annua. Artemisinin biosynthesis begins with two pathways operating independently in cytosol and plastid. The cytosolic mevalonate pathway (MVA pathway) originates

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with acetyl-coA; however, plastidic MEP pathway (non-mevalonate pathway) starts with pyruvate and glyceraldehydes-3-phosphate. Both pathways lead to the synthesis of isoprenoid precursors: dimethyl allyl diphosphate (DMAPP) and isopentenyl diphosphate (IPP), both are interconvertible. After the synthesis of IPP and DMAPP, the rest of the pathway doesn’t lead independently, and cytosolic DMAPP is suggested to be transferred to plastid, and plastidic IPP leads to majority of farnesyl diphosphate (FPP) which ultimately forms artemisinin [40]. Key enzymes of MVA pathway are HMGS (3-hydroxyl-3-methyglutaryl CoA synthase), HMGR (3-hydroxyl-3-methyglutaryl CoA reductase), and IPI (IPP isomerase), while DXS (1-deoxy-D-xylulose-5-phosphate synthase), DXR (1-deoxy-D-xylulose-5-phosphate reductoisomerase), and IDS (IPP/DMAPP synthase) are key enzymes of plastidic MEP pathway. Cyclization of FPP is the rate-limiting step of artemisinin biosynthesis and leads to the formation of amorpha-4,11-diene with the help of amorpha-4,11-diene synthase (ADS). Beside this, many other sesquiterpene synthases are present such as caryophyllene synthase (CPS), germacrene synthase (GAS), epicedrol synthase (ECS), β-farnesene synthase (BFS), and squalene synthase (SQS) which leads to the synthesis of β-caryophyllene, germacrene A, 8-epicedrol, β-farnesene, and squalene, respectively. Another key enzyme from the downstream artemisinin biosynthetic

Fig. 3 Artemisinin biosynthetic pathway operative in cells of glandular trichomes on Artemisia annua

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pathway is cytochrome P450 enzyme, CYP71AV1, with dehydrogenase and reductase activities and is responsible for the oxidation of amorpha-4,11-diene into artemisinic alcohol and further into artemisinic aldehyde and finally to artemisinic acid. Further activities of DBR2, ALDH1, and RED1 enzymes ultimately form artemisinin and its derivatives (Fig. 3). Many transcription factors are known to regulate artemisinin biosynthesis by targeting key biosynthetic enzymes. AaWRKY1 transcription factor is a positive regulator of artemisinin biosynthesis by targeting ADS, CYP71AV1, and DBR2 gene [16, 35]. Ethylene insensitive3 (EIN3), an important transcription factor of ethylene signaling, targets three key genes of artemisinin biosynthesis, i.e., ADS, DBR2, and CYP71AV1, and causes downregulation of these genes in AaEIN3 overexpressing lines [43]. Promoters of ADS and CYP71AV1 are also known to contain E-box which is putative binding sites for an important group of transcription factors: basic helix-loop-helix (bHLH). In A. annua AabHLH transcription factors are known to have positive regulatory effect on artemisinin biosynthesis [15]. In addition, a trichome-specific transcription factor AaORA (expressed in both glandular and nonglandular trichomes of A. annua) also targets ADS, CYP71AV1, and DBR2 gene and positively regulates artemisinin synthesis in A. annua [24].

5

Metabolites of Glandular Trichomes of Artemisia annua

Glandular trichomes are specialized structure for synthesis and storage of secondary metabolites. Not all secondary metabolites are synthesized in glandular trichomes; however, they serve as storage organ for majority of terpenoids and phenylpropanoids. In addition, flavonoids, methylketones, and acyl sugars are also known to be stored in glandular trichomes [13]. A. annua is a terpenoid-rich species and produces some valuable terpenes including artemisinin and its various derivatives. By using gas chromatography-mass spectroscopy techniques, it becomes easy to know the secondary metabolite population of glandular trichomes. Table 2 gives a detailed list of terpenes present in glandular trichomes of A. annua. Three major terpene classes were found after conducting GC-MS, monoterpenes with carbon number 10, sesquiterpenes with carbon number 15, and diterpenes with carbon number 20. Apart from secondary metabolites, many other proteins and metabolites are present in glandular trichomes which perform diverse functions. Using different biochemical and molecular techniques, several genes specific to glandular trichomes have been identified, cloned, and characterized. Transcriptome and proteome of glandular trichomes of A. annua have been elucidated using high-throughput sequencing and comparative proteomic approaches [47, 49]. With the aim of identifying proteins involved in artemisinin biosynthesis, its transport, and trichome development, [49] isolated glandular trichomes from 4-year-old A. annua leaves for protein profiling using 2D gel electrophoresis and MALDI-TOF/TOF-MS (matrixassisted laser desorption/ionization time of flight mass spectrometry). More than

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Table 2 List of major terpenes present in glandular trichomes of Artemisia annua (data unpublished) Three major terpenes stored in glandular trichomes Monoterpene (C10) Sesquiterpene (C15) 1,8-Cineole Acetyl cedrene α-Terpineol Aristolene epoxide Camphene Thymol Camphor α-Bisabolol trans-Carveol γ-Cadinene Cis-Carveol β-Cadinene Carvone β-Caryophyllene Citronellol Δ-Elemene β-Cymene β-Guaiene Isoborneol α-Copaene Jasmone Corymbolone Lavandulyl acetate Cubebanol Limonene oxide Cyclocolorenone Myrcene 8-Epicedrol Myrtenol β-Farnesene α-Pinene (E, E)-α Farnesene β-Pinene Farnesol Pinocarvone γ-Elemen Sabinene Germacrene A γ-Terpinene Himachalol 4-Terpineol γ- Selinene Thujopsanone α-Humulene Verbenol Intermedeol Cymene Isogernial Isoeugenol Isocomene Sesquicineole α-Selinene α-Phellandrene β-Isocomene (Z)-β-Ocimene Υ-Eudesmol (E)-β-Ocimene Lanceol acetate δ-3-Carene Longifolol Terpinolene β-Longipinene Santolina alcohol β-Malliene trans-Sabinene hydrate β-Selinene cis-p-Mentha-28-dien-1ol Spathulenol Chrysanthenone Silphinene Myrtenal Valerenyl acetate trans-Pinocarveol Arteannuic alcohol trans-Sabinol β-Bergamotene Υ-Cadinone Dihydroartemisinic acid Cuminaldehyde Elemol para-Cymen-8-ol β-Ionone

Diterpene (C20) Larixol Phytol 3-Decanone Capillene Decanoic acid 4-Caprylmorpholine Hexadecanoic Caprylmorpholine n-Butyro-morpholine Simvastatin Tetradecane Tridecanal Docosane-n Lignocero Linoleic acid Octadecanoic acid Heptadecane Nonadecane Methyl hexadecanoate Hexadecanoic acid Methyl octadecanoate Octadecanoic acid Ethyl 2-methyl butyrate Octanal n-Hexanol Hexyl 3-methyl butyrate (Z)-3-Hexenol 1-Octen-3-ol (Z)-3-Hexenyl isovalerate Linyl acetate Hexyl tiglate 9-Decen-1-ol Methyl salicylate para-Mentha-1,4(8)-dien-3-ol Octadecane Benzyl isovalerate Eicosane Heneicosane Sanderacopamira-8(14),15-diene Tricosane Tetracosane (continued)

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Table 2 (continued) Three major terpenes stored in glandular trichomes Monoterpene (C10) Sesquiterpene (C15) cis-Sabinol β-Selinene Eugenol – Carvacrol – Tricyclene –

Diterpene (C20) Pentacosane Hexacosane – –

Fig. 4 Different functional classes of proteins identified from glandular trichomes of Artemisia annua

700 spots were identified, and functional classification revealed diverse classes of proteins (Fig. 4).

6

Correlation Between Secondary Metabolite Accumulation and Size of Glandular Trichomes

Leaves, green stem, and flowers of A. annua are covered with glandular secretory trichomes containing valuable drug, artemisinin. Since artemisinin is synthesized and stored in trichomes, it is obvious that greater size or density of these trichomes would result in higher amount of artemisinin in plant. Indeed, many workers have confirmed the existence of strong relationship between more or bigger trichomes and more artemisinin. Ferreira and Janick [55] had found greater density of trichomes on younger leaves as compared to older leaves. However, mature plants with maximum

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flowering are known to contain the highest amount of artemisinin, and it is at flowering and post-flowering stage when artemisinin is extracted from A. annua [28]. Lommen and team [23] had observed that out of the total artemisinin content of a plant, nearly 50% can be extracted from only senescent leaves. They also had observed that at mature stages these trichomes collapse and artemisinin is released which probably inhibits artemisinin biosynthesis. Further Arsenault and his team [2] confirmed the inhibition of artemisinin biosynthetic pathway after the trichomes are matured and ruptured. Trichome size and density in A. annua are regulated by many environmental factors.

6.1

Growth Regulators, Trichomes, and Artemisinin

Various phytohormones are known to regulate artemisinin biosynthesis as well as trichome development in A. annua. Gibberellic acid, cytokinins, and abscisic acid are among the regulators of artemisinin biosynthesis as well as trichome development [26]. In another study conducted by Maes et al. [27], BAP and gibberellic acid treatments were found to stimulate the development of trichomes in A. annua. Interestingly low-artemisinin-producing cultivars showed greater stimulation in terms of glandular trichome size and artemisinin accumulation as compared to high-artemisinin-producing cultivars of A. annua. Another growth regulator chlormequat is also known to regulate trichome development [21]. Growth hormone-responsive artemisinin production is in turn regulated by a number of transcription factors which interact with genes involved in its biosynthesis. Abscisic acid-mediated enhanced artemisinin biosynthesis involves active binding of AaABF3 (an abscisic acid-responsive transcription factor) onto the promoters of ALDH1, an important gene of artemisinin biosynthetic pathway [53]. Further role of AaABF3 in artemisinin biosynthesis was confirmed by performing gene overexpression and RNAi-mediated gene silencing. Overexpression and RNA interference of AaABF3 in A. annua resulted in increased and decreased artemisinin biosynthesis, respectively. Plant signaling molecules like jasmonic acid and salicylic acid are known stimulators of artemisinin biosynthesis [27, 36]. Spraying A. annua plants with jasmonic acid (100 μM) led to development of greater sized trichomes with more artemisinin content. It also caused higher density of trichomes on aerial parts of the plant. Greater artemisinin biosynthesis in response to jasmonic acid treatment was found to be associated with overexpression of many artemisinin biosynthetic pathway genes such as ADS, CYP71AV1, CPR, and ALDH1. Overexpression of these genes in response to jasmonic acid is suggested to be governed by AabHLH1 transcription factor which express to a great extent in glandular trichome-enriched tissues [20]. In addition, octadecanoid-responsive AP2/ERF (AaORA) transcription factor, which again is a trichome-specific transcription factor from A. annua, is involved in jasmonic acid-responsive trichome development and artemisinin biosynthesis. This AaORA was further shown to bind synergistically with AaTCP14 onto the promoters of DBR2 and ALDH1 gene of artemisinin biosynthetic pathway [25].

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Another signaling molecule, salicylic acid, was also found to upregulate artemisinin biosynthetic pathway genes which ultimately lead to enhanced artemisinin accumulation in glandular trichomes [36]. To attest these results, our team conducted an experiment [19] where arsenic-stressed plants were sprayed with salicylic acid which resulted in significant upregulation of ADS, CYP71AV1, DBR2, and ALDH1 genes. Salicylic acid treatment also had stimulatory effect on size of glandular trichomes and artemisinin accumulation.

6.2

Abiotic/Biotic Stresses, Trichomes, and Artemisinin

Environmental stress signals are commonly found to have stimulatory impact on secondary metabolite synthesis in plants. In A. annua, many abiotic and biotic stresses have been tested for their potential to enhance artemisinin synthesis. Concentrations of various elements which are present in soil also regulate secondary metabolite synthesis in plants. In A. annua also, many elements are suggested to have stimulatory effect on artemisinin biosynthesis and glandular trichomes. Nitrogen (N) is one of the most abundant elements of fertilizers. Therefore the effect of nitrogen fertilization on trichomes and artemisinin synthesis in A. annua was monitored by Bilkova et al. [4]. They found that plants grown in low nitrogen level had smaller leaves but greater trichome density as compared to plants grown in high nitrogen levels (more than 175 kg N ha 1). Application of many other minerals such as phosphorus and boron has been found to affect artemisinin biosynthesis. In addition boron application also caused oxidative stress which is suggested to be reduced with methyl jasmonate treatment [1]. However, artemisinin levels remain higher in both boron-treated and methyl jasmonate-treated plants. Our team was involved in deciphering the role of arsenic treatment on artemisinin, and we have found significant elevation in artemisinin content and greater size of trichomes in plants treated with arsenic [19, 37]. Silicon is one of the abundant elements found in soil and also accumulates in glandular trichome. Knowing this fact, Rostkowska et al. [38] found that silicon application in the form of calcium/magnesium silicate can increase trichome size with higher artemisinin accumulation. Similar to these elements, cadmium also has stimulatory impact on artemisinin biosynthesis in A. annua. However, its effect on trichome size or density has not been investigated, but as in other cases, higher artemisinin accumulation is related to greater trichome size; here also we may hypothesize bigger size of trichome in response to cadmium as well. While many elements were shown to have stimulatory impact on artemisinin biosynthesis, Ferreira [10] had shown significant increase in artemisinin production by creating specific macronutrient deficit conditions for A. annua. They have grown A. annua plants in acidic and macronutrient deficit soil which reduced leaf biomass; however, potassium deficiency significantly increased (21% higher than control) artemisinin production. Based on the results, they also recommended that farmers can increase the profitability of artemisinin production while saving on potassium fertilizer.

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Table 3 List of external factors influencing glandular trichomes (GT) and artemisinin content in Artemisia annua Stress Silicon

Effect on GT Increase in GT size

Arbuscular mycorrhiza (Rhizophagus irregularis) Growth regulator (chlormequat) Gibberellic acid

Increase in GT density Increase in GT size

Abscisic acid Cytokinin BAP Jasmonic acid Salicylic acid Salt Lower nitrogen Arsenic stress

Increase in GT density Stimulate GT development Increase in GT size and density Increase in GT size Increase in GT density Increase in GT size Increase in GT density Increase in GT density Increase in GT size

Effect on artemisinin Higher artemisinin Higher artemisinin Higher artemisinin Higher artemisinin Higher artemisinin Higher artemisinin Higher artemisinin Higher artemisinin Higher artemisinin Higher artemisinin Higher artemisinin Higher artemisinin

References Rostkowska et al. [38] Kapoor et al. [18] Liersch et al. [21] Maes et al. [27] Zhong et al. [53] Maes et al. [27] Maes et al. [27] Lui et al. [22]; Maes et al. [27] Pu et al. [36]; Kumari et al. (2017) Zhou et al. [54] Bilkova et al. [4] Rai et al. [37]

Trichome density was also improved when A. annua plants were inoculated with arbuscular mycorrhiza. This ultimately improved the artemisinin content of the plant [18]. Salinity is among the major abiotic factors responsible for reducing crop yield globally. However, when its effect on glandular trichome density and secondary metabolite accumulation was tested in A. annua, reverse results were observed. Moderate to severe salt stress caused significant increase in glandular trichome density on both sides of leaves in A. annua [54]. There are many abiotic factors whose effects on glandular trichomes are still not explored and need to be focused in near future (Table 3).

7

Conclusion

Artemisinin is a valuable drug and its global demand is very high. Glandular trichomes are the biofactories that synthesize and store artemisinin and many other secondary metabolites. Although the semisynthetic artemisinin production has been

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achieved and it is contributing well to the global supply, recent reports suggest consumption of natural artemisinin over synthetic drug for better absorption and results. This again generates the need for increased in planta production of artemisinin along with its derivatives and flavonoids which are supposed to synergize the antimalarial effect of drug. In this regard it becomes necessary to decipher complete regulatory pathways involved in glandular trichome development and artemisinin biosynthesis. As many reports suggest, these two events (trichome development and artemisinin biosynthesis) are highly coordinated. Quite a few reports suggest higher artemisinin content in leaves having greater or bigger glandular trichomes. We have made a sound progress in revealing the molecular players regulating trichome development and artemisinin biosynthesis. This provides ample opportunities for bioengineering glandular trichomes of A. annua and enhancing artemisinin production in plants. Although many genes from the artemisinin biosynthetic pathway have been cloned and overexpressed in A. annua which resulted in higher artemisinin, lesser studies have been conducted for targeting glandular trichomes. It is high time to go for glandular trichome engineering by targeting upregulation or downregulation of genes involved in trichome development, targeting endogenous miRNAs or manipulating external environment where the plant is growing. This will surely help us in winning the battle against low supply to demand ratio of artemisinin.

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5

A Model for Resin Flow Paulo Cabrita

Contents 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Duct Structure, Development, and Distribution . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1 Gymnosperms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2 Angiosperms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Synthesis and Secretion into Ducts . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 A Hydrodynamic Model . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1 Model Assumptions and Governing Equations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2 Flow Properties . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5 Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Abstract

Resins are plant exudates of economic importance composed of volatile and nonvolatile compounds, sometimes including gum, that are used by plants mostly as defense against invading pathogens. They are synthesized by epithelial cells surrounding intercellular spaces, called ducts, where they are stored under pressure. Despite the differences in the structure and distribution of resin ducts between gymnosperms and angiosperms, as well as in the mechanisms of resin loading, there are similarities in the importance of resin in both groups of plants. This chapter presents a model that applies the unsteady Stokes equation and describes resin flow in plants, considering resin viscosity, the structure of resin ducts, and a pressure-driven granulocrinous loading of resin through the duct wall. Resin flow is shown to increase towards the duct open end. Both flow and P. Cabrita (*) IAPN – Institute of Applied Plant Nutrition, Georg-August University of Göttingen, Göttingen, Germany e-mail: [email protected] © Springer Nature Switzerland AG 2021 K. G. Ramawat et al. (eds.), Plant Cell and Tissue Differentiation and Secondary Metabolites, Reference Series in Phytochemistry, https://doi.org/10.1007/978-3-030-30185-9_5

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pressure within the duct depend on the loading of resin and on the duct specific resistance, which depends on the duct geometry, viscosity, and duct wall permeability to resin. The dynamics of flow within resin ducts seems physiologically advantageous for the defense role commonly attributed to resin and agrees with previous measurements and observations. Understanding how resin flow is affected by these physiological and morphological parameters might be useful not only to better understand the physiological role of resin but also improve and develop more efficient and sustainable tapping methods. Keywords

Resin · Resin flow · Gymnosperms · Angiosperms · Resin ducts · Gum · Unsteady Stokes equation · Granulocrine secretion

1

Introduction

The use of resin by human societies is quite old and diverse, going through the preindustrial era and surviving up to the present day, similarly to other plant exudates like essential oils, gum, gum resin, mucilage, and latex, as shown by different historical records. Resin is a viscous plant exudate composed of a heterogeneous mixture of volatile and a smaller fraction of nonvolatile terpenoid and/or phenolic secondary compounds, which vary among plant species and are typically secreted and stored under pressure in specialized structures, called ducts, or cells that are located either internally (idioblasts) or on the surface of the plant body (trichomes). The synthesis of the different components of resins involves plastids, the endoplasmic reticulum, and the cytosol. When present, the gummous components originate from the sloughing of the cell walls lining the duct lumen. Occurring in cell interiors as well as cell walls and tending to stop or lessen the loss of water from plant tissues, resin is often mentioned as playing an important role in plant defense mechanisms against herbivory, as well as fungi and bacteria, thus making wood resistant to decay. In fact, a big part of the continued success of conifers, which appeared around 200 million years ago [1], stems from the mechanisms developed to defend them against invading organisms, including the synthesis of resin, and sterilize, seal, and heal damaged tissues. However, despite its potential significance in ecological interactions with plants, the physiological role of resin is not yet completely understood. For example, resin seems to have little or no importance in the primary or fundamental physiological processes of plants, e.g., serving as reserve food. On the other hand, resin is also secreted in the floral structures and fruits of some angiosperms to attract pollinators and facilitate the dispersion of seeds [2–4]. Typically and contrary to gum, resin is insoluble in water and differently soluble in vegetable oils and organic solvents [5, 6]. It may occur alone or in combination with essential oils, gums, or both, being called oleoresins, gum resins, and oleo-gum-resins, ranging in color from white to yellow, pink, brown, or even black. Within oleoresins, some authors distinguish yet balsams as having cinnamic acid and/or benzoic acid or

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their esters [5]. Among angiosperms, Pistacia lentiscus L. is, perhaps, the most wellknown source of oleoresin, while Boswellia, Calophyllum, Commiphora, and Ferula are probably the most important sources of oleo-gum-resins [5, 7]. However, resin composition may differ significantly between species and even individuals [8]. Commonly, resin yields are greater in conifers than in angiosperms, and within conifers higher yields are noticeably observed in members of Araucariaceae and Pinaceae and within these particularly in the genus Pinus. Several aspects of resin synthesis such as biochemistry, anatomy, and physiology of the secretory structures, as well as its ecological role, have been investigated more frequently and intensively in conifers than angiosperms. This difference in interest seems to reflect the higher economic value of conifers compared to angiosperms, as well as their prominence in the forests of temperate zones, where, for historical reasons, most of the associated activities and applications of resin originated. In this chapter, flow within resin ducts is discussed by applying fluid dynamics considering several physiological and anatomical aspects. The main goal is thus to present how physiological factors such as the loading, solidification, and viscosity of resin, as well as anatomical factors such as duct size and structure, affect flow within ducts and could explain differences between species. It is hoped that this model could not only serve as basis to a better understanding of resin synthesis and its physiological role but also improve and develop more efficient and sustainable tapping methods.

2

Duct Structure, Development, and Distribution

Resins, gum resins, and oleo-gum-resins are distributed widely among plant species, occurring in almost all plant organs, from roots to flowers. The synthesis of resin can occur in trichomes, idioblasts (resin cells), and secretory cells surrounding intercellular spaces that form ducts. The development of these structures and the subsequent synthesis of resin can be natural or induced. Those structures induced by damage of the cambium, which is caused by frosty, mechanical, chemical, or pathogenic origin, are called traumatic, opposite to natural or constitutive structures. Their occurence and distribution vary among plant species [9]. Elongated ducts of more or less uniform tubular shape that can attain considerable size are often called canals, otherwise when enlarged and irregularly formed they are designated by cavities. Smaller ducts of approximately spherical or ellipsoidal shapes, occurring isolated or in series, are called pockets or cysts. In common, all these specialized intercellular spaces are surrounded by one or more layers of specialized parenchyma cells that make the epithelium and secrete resin, gum resin, or oleo-gum-resin into the intercellular spacesand lumina. Epithelial cells can be further associated to neighboring specialized cells (parenchyma and tracheids, if present), called subsidiary cells [10]. Given the common origin of both epithelial and subsidiary cells from fusiform initials that underwent mitotic divisions [9, 11] and their close physiological relationships, Wiedenhoeft and Miller [10] suggested the term resin canal complex in referring to these cells, similar to the terminology used in leaf anatomy for the stomatal complex [12] or the sieve element-companion cell complex (SE-CC)

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observed on the phloem [13]. Resin ducts can be distributed axially and radially throughout the plant body, but radial ducts occur in the secondary phloem in fusiform rays only, and by connecting vertical ducts, they can sometimes form considerable three-dimensional networks [14–17]. Traumatic resin ducts, caused by wounding and often composing an anastomosing network, are frequently cystlike and occur in tangential series mostly [9]. In some species, they can be longer and wider than normal vertical ducts, e.g., in Picea abies (L.) H. Karst., traumatic resin ducts can be up to four times wider than normal vertical ducts, becoming further dispersed at a distance from the wound [9, 18, 19].

2.1

Gymnosperms

Although a normal feature easily recognized and associated with conifers, the structure and distribution of resin ducts in number as well as plant organs differ among gymnosperms [20]. Normal or constitutive resin ducts are scattered and found in all plant organs in both primary and secondary tissues, while in some genera, only traumatic ducts are found in the wood (Table 1). Normal resin ducts develop similarly throughout the primary and secondary tissues of the plant body [21, 22]. Typically, a small group of duct initials is formed by meristematic tissues, i.e., the procambium in the primary growth and the cambium in secondary growth. An intercellular space is then created by the dissolution of the middle lamella between the duct initials, causing their separation at the center of the group in what is called schizogenous development [16]. Also, either prior to, during, or after the formation of the intercellular space that will become the duct lumen, the initials undergo some mitotic divisions. Concomitantly, the initials enclosing the duct thus formed differentiate into epithelial cells, distributed in one to three layers surrounding the duct lumen. As development proceeds, the intercellular space formed is further widened by tangential flattening, separation, and anticlinal divisions of the epithelial cells that are oriented longitudinally among tracheary elements and radially through vascular rays [22, 23]. The epithelial cells of vertical ducts are derived from derivatives of fusiform cambial initials, whereas those of the radial ducts are derived from derivatives of fusiform ray initials in developing uniseriate rays. As the resin duct develops, the ray expands and becomes multiseriate. The epithelium is composed of varying proportions of thin- and thick-walled, living and dead cells, depending on age, organ of the plant, and species [23]. In some species, epithelial cells become thick-walled, lignified, and nonfunctional at maturity, hence with determinate growth, e.g., Picea, Larix, and Pseudotsuga [9, 35], while in others they remain thin-walled, unlignified, and functional for longer periods of time, e.g., Pinus [9]. Additionally, the epithelium may be surrounded by one to three layers of pectin-rich subsidiary cells (specialized parenchyma cells and/ or tracheids) with relatively thick walls, sometimes unlignified, which become nonfunctional and may become crushed as the duct develops and enlarges [10, 22, 36]. Epithelial cells are easily distinguishable from the neighboring parenchyma and subsidiary cells by having an organelle-rich dense cytoplasm. Compared with the

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Table 1 Distribution of secretory structures in gymnosperm genera known to yield gum resins, oleoresins, and resins Family Araucariaceae

Cephalotaxaceae

Cupressaceae Pinaceae

Podocarpaceae Sciadopityaceae

Genera Agathis Araucaria Wollemia Amentotaxus Cephalotaxus Torreya All Abies Cathaya Cedrus Keteleira Larix Picea Pinus Pseudolarix Pseudotuga Tsuga All Sciadopitys

Resin ducts Stem C, Pa, Phda C, P Absentb Cc Cc Cc C, Pd, Xd C, P, Xa C, X C, P, Xa C, Xe X C, P, X C, P, X Ca C, X C, Xa Absentb Absent

Leaves VB, M VB, M VB, M VB VB VB VB, M VB, M VB, M M VB, M M M VB, M VB, M VB, M VB, M VB, M VB, M

References [24, 25] [26, 27] [28, 29] [30, 31] [30, 31] [30, 31] [25, 31, 32] [23, 25] [23, 33] [23, 25] [9, 23, 34] [23] [23, 25] [23, 25] [23] [23, 25] [23, 25] [25, 31] [31]

Abbreviations: C cortex, M mesophyll, P phloem, Phd phelloderm, VB vascular bundles, X xylem a Only traumatic b Resin cells only c Only traumatic, otherwise resin cells are present under normal conditions d Only traumatic and in specific genera e Only in some species and in most cases of traumatic origin

neighboring parenchyma cells, they are typically elongated in the axial direction of the duct and have many vacuoles, a prominent nucleus, higher number of plastids, and high concentration of proteins, lipids, and carbohydrates, which reflect their main function of synthesis and secretion of resin [7, 37]. Also, compared to the plastids of adjacent parenchyma tissue, with well-developed grana, plastids of the resin ducts epithelium have smaller surface-to-volume ratio reflecting their smaller and less developed internal structure. Their external membrane is completely enveloped by a sheet of endoplasmic reticulum, forming a continuous and permanent membrane system [37, 38]. The epithelium and the surrounding subsidiary cells are thin-walled in the mesophyll and cortex, while in the secondary xylem, both cell types have thickened and lignified cell walls, with many of them being nonfunctional and lacking protoplasts [10, 23]. The inner tangential wall of epithelial cells facing the duct lumen is convex, thin, and devoid of plasmodesmata, contrary to other tangential and radial thicker walls with many plasmodesmata facing and linking other epithelial cells and subsidiary cells [7, 39, 40]. Therefore, the hydraulic

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conductivity of the epithelial cell walls is anisotropic, favoring water exchanges with the subsidiary cells and surrounding tissues only. Additionally, this anisotropic distribution of plasmodesmata also allows the transport of precursors of resin components and other metabolites from the phloem through rays to the epithelium of axial and radial resin ducts. In most gymnosperms, the epithelium is short-lived, but in Pinus, presenting commonly a multilayer epithelium, it remains alive and functional for several years [10, 23]. One notable exception of the typical resin duct structure observed in Pinaceae is the apparent absence of subsidiary cells in the mesophyll resin ducts of Tsuga, where the epithelial cells are bordered by large parenchyma cells [23]. After the secretory phase has ended and once transitioned into heartwood, resin ducts of Pinaceae are frequently occluded with tylosoids (protrusions of epithelial cells), e.g., Picea [41] and Pinus [42, 43]. Subsidiary cells are a characteristic feature of the vertical and radial ducts of many species of Pinaceae [9, 23]. These cells are generally characterized by having large vacuoles and a peripheral cytoplasm with fewer organelles than their neighboring epithelial cells [7] and are closely connected not only to the enclosed associated epithelial cells but also to surrounding parenchyma and neighboring tracheids [40]. A complete cylinder of nonfunctional subsidiary cells surrounding the epithelium is found in the secondary xylem only. Although the crushing of nonfunctional subsidiary cells contributes to the expansion of the already developed duct lumen, it is not the way through which resin ducts are formed in gymnosperms. In the primary body, occasional single nonfunctional subsidiary cells are observed, while being totally absent in the secondary phloem [22]. Quite often, this ring of nonfunctional subsidiary cells is bridged at one or more locations by living adjacent parenchyma [9]. The physiological role of this layer of air-filled and nonfunctional subsidiary cells is not clear, but it was considered to have a regulatory function by serving as an air mantle enclosing the epithelium, thus preventing water from entering its osmotically active epithelial cells too rapidly. Therefore, this more or less firm sheath of nonfunctional subsidiary cells surrounding the epithelium may play an important role in the secretion of resin in the secondary vascular system [9, 44, 45]. The size of resin ducts varies considerably between species and even individuals [9, 46]. In many species, the proportion of shorter ducts seems higher in younger parts of the plant body, e.g., Pinus halepensis Mill. [22]. Vertical ducts are commonly three or four times wider than radial ducts [42, 43, 47]. Axial ducts with more than 200 μm in diameter are common in conifers, e.g., Picea, varying between 60 and 300 μm in diameter [48], Pinus caribaea var. hondurensis (Sénécl.) W.H. Barrett & Golfari [40], Pinus canariensis C. Sm. [36], Pinus densiflora Siebold & Zucc. [49], and Pinus elliottii Engelm. [42, 43]. Longer ducts of more than 1 m in length are mostly observed in Pinaceae, e.g., Pinus taeda L. [50]. The distribution of resin ducts within the plant body varies greatly between gymnosperms; however radial (horizontal) ducts are always less frequent than vertical (axial) ducts and occur in few fusiform rays only [42, 43]. Vertical resin duct density is higher in Pinus, varying between 4 and 5 ducts.mm
2 [23, 42, 43], while other conifers present a more dispersed distribution of vertical ducts: Cathaya 0.32; Keteleeria between 0.37 and 2.5; Larix from 2.5 to 2.8; and Picea varying between 2.4 and 3.5 ducts.mm
2

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[23]. The highest radial duct densities are also observed in Pinus, between 0.5 and 2.0 ducts.mm
2, while other conifers present smaller values: Cathaya 0.7; Larix between 0.15 and 0.7; Picea from 0.15 to 0.4; and Pseudotsuga varying between 0.15 and 0.2 ducts.mm
2. In Pinaceae, there are significant differences in the structure and distribution of resin ducts between genera but not within species of the same genus [23]. Although differently distributed within the plant body, in some cases, radial ducts may connect vertical ducts but in the same radial plane only, forming extensive two-dimensional networks occasionally [19, 22, 50]. For example, in Pinus halepensis Mill. there are many two-dimensional networks, each situated in a different radial plane [22]. In Pinus taeda L., axial and radial ducts often are in close proximity and even share epithelial cells, but direct openings between the two are rare [50]. Therefore, the limited size and distribution of vertical ducts and the lack of tangential connections between them may explain the differences in resin composition between different plant organs [51–53]. Usually, traumatic resin ducts are cyst-like or short, of a few centimeters, occurring mostly in tangential series due to their cambial origin. However, in some cases they can develop into large canals, longer and wider than the normal vertical ducts and often scattered and dispersed at distant regions from the wound [9, 18, 19]. Pinus is particularly known for presenting scattered and highly dispersed traumatic resin ducts that resemble and are often mistaken by the normal vertical resin ducts typical of the genus. Not uncommonly, traumatic resin ducts can anastomose tangentially forming a continuous network ramifying throughout the surrounding adjacent parenchyma tissue and, in some cases, extending greatly throughout the plant body forming a barrier zone to seal off the wound or the invading pathogen and protect neighboring healthy tissues [54–57]. The fusion of neighboring traumatic ducts, due to their tangential anastomosis, also contributes to the presence of tangentially wider ducts [19]. However, the character and distribution of traumatic resin ducts vary between genera depending on the type and circumstances of injury [9, 16]. Traumatic resin ducts can also connect to radial resin ducts, thus increasing the extent of their network. Therefore, this organization facilitates the flow of resin not only axially, along the trunk, but also radially towards the surface of bark, thus improving the defense strategies of plants in which they occur [9, 15, 16, 19]. The formation of traumatic resin ducts in gymnosperms is schizogenous, in the same way as that of normal resin ducts, and can occur quite rapidly after injury, e.g., in 2–4 weeks in Picea abies (L.) H. Karst. [16]. Traumatic ducts can originate from cambial xylem mother cells (Pinaceae, except Pseudolarix), phloem mother cells (Agathis, Araucaria), or both tissues (e.g., Cupressaceae) (Table 1) [16, 25]. The vast majority of traumatic resin ducts are surrounded by epithelial cells and subsidiary cells with thickened lignified walls derived from cambial traumatic parenchyma tissue [9, 19].

2.2

Angiosperms

In angiosperms, resin, more commonly mixed with gum and/or essential oils, is produced primarily by woody species and some few herbaceous plants, with higher

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yields being observed in trees of subtropical and tropical climates. Similarly to gymnosperms, most of what is known about the anatomy and physiology of internal secretory systems in angiosperms comes from studies done on species that are the sources of gum resins, oleo-gum-resins, and resins of higher economic value belonging to families Altingiaceae, Anacardiaceae, Apiaceae, Arecaceae, Asparagaceae, Burseraceae, Calophyllaceae, Cistaceae, Clusiaceae, Compositae, Convolvulaceae, Dipterocarpaceae, Leguminosae, Rubiaceae, Rutaceae, Simaroubaceae, Styracaceae, Xanthorrhoeaceae, and Zygophyllaceae (Table 2) [5, 7]. However, the secretion of resin, gum resin, and oleo-gum-resin is quite widely distributed among angiosperms, being mostly observed in dicotyledons compared with monocotyledons, where it occurs in five families only (Table 2). Contrary to gymnosperms, many angiosperms secrete resin in floral structures, to attract pollinators [2, 4, 58–60 ], and fruits [59, 61–64], possibly to facilitate seed dispersal by animals [3]. Hence, it seems that in angiosperms the role of resin goes beyond its typical importance in defense strategies, as commonly attributed in gymnosperms. Possibly, reflecting its broader distribution and physiological function, the development of resin ducts in angiosperms has been observed to occur in three different ways, sometimes in the same plant, but in different tissues, e.g., Anacardiaceae [17, 59, 65, 66] and Compositae [67]. In the same way as in gymnosperms [22, 23], the schizogenous development of resin ducts has been observed in angiosperms both on the primary and secondary vegetative tissues similarly. With very few exceptions, e.g., Ailanthus excelsa Roxb. [73, 74], where resin ducts are found in the xylem, normal vertical and horizontal schizogenous resin ducts of angiosperms occur in the phloem, thus following the vascular bundles, e.g., Anacardiaceae [17, 59, 63, 64, 75], Araliaceae [76], and Burseraceae [77–79]. They can also be found in both xylem and phloem, e.g., Inula helenium L. [67]. Horizontal ducts can be absent [75], but when present, although less numerous than vertical ducts, similarly to what is observed on gymnosperms, they are found in rays, e.g., Boswellia serrata Roxb. ex Colebr. [80] and Lannea coromandelica (Houtt.) Merril [64]. Vertical ducts anastomose frequently in tangential planes [81], thus forming a continuous but irregular network extending throughout the plant body. Normal resin ducts can also be found in the phelloderm, arranged tangentially but irregularly, e.g., Lannea coromandelica (Houtt.) Merril [64]. In many angiosperms, vertical, tangential, and radial ducts form a continuous three-dimensional system connecting leaves, stems, and roots throughout the whole plant body [17, 64]. Considering the environment in which angiosperms presenting this organization of internal secretory structures live and the composition of their exudates, i.e., most frequently resin mixed with gum, the presence of gum resin ducts organized in this way seems advantageous by contributing to an increase of the water-holding capacity of tissues, thus helping plants to prevent desiccation [64, 82]. This type of organization of secretory structures has not been observed in gymnosperms and apparently suggests that the importance of resin in angiosperms goes beyond its commonly associated and most observed defense role. In some species of angiosperms, the column of cells above an existing duct undergoes a few divisions initially with concomitant expansion and splitting of the

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Table 2 Angiosperm genera known to yield gum resins, oleo-gum-resins, and resins [5, 7, 31, 68–72] Family Altingiaceae Anacardiaceae Apiaceae Apocynaceae Araceaea Araliaceae Arecaceaea Asparagaceaea Asphodelaceaea Berberidaceae Boraginaceae Betulaceae Burseraceae Calophyllaceae Cannabaceae Celastraceae Cistaceae Clusiaceae Compositae Connaraceae Convolvulaceae Cornaceae Dipterocarpaceae Gesneriaceae Goodeniaceae Gyrostemonaceae Hypericaceae Humiraceae Koeberliniaceae Lamiaceae Leguminosae

Marcgraviaceae Olacaceae Phrymaceae Pittosporaceae Plumbaginaceae Podostemaceae

Genera Altingia, Liquidambar All genera All genera Cryptostegia, Plumeria Monstera, Philodendron All genera Daemonorops Dracaena Xanthorrhoea Podophyllum Eriodictyon, Halgania All genera All genera Caraipa, Clusiella, Haploclathra, Kayea, Kielmeyera, Mahurea, Mammea, Marila, Mesua, Poeciloneuron Cannabis, Humulus Mortonia, Paxistima Cistus Allanblackia, Calophyllum, Chrysochlamys, Clusia, Garcinia, Montrouziera, Moronobea, Platonia, Symphonia, Tovomita Nearly all genera of Asteroideae, many genera of Mutisioideae, Scorzonera, Scolymus, Tragopogon Connarus, Vismianthus Convolvulus, Ipomoea, Operculina Cornus, Mastixia All genera except Marquesia and Monotes Rhynchoglossum Coopernookia, Goodenia Gyrostemon All genera Humiria Koeberlinia Cyanostegia, Newcastelia, Prostanthera Afzelia, Colophospermum, Copaifera, Cordyla, Daniellia, Detarium, Eperua, Gossweilerodendron, Guibourtia, Hardwickia, Hymenaea, Kingiodendron, Millettia, Myroxylon, Oxystigma, Peltogyne, Piscidia, Prioria, Pterygopodium, Sindora, Tessmania, Trachylobium Marcgravia Coula Diplacus, Mimulus All genera Plumbago Weddellina (continued)

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Table 2 (continued) Family Rhamnaceae Rubiaceae Rutaceae Salicaceae Sapindaceae Scrophulariaceae Simaroubaceae

Solanaceae Styracaceae Thymelaeaceae Xanthorrhoeaceaea Zygophyllaceae a

Genera Reynosia Carphalea, Cinchona, Coffea, Coutarea, Elaeagia, Gardenia Amyris, Cneorum, Cneoridium, Harrisonia, Picrella, Spathelia Populus, Salix Aesculus, Diplopeltis, Dodonaea Bontia, Eremophila, Myoporum Ailanthus, Amaroria, Brucea, Eurycoma, Leitneria, Picrasma, Picrocardia, Picrolemma, Quassia, Samadera, Simaba, Simarouba, Soulamea Anthocercis Styrax Aquilaria, Gonystylus Xanthorrhoea Balanites, Fagonia, Guaiacum, Larrea, Metharme, Pintoa, Plectrocarpa, Porlieria, Sericodes

Monocotyledons

cell walls located at the center of the column. Afterwards, the entire column of cells disintegrates centrifugally, leaving a lumen continuous with the duct beneath. The cells located most outwards radially differentiate into an inner layer of secretory cells surrounded by a layer of thin-walled subsidiary cells. This type of duct development is called schizolysigenous and involves first the division and splitting apart of cell walls and subsequently the breakdown of surrounding tissues both contributing to the development and widening of the duct lumen. It can be observed in vegetative organs [75, 81, 83, 84], but it is mostly found in reproductive organs, such as flowers and fruits [59, 61, 85]. Alternatively, in some species, after schizogenous widening of the duct, the innermost epithelial cells lining the duct lumen disintegrate themselves releasing their contents into it, thus widening the duct further [17]. The lysis of epithelial cells can occur on isolated cells, or it can happen as a programmed systematic process extending throughout the whole duct length. Ducts in which their development involves the lysis of epithelial cells can attain relatively large diameter when mature, as their development requires the destruction of hundreds of cells, which can extend throughout tissues considerably. The disintegration of cells may even extend to neighboring tissues, where a great number of phloem cells, including well-differentiated sieve tube elements and their respective companion cells, disintegrate to form ducts lysigenously. Finally, exclusively lysigenous development of resin ducts, i.e., involving the disintegration of an epithelium from which a lumen originates, has been observed on some angiosperms, e.g., the secondary phloem and phelloderm of Lannea coromandelica (Houtt.) Merril [64] and stem and roots of Trichoscypha spp. [65]. After lysis of the epithelium cells facing the duct lumen, adjacent subsidiary cells differentiate into epithelial cells, thus taking over the secretory function and maintenance of the secretory activity in mature ducts, throughout the development and expansion of the organ in which they are located

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[78, 82, 86]. However, the lysis of epithelial cells and its subsequent substitution by new secreting cells in mature ducts are not always obvious, e.g., Pistacia lentiscus L. [17]. Therefore, some authors consider the development of ducts involving lysigeny as a variation of the more common schizogenous duct development. The epithelial cells of the resin ducts found in angiosperms share many features observed in gymnosperms as well as typical of cells specialized in secretion like those known in glands. Normally, there is one to three layers of elongate tangentially flattened epithelial cells surrounding angiosperm resin and gum resin ducts, sometimes with large vacuoles, e.g., Boswellia serrata Roxb. ex Colebr. [80]. Occasionally, epithelial cells may protrude into the duct lumen forming trabeculae, e.g., Protium heptaphyllum (Aubl.) Marchand. [78]. Although being in some cases thin-walled, e.g., Copaifera langsdorffii Desf. [82], like many gymnosperms, epithelial cells of angiosperm resin ducts are mostly thick-walled and lignified. Lignification can, in some cases, progress further so that the epithelium becomes sclerified with even thicker walls, with large, ramified pits, resembling stone cells. In these cases, the sclerification usually extends to the surrounding subsidiary cells, e.g., Protium heptaphyllum (Aubl.) Marchand. [78] and Delarbrea paradoxa Vieill. [76]. In the same way as in many glands, epithelial cells have a dense granular cytoplasm, rich in plastids, mostly lying near the inner tangential wall facing the duct lumen [80]. Plastids have no well-defined internal structures and are also surrounded by the endoplasmic reticulum, similar to what is observed on conifers [37]. Polysomes; ribosomes (free and attached to endoplasmic reticulum cisternae [75]); numerous mitochondria, sometimes branched [81]; Golgi apparatus [63]; and membranous or multi-vesicular bodies (called paramural bodies) make the rest of the cytoplasm typically found in angiosperm epithelial cells surrounding prominent nuclei. Like in gymnosperms, plasmodesmata, in some cases branched [77], are absent in the tangential wall facing the duct lumen while connecting adjacent epithelial and subsidiary cells radially and tangentially [63]. Exceptionally and contrary to what is known of gymnosperms, plasmalemma invaginations and folds were observed on epithelial cell walls of some angiosperms, e.g., Inula helenium L. [67] and Ailanthus excelsa Roxb. [73], thus evidencing the transfer character of these cells in those species. In Rhus glabra L., these cytoplasmic protrusions connect the duct lumen and originated from plasmodesmata that became disconnected during the shizogenous development of the duct [81]. This suggests that they may participate in the loading of resin into the duct lumen, thus evidencing different mechanisms of resin loading between angiosperms. Parenchyma subsidiary cells, mostly with thick but unlignified cell walls, resemble their enclosed epithelial cells with large vacuoles but, comparatively, with a smaller cytoplasm with fewer organelles [7, 64, 78]. However, subsidiary cells may be absent, e.g., Mangifera indica L. [59] and Rhus glabra L. [83]. Similar to the lysis of epithelial cells that leads to the development of normal resin ducts [17, 64, 65], the development of traumatic resin ducts in angiosperms involves the lysis of the surrounding epithelium after intense and rapid anticlinal and periclinal cell divisions of the cambium derivatives induced by injury. In some cases, this response to trauma can be as fast as some hours after injury [79]. As epithelium lysis

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progresses enlarging the duct lumen, the traumatic parenchyma cells adjacent to the epithelial cells become meristematic and cambiform forming additional epithelial cells that will autolyse eventually, thus further enlarging the resin duct lumen [79, 87, 88]. In this way, and similar to what is observed in conifers [19], traumatic resin ducts of angiosperms can anastomose tangentially so that neighboring traumatic ducts may fuse together, hence contributing to the presence of tangentially wider ducts [78]. In some cases, e.g., Ailanthus excelsa Roxb., the anastomosing and fusing of traumatic resin ducts can be quite extensive due to the total disintegration of axial parenchyma cells of the traumatic tissue, forming a network of tangentially anastomosing cavities that appear as a vertically elongated continuous system only interrupted by multiseriate rays that remain intact [87]. The main function of traumatic resin ducts in angiosperms is identical to that observed in gymnosperms, that is, the formation of barrier zones that protect healthy tissues isolating them from injury or pathogen attack [56]. Compared to gymnosperms, there are not many studies presenting quantitative data on the secretory ducts of angiosperms. However, like in gymnosperms, high variability on the dimensions of secretory ducts between species and individuals is also recognized [70]. For example, the size of normal vertical ducts is considered useful in distinguishing some species belonging to Dipterocarpaceae [89]. Resin ducts of 10 to 300 μm in diameter have been observed on Inula helenium L. [67], while they can be as wide as 250 μm in Schinus terebinthifolius Raddi [59]. Species like Boswellia serrata Roxb. ex Colebr., with ducts up to 100 μm wide [90], and Delarbrea paradoxa Vieill., with ducts up to 50 μm wide [76], seem to present narrower secretory structures. Resin ducts of Boswellia papyrifera (Delile ex Caill.) Hochst. have been observed to vary between 20 and 230 μm in diameter while occurring with a density between 0.3 and 1.51 duct.mm- 2 [91, 92]. The fusion and anastomosing of traumatic ducts leading to extensive internal intercellular spaces like in the extreme case of Ailanthus excelsa Roxb. [87] further complicate the distinction and measurement of the dimensions of specific ducts.

3

Synthesis and Secretion into Ducts

The very first studies carried on the ultrastructure of secretory systems of plants using electron microscopy suggested that different organelles are involved in the synthesis of different resin components derived from photosynthetically produced carbohydrates. First focusing on conifers and less so on angiosperms, two prominent organelles, the plastid and the endoplasmic reticulum, stood out distinctively in the epithelium secretory cells [37, 75, 77, 93–95]. Complemented with biochemical analysis, it was then possible to show that the synthesis of monoterpenes, the major components of the volatile fraction of the resin in conifers [5], was initiated in the plastids but completed in the cytosol and endoplasmic reticulum [96–98]. The synthesis of sesquiterpenes, the dominant components of the volatile fraction of most angiosperm resins, e.g., Dipterocarpaceae and Leguminosae, occurs in the endoplasmic reticulum [99, 100]. Diterpenes, the dominant components of the

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nonvolatile fraction in the resin of Leguminosae, are produced in plastids, while the synthesis of triterpenes, which dominate the nonvolatile fraction of the resin of Anacardiaceae, Burseraceae, and Dipterocarpaceae, involves both the cytosol and the endoplasmic reticulum [5]. Compared with terpenes, the synthesis of other components of resin, e.g., phenolic compounds, is less clear as well as which organelles are involved. After synthesis in both plastids and endoplasmic reticulum, terpenoid resin components are then transported across the cytosol enclosed in vesicles originated from the continuous membranous system formed by these organelles, which is specific to resin duct epithelium [37, 38]. The resin-containing vesicles will then fuse with the plasmalemma releasing their contents by exocytosis into the space between the plasmalemma and the tangential wall facing the duct lumen, in what is called by granulocrine secretion. The loose and dispersed microfibrils of the tangential cell wall facilitate the transport of resin through it into the duct lumen [63, 73, 101, 102]. The simple diffusion of these components through the cell wall is then suggested as the main process of resin loading into the duct lumen [39, 73, 95, 103]. In some cases, dense vesicular bodies between the plasma membrane and the cell wall lining the duct lumen have also been observed. Considering the almost ubiquitous absence of plasmodesmata in the tangential wall lining the duct lumen, it seems that, in these cases, resin loading into the duct is made through a relatively porous cell wall via vesicles, e.g., Commiphora mukul (Hook. ex Stocks) Engl. [62]. In Rhus glabra L., the presence of cytoplasmic protrusions, derived from plasmodesmata during shizogenous duct development, connecting the duct lumen, thus contributing to a higher permeability of the cell wall, may facilitate even further the transport of resin components through it [81]. In some angiosperms, however, plasmalemma folds and invaginations in the tangential wall facing the duct lumen, similar to those found in transfer cells, suggest that, at least, some components of resin may be actively loaded into the lumen, in what is called eccrine secretion [7, 67, 73, 104]. Gum, one of the main components of gum resins, is a mixture of complex chains of hydrophilic polysaccharides, mostly known from angiosperms, e.g., Ailanthus excelsa Roxb., Cistus ladanifer L., and Dorema ammoniacum D. Don. [7]. Therefore, gum resins present characteristics of both resins and gums, including being hydrophilic. Some gymnosperms also produce gum resins, e.g., Araucaria and Podocarpus [105, 106]. The gummous components of gum resins result from the breakdown of celluloses and hemicelluloses in the outer epithelium cell wall layers, lining the duct lumen, into unorganized amorphous substances that are loaded into the duct lumen, in what is called gummosis. Simultaneously, new material is added to the inner surface of the cell wall [77, 81]. The Golgi apparatus is the main site of synthesis, transport, and secretion of polysaccharides in higher plants. After synthesis in the Golgi apparatus cisternae, gum is then transported by Golgi-mediated smooth vesicles that deposit it in the space between the plasmalemma and the cell wall by exocytosis [61, 63, 73, 75, 77, 102, 107–110]. Gummosis can occur as a normal physiological process or a pathological response of certain plant species to bacterial or fungal infection [111]. Combined with the special organization of gum resin ducts, the hydrophilic nature of the gummous components of gum resin seems

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thus to help angiosperms to withstand water shortages, induced by adverse environments, by increasing the water-holding capacity of tissues, and prevent desiccation, as suggested by the increase of gum yield under those conditions [64, 82]. Although the secretion of resin in gymnosperms is mostly considered of granulocrinous nature, resin secretion in angiosperms, perhaps due to its more diverse composition, involves in some cases eccrine, granulocrine, and holocrine processes, e.g., Anacardium occidentale L. [63], Commiphora mukul (Hook. ex Stocks) Engl. [62], Copaifera langsdorffii Desf. [82], Lannea coromandelica (Houtt.) Merr. [64], Pistacia lentiscus L. [17], and Protium heptaphyllum (Aubl.) Marchand [78].

4

A Hydrodynamic Model

The first mathematical model of resin flow was presented by Schopmeyer et al. [112] by applying the Hagen-Poiseuille equation to the resin flow of the duct system of Pinus elliottii Engelm and comparing it to pressure and flow measurements of 12 trees. It was very simple, for example, it did not consider resin loading into the duct lumen. However, despite its limitations, it showed that the specific resistance of the duct system, given by the product of the reciprocal of the viscosity of oleoresin by the number and cross-sectional area of the cut ducts, for when N ducts were cut and exposed through a tangential wound surface, accounted for 83% of the variation in flow rate between the 12 specimens studied. Also, by observing a correlation between the yield accumulated in the first 24 h and the annual yield of 25 trees, Schopmeyer et al. [112] suggested that the specific resin duct resistance, dependent on these physiological and anatomical parameters, could also correlate to the annual yield, the most used indicator of resin yielding capacity. Perhaps, the biggest merit of the work of Schopmeyer et al. [112] was that it showed that a set of measurable physiological and anatomical parameters could be a useful criterion for the selection of parent specimens, which could be then used in breeding programs to produce progeny with high oleoresin yielding capacity. A more realistic model based on what is currently known about the resin ducts structure and physiology, namely, the loading of resin, was suggested by Cabrita [113].

4.1

Model Assumptions and Governing Equations

Considering the conceptual model of resin loading first proposed by Büsgen and Münch [114], which partially explained the buildup of pressure in the duct lumen, the approximate cylindrical geometry of resin ducts systems in plants, and the granulocrine loading of resin, Cabrita [113] applied the unsteady Stokes equation to describe the dynamics of resin flow within ducts: !

@u 1 ! ¼
∇ p þ ν∇2 u ρ @t

ð1Þ

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Resin, behaving like a homogeneous Newtonian fluid, i.e., of constant kine! matic viscosity ν and density ρ, flows with velocity u under pressure p out of a duct, of constant radius R, open at the end of its length L through a wound. Once in contact with the air outside, resin solidifies, decreasing its flow with time, eventually closing the wound. The viscous nature of resin [112, 115] combined with the long and narrow structure of resin ducts [23, 50] make flow inside them very slow. For this reason, it is thus classified as Stokes flow or creeping flow, where the advective inertial forces are small compared with the viscous forces. Compared to previous models [112, 114], the model of Cabrita [113] is more realistic in considering the physiology and the anatomy of resin secretory systems of plants. In this respect, the resin duct is considered surrounded by an epithelium in which pressure, pe, constant with distance, follows the diel cycle of the water status of the surrounding tissues, i.e., with a period τ of 1 day, fluctuating around pe0 with amplitude A ( Qieketou (Table 2).

Table 2 Annual yield of raw lacquer in the four cultivated T. vernicifluum varieties for each cut Varieties Average yield of one time tapping (g) Average annual yield (g)

Dahongpao 8.95  0.54

Gaobachi 8.76  0.43

Huoyanzi 3.44  0.32

Qieketou 1.90  0.33

223.79  4.75

219.04  5.34

85.89  3.18

47.38  3.20

Five trees of each variety were used in the experiment.

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Discussion

The varieties of cultivated plants are usually distinguished by their external features, harvest quality, and yield [35]. In previous studies, tree crown sizes, characteristics of reproductive organs, and morphology of different organs have been considered as the index to differentiated lacquer tree varieties. Recently, amplified fragment length polymorphism (AFLP) molecular markers also have been employed as novel indicators for identifying cultivated high-yield lacquer tree varieties. For raw lacquer production of T. vernicifluum, the yield and quality are considered as the main economic characters, in previous studies, investigators mainly focused on economic characters for cultivated lacquer tree varieties [16]. These studies were carried out separately, and thus information on the yield-quality relationship is still lacking. To resolve this drawback, comparative anatomical approach on bark structures with economic characters and phytochemistry were employed in the present study. Our study revealed that raw lacquer tree varieties were similar in bark structure but different in the quantity of resin canals per unit area, diameters of resin canals, thickness of bark, and other aspects. The differences were steady among these varieties and may be used as an anatomic classification index for lacquer tree varieties, which were similar to the H. brasiliensis research findings [3]. As another cave secretion, the commercial resin was tapped from pine trees with the similar method applied for raw lacquer harvest; the pine trees’ oleoresin is synthesized and contained in the mature laticifers [36]. Laticifers are distributed within all the pine organs, and oleoresin is produced in the secondary xylem. The previous study showed that the number, diameter, depth, and density of inner and outer resin canal measurements differed significantly among pine species [37]. Therefore, these characters should serve as the important index for distinguishing cultivated species with the aim to use the combination of the biological and economic characters in T. vernicifluum variety classification. Resin canals are mainly formed and extended in the conducting phloem, but most mature resin canals occur in the nonconducting phloem [21]. The microstructure of lacquer tree resin canals showed that raw lacquer is synthesized in the secretory cells surrounded by canals. The raw lacquer deposited in caves of resin canals through cell walls by plasmodesmata or diffusion penetration. Furthermore, the surrounding tissues of resin canals participated in the synthesis of raw lacquer precursor substances [25]. The differences in the quantity and diameter size of resin canals with bark thickness of lacquer tree varieties can affect the ability of composing and storing raw lacquer. Sieve tubes are the channels that transport organic compounds inside the plants. Those compounds may serve as precursors for raw lacquer components, and the average diameter of sieve tubes may affect the ability to produce/transport raw lacquer. Investigation into the relevance between bark structure and raw lacquer output of lacquer trees showed positive correlations in amount of resin canals, average diameter of resin canals and tubes, and the average of bark thickness with annual raw lacquer yields. For example, the Dahongpao variety had the highest annual raw lacquer yield with its average diameter of resin canals and average of bark thickness. Thus, the

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quantity and the average diameter of resin canals and tubes and the average of bark thickness were the key factors for the raw lacquer yield. These characteristics should be considered as the anatomical index for the selection of lacquer tree varieties. Resin canals are specialized cells containing latex and have been found to take part in the biosynthesis of specialized metabolites in many plants, such as H. brasiliensis [22]. In the present work, structural guidance of phloem and resin canals were analyzed jointly. The results showed that those structural factors had the relationship with the annual raw lacquer yields. These findings should be considered for raw lacquer production and for further investigation of lacquer tree varieties in support of appraisal and selection.

4

Ultrastructural Study of the Development of Resin Canals and Lacquer Secretion in T. vernicifluum

The secretory products of plants have a variety of ingredients like gum, resin, latex, essential oil, and alkaloid; those products are regarded as one of the most significant groups of secondary metabolites in plant defense; these substances work as defense mechanisms against insects, herbivores, and pathogens [38, 39]. The study on the secretion mechanism of secretory product reveals that there are two possible ways of secretion. One way is the active molecules synthesized by secretory cells that are secreted to the outside through cell walls. The other is that vesicles are formed in secretory cells. The membranes of these vesicles can be fused with plasma membrane, and the secretory product is secreted in this way [40]. The storage structure for raw lacquer is the resin canals which are distributed in phloem of vascular bundles in all organs of lacquer trees; the resin canal structure and development in T. vernicifluum have been described in detail [16]. There are most resin canals in secondary phloem of stem, so this is the major part for raw lacquer production [21]. Although much work has provided morphological, developmental, and ultrastructural details of the resin canals, but apparently nothing is known about its ontogeny or details of raw lacquer secret. Therefore, the objective of this study was to perform a detailed analysis of the anatomy and ultrastructure of the secretory cells, to investigate the structural characteristics and ontogeny of secretory cells and the secretion of raw lacquer synthesizer and secretory.

4.1

Ultrastructure of Secretory Cells and the Development of Resin Canals

During development of resin canals in phloem, the initial cells of resin canals have a series of changes in morphology and ultrastructure. Before the formation of resin canals, 5–8 initial cells gathered in a shape of lotus throne (Fig. 6a). The initial cells of resin canals were rich in protoplasm. Compared with phloem elements in surrounding tissue, the initial cells of resin canals have more Golgi bodies and plastids. Most of the plastids were round or oval. They were large and even distributed in

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Fig. 6 Ultrastructure of secretory cells and raw lacquer secretion from secretory cells in resin canals. (a) Original cells of a resin canal arranged in a rosette, bar = 0.48 μm; (b) Endoplasmic reticulum, mitochondria, and other organelles in the original cell, bar = 0.5 μm; (c) Intercellular space appearing between the original cells of resin canals, the arrow showed the initial stage of resin canal, bar = 0.75 μm; (d) Resin canals in slit shape at initial stage, secretory cells have secretion

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cytoplasm. There were also other organelles like endoplasmic reticulum and mitochondria. There was a conspicuous cell nucleus and a lot of small vacuoles in the cell (Fig. 6b). At the first stage of resin canals formation, intercellular space appeared at middle lamella of initial cells. The combining sites of cell walls separated and formed lumen gap as a narrow gap at first (Fig. 6c). Afterward, the intercellular space extended toward two sides along the middle lamella and develops into 5–8 cells which formed cavities of different shapes. In this process, the surrounding secretory cells got mature, the plastid in these cells was increasing, and the types of mitochondria, Golgi bodies, and endoplasmic reticulum were also increasing (Fig. 6d). The endoplasmic reticulum was developed with a large number of flocculent materials (Fig. 6e).

4.2

Raw Lacquer Secretion from Secretory Cells in Resin Canals

With development of resin canals and secretory cells, there were more organelles in mature secretory cells like plastid, Golgi bodies, and mitochondria. It contained a lot of the endoplasmic reticulum in secretory cells (Fig. 6f). The cell nucleus was very distinct, and there was no central vacuole. At this the cells have a strong secretion function. In cytoplasm, there were a lot of flocculent materials covered by monofilm (Fig. 6g). These secretory granules covered by monofilm were formed by Golgi bodies and then fused and transported to near plasma membrane. Later, the vacuolar membrane and secretory cells fused together toward the plasma membrane of resin canals lumen. The vesicles which are filled with flocculent materials were released to periplasmic space (Fig. 6h), and then the electron dense inclusion was spread and released to resin canals lumen through permeating the cell wall (Fig. 6i). The flocculent materials in resin canals lumen would then fused, integrated, and formed raw lacquer; the lacquer was abundant in the lumen (Fig. 6j).

4.3

Aging and Disintegration of Secretory Cells in Resin Canals

In the region of nonconducting phloem at secondary phloem, the secretory cells of resin canals gradually lost the function of synthesizing and secreting lacquer. ä Fig. 6 (continued) function, bar = 0.4 μm; (e) Secreting cells gradually mature, containing a variety of organelles, plastids, mitochondria, Golgi apparatus, and endoplasmic reticulum, bar = 0.15 μm; (f) Endoplasmic reticulum in a secretory cell, bar = 0.1 μm; (g) Mature secretory cells, vesicle fusion appeared near the plasma membrane, bar = 0.25 μm; (h) Vesicle fusing with the plasma membrane and electron dense substances being released into periplasmic space, bar = 0.15 μm; (i) Raw lacquer passing through the cell wall to the lumen, bar = 0.15 μm; (j) Flocculent materials formed raw lacquer in resin canals lumen; (k): Senescence cell without secretory function, bar = 0.75 μm; (l) Senescence secretory cell, bar = 0.45 μm. Notes: C, cambium; Com, companion cell; CP, conducting phloem; CW, cell wall; PE, periderm; Pla, plasmodesma; PR, phloem ray; RL, raw lacquer; SC, stone cells; SE, secretory cells; SH, sheath cells; VES vesicles; Star«, resin canals

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Senescent secretory cells were hollowed cells and lacking certain cellular components (nucleus, Golgi bodies, ribosomes, and dictyosomes); there were few flocculent materials covered by monofilm in cytoplasm. Large central vacuoles were formed in secretory cells. Organelles were significantly reduced. There were only few plastids and mature starch grain (Fig. 6k). In the region of nonconducting phloem around periderm, the secretory cells of resin canals aged and disintegrated. The aged secretory cells vacuolize and shrink in size. There was few starch grain in the cells, but the organelles like endoplasmic reticulum and Golgi bodies disappeared, and few inclusions in the cells were released to resin canals lumen through cell walls (Fig. 6l). Eventually, the cell wall got fibering and disintegrated (Fig. 7a).

4.4

Ultrastructure of Sheath Cells and Phloem Elements

The sheath cells were located at the outside of secretory cells; when the resin canals formed, the initial cells of sheath cells were rich in protoplasm and have a lot of plasmodesma with secretory cells (Fig. 7b). With the formation of secretory cells, sheath cells extend gradually and cover the secretory cells flatly. At this time, sheath cells have thick cell walls, electron density which was lower than secretory cells and distinct cell nucleus. There were a variety of organelles in cytoplasm like plastid, endoplasmic reticulum, and Golgi bodies. Different from secretory cells, they have large central vacuoles (Fig. 7c). Light microscope (LM) and transmission electron microscope (TEM) research on the secondary phloem of lacquer tree showed that the secondary phloem was composed of sieve tube, companion cell, parenchymal cell, stone cell, and phloem ray. The sieve tubes in the functioning phloem possess a central large vacuole, clear nucleus, and organelles such as plastids and endoplasmic reticulum (Fig. 7d). In the region of nonconducting phloem, the secretory cells lost their function of synthesizing and secreting lacquer; at the same time, the sieve tubes were squeezed to shrinkage and deformation by parenchymal cells; they displayed quite irregular shape, narrow intracellular space, and loss of nuclei, with intracellular P-protein and obvious thickening wall (Fig. 7e). Lignin deposition in concentric ring and formed mature stone cells finally (Fig. 7f).

4.5

Discussion

Secretory structures were found in about 12,500 plants of 22 families; secretory structures and ontogeny formation have been an important source of evidence for establishing the evolutionary patterns of characters and for re-examining phylogenetic relationships in a number of groups, particularly for orchids [1]. Among them, there are more tropical plants than temperate plants [2]. Structure and development of sumac phloem have the characteristics of temperate-zone trees. The resin canals exist in the molecules of phloem. They are derived from cambial cells [17]. The secretory

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Fig. 7 Ultrastructure of aging and disintegration of secretory cells, sheath cells, and phloem elements. (a) Cell wall fibrotic of the senescence secreting cells, bar = 1 μm; (b) Original sheath cells show the plasmodesma, bar = 0.65 μm; (c) Ensheathing cells gradually elongated with a significant nuclei and variety of organelles, bar = 0.65 μm; (d) Sieve tubes and companion cells, bar = 0.6 μm; (e) Deformation sieve tubes in nonconducting phloem; bar = 0.6 μm; (f) Lignin deposition in concentric ring and formed stone cell, bar = 0.5 μm. Notes: C, cambium; Com, companion cell; CP, conducting phloem; CW, cell wall; PE, periderm; Pla, plasmodesma; PR, phloem ray; RL, raw lacquer; SC, stone cells; SE, secretory cells; SH, sheath cells; VE, vesicles; Star«, resin canals

structures are generally formed in schizogenic, lysigenic, and schizolysigenous ways [1]. Secretory structures have been investigated in several genera of Anacardiaceae. Historically, Venning (1948) was the first author who described these secretory structures in Anacardiaceae [20]. Afterward, much attention has been dedicated to their original location and their further differentiation. The ontogeny and formation of the secretory canals have been considered as an important character in classification of Anacardiaceae. It was found in structures of many Anacardiaceae plants like Pistacia chinensis, Rhus typhina, Rhus potaninii, Semecarpus anacardium, Trichoscypha, Mangifera indica, and Schinus terebinthifolius; all these plants have schizogenic resin canals. This is believed to be a common characteristic of Anacardiaceae in anatomical structure [20]. Through microstructure and ultrastructure observation of secretory cells in lacquer trees, our research found that the resin canals of lacquer trees are also typical schizogenic. In the previous study on Ainaceae plants, Anacardiaceae, Asteraceae, Hypericaceae, and Umbelliferae, it shows that the secretory cells have the following

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characteristics: the cytoplasm is rich in plastid, and part of the plastid is covered by endoplasmic reticulum. The large amount of flocculent materials in cytoplasm is the most typical characteristic in ultrastructure of secretory cells; there is osmiophilia in various organelles in cytoplasm [40]. The observation to ultrastructure of secretory cells reveals that there are a lot of endoplasmic reticulum, Golgi bodies, and plastid in secretory cells with active secretion function. This is similar to the ultrastructure of secretory cells in various plants [38, 41]. Research on secretion structures of plants indicates that the endoplasmic reticulum (ER) and Golgi bodies carried out a multiplicity of functions in the synthesis and secretion of product. The endoplasmic reticulum and (or) Golgi bodies in active secretory cells produce a lot of cisternae, and these cisternae form vesicae, membranes of the ER, are functionally connected to all membranes of the secretory and endocytic pathways via vesicular transport. Afterwards, the vesicae membrane and plasma membrane fuse and release secretion product [42–45]. In the secretory cells with secretion function, there are a lot of vesicae of osmiophilic secreta. This is one of major features that secretory cells are different from other cells in phloem. These vesicae contain osmiophilic substance and gradually fuse with plasma membrane. In this way, they transport the secretion product through membrane and spread it to the resin canals lumen and form lacquer. The major physiological function of phloem is to transport many sorts of nutrients, including saccharides, amino acid, micromolecular nutrients, lipid, and hormone. It plays a leading role in the growth process of the plants [46]. Long-distance signal transmission of plants is mainly carried out through the phloem [47, 48]. In addition, the molecules in phloem also participate in synthesis of various secondary metabolites like latex, alkaloid, and tannin [49–51]. Our results showed that the secretory cells and sheath cells of resin canals and other molecules in conducting phloem of lacquer trees got mature at the same time. The previous study on development of secretory structure and molecules in phloem of Papaver somniferum indicated that before secretory structures get mature, alkaloid of Papaver somniferum has always been synthesized in molecules of phloem. Some enzymes synthesized by alkaloid also exist in molecules of phloem [50]. The study also finds that the molecules in phloem around resin canals are high in flocculent materials. Therefore, these molecules serve as precursor substance in synthesis of lacquer. When the initial cells of latex cells form the rudiment of slot-shaped resin canals lumen, there are already vesicles in initial cells. In previous research on the development of secretory lumen in Copaifera langsdorffii fruits, it shows that the formation of secreta is earlier than the development of secretory cavity [38].

5

Conclusions

The vertical system of the lacquer tree secondary phloem consists of sieve tubes, companion cells, parenchyma cells, stone cells, and resin canals. The phloem structures were similar with different varieties but displayed differences in secondary phloem thickness and raw lacquer yields. The diameter of resin canals and the bark thickness were positively correlated with the raw lacquer yields. The resin canal was

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formed by the initial cells of secretory cells through schizogeny. Secretory granules with monofilm in secretory cells were shaped by Golgi bodies and transported to the plasma membrane and fused with the membrane, and the interior flocculent materials were released and spread to the resin canals through the cell walls and formed raw lacquer. Acknowledgments This study was supported by the National Natural Science Foundation of China (Grant No.: 31300157, 31401435).

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Part II Production of Secondary Metabolites in Shoot Cultures

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Shoot Organogenesis, Genetic Stability, and Secondary Metabolite Production of Micropropagated Digitalis purpurea Elizabeth Kairuz, Naivy Pérez-Alonso, Geert Angenon, Elio Jiménez, and Borys Chong-Pérez

Contents 1 2 3 4

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Digitalis purpurea L. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Phytochemistry and Medicinal Uses . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . In Vitro Culture of Digitalis purpurea and Secondary Metabolite Production . . . . . . . . . . . . 4.1 Indirect Organogenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2 Direct Organogenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5 Genetic Stability . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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E. Kairuz Departamento de Biología, Facultad de Ciencias Agropecuarias, Universidad Central Marta Abreu de Las Villas (UCLV), Santa Clara, Cuba Instituto de Biotecnología de las Plantas (IBP), Universidad Central Marta Abreu de Las Villas, Santa Clara, Cuba Laboratory of Plant Genetics, Vrije Universiteit Brussel (VUB), Brussels, Belgium e-mail: [email protected] N. Pérez-Alonso Instituto de Biotecnología de las Plantas (IBP), Universidad Central Marta Abreu de Las Villas, Santa Clara, Cuba e-mail: [email protected] G. Angenon Laboratory of Plant Genetics, Vrije Universiteit Brussel (VUB), Brussels, Belgium e-mail: [email protected] E. Jiménez Florida Crystals Corp, Belle Glade, FL, USA e-mail: [email protected] B. Chong-Pérez (*) Sociedad de Investigación y Servicios BioTECNOS Ltda, San Javier, Chile e-mail: [email protected] © Springer Nature Switzerland AG 2021 K. G. Ramawat et al. (eds.), Plant Cell and Tissue Differentiation and Secondary Metabolites, Reference Series in Phytochemistry, https://doi.org/10.1007/978-3-030-30185-9_16

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6 Biotechnological Approaches for Biomass and Cardenolide Production . . . . . . . . . . . . . . . . . . 180 7 Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 182 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 183

Abstract

Digitalis purpurea L. is a cardenolide-producing medicinal and ornamental plant. Cardenolides, like digoxin, are commonly used to treat congestive heart failure, cardiac arrhythmia, and atrial fibrillation. More recently, its cytotoxic activity on several types of cancer and antiviral effect have been confirmed. As chemical synthesis is not viable, D. purpurea plants are one of the major sources of cardenolides for the pharmaceutical industry. However, cardenolide content is highly variable under natural conditions. Therefore, D. purpurea in vitro culture has been a focus of research since the second half of the twentieth century. This chapter is a compendium of these reports with emphasis on the effects of morphogenesis, culture conditions, and in vitro culture medium composition on cardenolide content. Besides, some studies on the genetic stability of Digitalis plants are summarized. Finally, we describe the biotechnological approaches reported so far to obtain a higher yield of cardenolides in vitro, such as elicitation and metabolic engineering, both recognized as promising strategies. Keywords

Cardenolides · Common foxglove · Digitoxin · Digoxin · Growth regulators · Organogenesis · RAPD · Temporary immersion systems · Elicitation Abbreviations

2,4-D 2iP ABA BA BTOA DW FW GA3 IAA LED LS medium MS medium NAA PEG RAPD RuBisCO TDZ TIS

2,4-Dichlorophenoxyacetic acid N6-[2-Isopentenyl] adenine Abscisic acid Benzyladenine 2-Benzothiazole-oxyacetic acid Dry weight Fresh weight Gibberellic acid Indoleacetic acid Light-emitting diode Linsmaier and Skoog medium Murashige and Skoog medium Naphthaleneacetic acid Polyethylene glycol Random amplified polymorphic DNA Ribulose-1,5-bisphosphate carboxylase/oxygenase Thidiazuron Temporary immersion system

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Introduction

To deal with high disease incidence, there is a continuous search for new natural compounds that can serve as drugs and for more efficient production systems for known bioactive compounds [1]. Leaf extracts of Digitalis species have been used to treat heart diseases for centuries [2]. Moreover, Digitalis-derived cardenolides have been reported to have antiviral activity, a strong cytostatic effect against several kinds of tumors, and potential beneficial impact on cystic fibrosis [3]. Since 1950, cardiovascular diseases have been the leading cause of death in humans (17.9 million deaths per year, 44% of all noncommunicable diseases), followed by cancer (9.0 million deaths per year, 22%) [4]. However, more recently, an increase due to cancer has been recorded in some populations [5]. Therefore, bioactive compounds to treat these diseases are highly demanded worldwide. As chemical synthesis of cardenolides is not viable, Digitalis and Isoplexis plants are the only sources of cardenolides, with Digitalis purpurea L. and Digitalis lanata L. as the major sources for the pharmaceutical industry. Nonetheless, the intensive cultivation is constrained by several factors including the biennial cycle of the plant, biotic and abiotic stress conditions, and germination frequency. Besides, cardenolide content and accumulation are influenced by climate, soil conditions, and genotype [6, 7]. Hence, several biotechnological methods (organogenesis, somatic embryogenesis, bioreactor culture, biosynthetic precursor feeding, elicitors, genetic transformation) have been applied to produce biomass and cardenolides in vitro. However, failed attempts to obtain commercial production from in vitro culture emphasize the need for new alternatives like the combination of metabolic engineering with other biotechnological strategies [2]. This review aims to summarize the main studies on shoot organogenesis of D. purpurea, as the leading propagation method. Also, genetic stability and secondary metabolite production of micropropagated D. purpurea are discussed. This chapter includes a compilation of the reports on the in vitro culture of D. purpurea and the effects of morphogenesis, light, growth regulators, and medium composition on the accumulation of cardenolides. Besides, studies on genetic stability determination in the Digitalis genus are reviewed, and some promising biotechnological approaches to enhance production of cardenolides in vitro, such as elicitation and metabolic engineering, are described.

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Digitalis purpurea L.

Digitalis purpurea L. (Plantaginaceae), also known as common foxglove, purple foxglove, or lady’s glove, is one of the current 23 species within the Digitalis genus [3]. This plant is a biennial or short-lived perennial (when flowering is delayed) herb native to Europe (Portugal, Spain, Germany, Norway, Ireland, and Great Britain). However, because of its medicinal and ornamental values, it was introduced in other temperate areas like South America, West Coast of the USA, New Zealand, and Canada [3, 7].

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D. purpurea can be found in natural and human-made areas like pastures and roadsides. The germination is inhibited by low-light conditions and takes place in spring. Leaves are alternate, petiolate with oval-lanceolate limb and dentate margin, and densely hairy mainly by the underside, gathered in a rosette at ground level in the first year. The flowering stem develops in the second year during the summer after vernalization in winter, but this process can be delayed for several years. Plants produce one or a few racemose inflorescences around 1.0 m height on average, with characteristic purple flowers [6]. This was the first species of the genus Digitalis described as a medicinal plant more than 200 years ago by William Withering [8]. However, due to the presence of cardenolides, it is considered a poisonous plant to humans and several animals.

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Phytochemistry and Medicinal Uses

D. purpurea is widely known for the production of cardenolides in leaves, tender stems, flowers, and seeds. These steroidal cardiotonic glucosides inhibit the Na+/K+ATPase causing a positive inotropic effect in the heart that increases myocardial contraction and regulates the heart rate [2]. Additionally, Digitalis drugs have a stimulating effect on the parasympathetic nervous system. This plant can produce more than 30 kinds of cardenolides, like digoxin, digitoxin, digitoxigenin, gitoxin, lanatoside C, and purpureaglycosides A and B, among others [3, 6]. Digitoxin, digoxin, and other cardenolides can be interconverted, and their content and accumulation are highly modulated by biotic and abiotic conditions [6]. Moreover, other pharmacological effects have been demonstrated for cardenolides like apoptotic effects in tumor cells at low concentrations. In addition, digitoxin and other cardenolides can be potentially used in the treatment of cystic fibrosis and viral infections [3]. On the other hand, a steroid called digoxigenin found exclusively in D. purpurea and D. lanata plants is used as a molecular probe to detect DNA or RNA. Besides, these plants can contain an arsenal of metabolites like digitanols (Δ5pregnenes), rare sterols, steroid saponins, anthraquinones, phenylethanoids, and flavonoids [6]. Moreover, polysaccharides (arabinogalactans, glucomannans, arabinoxyloglucan, acidic xylans) were also extracted from D. purpurea leaves.

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In Vitro Culture of Digitalis purpurea and Secondary Metabolite Production

In vitro culture application to produce secondary metabolites has several advantages in comparison to traditional cultivation methods. These techniques allow providing a continuous and uniform source of bioactive compounds using healthy selected genotypes as starting material. The culture is performed in small spaces and under controlled conditions avoiding seasonal effects. Besides, culture conditions and

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medium compositions could be optimized to achieve a higher multiplication rate and to enhance the production of specific metabolites. Furthermore, the main issue related to plant extract application is avoided due to the production of a homogeneous extract. Therefore, the Digitalis genus in vitro culture has been widely studied to obtain cardenolides [7]. Particularly in D. purpurea, the earliest report was focused on determining whether or not tissue cultures of cardiac glycoside-containing plants can biosynthesize these compounds [9]. In this work, seedlings and roots from Digitalis plants were sterilized and transferred aseptically to a nutrient medium with coconut milk to induce callus formation. The addition of casein hydrolyzate inhibited the growth of seedlings in all medium compositions. In contrast, seedlings were able to grow regardless of the mineral solution composition. The effects of the auxins 2-benzothiazole-oxyacetic acid (BTOA), 2,4-dichlorophenoxyacetic acid (2,4-D), and naphthaleneacetic acid (NAA) in callus formation were evaluated. The best callus formation from seedlings was achieved with combinations of BTOA (1 or 5 ppm) and 2,4-D (1 ppm). Moreover, callus tissue formed from roots was occasionally observed in the culture medium with all the auxins evaluated or their combinations [9]. In the following publications of this group, suspension cultures were established from callus tissue [10] and were incubated with selected substrates to evaluate the production of cardiac glycosides [11], but D. lanata and D. purpurea callus suspension cultures did not produce glycosides. A few years later, Furuya et al. [12] examined the biotransformation of progesterone by suspension cultures of D. purpurea. However, only 5α-compounds were found in metabolic products from the callus, whereas cardenolides (i.e., 5β-configured compounds) were never obtained. These results were further confirmed by these authors in normal and habituated callus, leading to the conclusion that the cardenolide biosynthetic pathway is inhibited in undifferentiated Digitalis cells [13, 14]. Finally, during the purification and characterization of sterol:UDPG glucosyltransferase in cell culture of D. purpurea, they observed that shoot differentiation was required for cardenolide biosynthesis [15]. Hence, shoot organogenesis is essential for cardenolide production in Digitalis.

4.1

Indirect Organogenesis

The regeneration of D. purpurea plants via indirect organogenesis has been described in several reports (reviewed by [7]). Plant regeneration from anthers was obtained in 1975 by Corduan and Spix [16]. These authors were able to obtain haploid callus when the anthers showed a dark-brown to a black color and were cultured on Nitsch and Nitsch medium [17]. Another important factor was the presence of 2,4-D (5 mg l 1) in the culture medium during callus induction, but not for the regeneration medium. Moreover, plant regeneration was highly susceptible to light conditions and kinetin effects. As a final result, a high frequency of haploid, diploid, and tetraploid plants was regenerated [16].

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From 1980 onward a series of crucial papers by Hagimori and co-workers was published [18–22]. These works aimed to study the production of cardenolides by plant tissue culture. First, digitoxin and digoxin contents were determined in calli and shoot-producing callus of several Digitalis species. Callus was induced from seedlings and leaf discs in MS medium [23] at 28  C with several combinations of growth regulators. The content of digitoxin and digoxin in leaves of D. purpurea was 180 and 8 μg gFW 1, respectively. The content of both cardenolides was undetectable after the second subculture of seedling-derived callus. Similar results were obtained with calli induced from leaf discs. In these experiments, the calli cultured under light did not become green [18]. Finally, shoot organogenesis was performed with 10 mg l 1 of benzyladenine (BA) for D. purpurea under a 12-hour daylight cycle, but the development of the leaves was imperfect. Besides, the biosynthesis of digitoxin and digoxin was restarted in redifferentiated shoots and in root-forming calli as previously reported Hirotani and Furuya [14]. These authors concluded that cardenolide contents of the six Digitalis species were variable and strongly related to leaf development [18]. These assumptions were further confirmed years later by the same research group [21]. The second publication was focused on the effect of the light and plant growth regulators on digitoxin production of D. purpurea grown in liquid media [19]. For this, cell lines (undifferentiated green and white lines; green and white shootforming cultures) were used to elucidate the effect of light in cardenolide production. As expected, digitoxin content was considerably higher in shoot-forming cultures (40 μg gDW 1) than in undifferentiated cells (up to 0.21 μg gDW 1). Moreover, light-grown cultures produce more digitoxin than dark grown cultures, but Digitalis cardenolide content was independent of chlorophyll content and RuBisCO activity. Finally, the effect of BA, indoleacetic acid (IAA), NAA, and 2,4-D was studied for green shoot-forming cultures. The digitoxin formation was stimulated by growth regulators at low concentrations (BA and IAA 0.01–1 mg l 1; NAA and 2,4-D 0.1 mg l 1) and suppressed by higher ones (BA and IAA 10 mg l 1; NAA 1 mg l 1; and 2,4-D 1 and 10 mg l 1) in parallel with shoot differentiation [19]. In the third study, Hagimori et al. [20] evaluated the effect of nutrients on digitoxin formation by shoot-forming cultures of D. purpurea grown in MS liquid media. This publication marked the guidelines for establishing the composition of the culture medium in many subsequent studies. Sucrose, glucose, and raffinose yielded higher growth and digitoxin content in comparison with other carbon sources evaluated. Further analysis of the sucrose concentration effect allowed them to determine the optimal concentration for digitoxin production (3%) and cell growth (5%). The nitrogen concentration of the MS medium was too high for the production of cardenolides; two-thirds of the basal concentration was recommended. Besides, a phosphate concentration of 510 mg l 1 is more suitable for digitoxin production and coconut milk for cell growth without affecting secondary metabolite. Thiamine HCl and myo-inositol are necessary for digitoxin production. Finally, the effect of other plant growth substances, i.e., gibberellic acid (GA3), abscisic acid (ABA), and kinetin, was analyzed. Digitoxin formation was slightly improved by concentrations of 0.01–0.1 mg l 1 of GA3 or ABA and repressed by

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higher concentrations. Kinetin effect was not clearly identified. In summary, this work demonstrates that the culture medium composition should be selected for optimal achievement of the goal pursued in the research. The revised medium previously described was further studied in jar fermenter culture [22]. The total cell yield and digitoxin concentration per jar were 38.5 g DW 1 and 1610 μg per jar, respectively. Besides, some recommendations to achieve higher digitoxin content were proposed. A similar medium composition was applied by Matsumoto et al. [24], but to evaluate the production of a group of phenolic glycosides in callus and leaves of D. purpurea. Five and four glycosides were isolated and identified from the leaves and callus, respectively. Both explants were able to synthesize different kinds of secondary metabolites, presumably related to plant defense against fungi and viruses. A few years later, the effect of the natural cytokinin N6-[2-Isopentenyl] adenine (2iP) on shoot differentiation was evaluated [25]. Leaf lamina and petiole were used as initial explant. The composition of the basal culture medium contained LS medium, B5 vitamins, sucrose, myo-inositol, and glycine, but not thiamine HCl as previously recommended by Hagimori et al. [20]. Callus proliferation was enhanced by the combination of 2,4-D and NAA. Moreover, the presence of BA and kinetin induced a synergistic effect on auxins. Bud and shoot differentiation occurred in leaf explants and callus cultures grown with 1.00–5.00 mg
l 1 2iP. Hence, direct shoot organogenesis was also performed in this work. Finally, the regeneration protocol allowed to obtain 7–10 plants per leaf explant [25]. D. purpurea tissue culture was further applied to analyze the activity of enzymes involved in cardenolide biosynthesis [26–28]. In these works D. purpurea progesterone-5β-reductase and 3β-hydroxysteroid-5β-oxidoreductase were studied in photomixotrophic shoot cultures [26, 27]. In addition, progesterone-5β-reductase was purified and characterized for this species [28]. Furthermore, the effects of auxins (IAA, NAA, BA, GA3) and phenobarbital on morphogenesis, ultrastructure, and digitoxin content were determined in callus derived from D. purpurea hypocotyls [29, 30]. The callus growth and vacuole/ cytoplasm ratio were increased with NAA and affected by the stress-induced effect of phenobarbital. This compound also caused several changes in mitochondria and chloroplast structures [30]. On the other hand, digitoxin production was increased with the combination of phenobarbital and IAA, even when biomass was reduced, but not with NAA. Therefore, these authors conclude that biomass production and the digitoxin biosynthesis process are not directly related. However, cardenolide production was positively correlated to mitochondrial activity in cells and morphogenesis of callus tissue [29]. More recently, a plant regeneration procedure in combination with an Agrobacterium-mediated genetic transformation protocol was described by PérezAlonso et al. [31] (Fig. 1). The effects of 2,4-D and BA in callus induction and IAA and BA in plant regeneration were studied. Callus formation (Fig. 1a) was observed in all treatments with 2,4-D, while BA affected callus induction. For plant regeneration, callus was segmented and transferred to a regeneration medium under light conditions. Shoot-producing callus became green after 7–10 days (Fig. 1b), but only some

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Fig. 1 Plant regeneration from leaf segments of in vitro-cultivated Digitalis purpurea plants as proposed by Pérez-Alonso et al. [31]. (a): Callus obtained on MS semisolid medium supplemented with 4.5 μM 2,4-D after 4 weeks. (b): Shoot-producing callus segments became green after 7–10 days in regeneration medium (MS semisolid medium supplemented with 4.4 μM BA and 0.57 μM IAA). (c): Callus segment after 2–3 weeks in regeneration medium. (d): Regenerated plants after 8 weeks. (e): Regenerated plant established in the pot. Bars: (a) 10 mm, (b, c), 4 mm, (d) 50 mm, (e), 1.5 cm

treatments lead to shoot formation. The best growth regulator combination was 4.5 μM 2,4-D for callus induction and 0.57 μM IAA and 4.4 μM BA for plant regeneration (Fig. 1c, d), resulting in 16 shoots per leaf segment [31]. Roots were easily developed in vitro using the same regeneration medium, and rooted shoots were able to be transferred to soil (Fig. 1e). The protocol obtained is applicable to functional studies for a better understanding of cardenolide biosynthetic pathways and the metabolic engineering of cardenolides to develop high-yielding improved genotypes.

4.2

Direct Organogenesis

Direct shoot organogenesis of D. purpurea was less developed in earlier studies compared with other Digitalis species, like D. thapsi [32] and D. minor [33, 34], although cardenolide production was known to be directly related to green-

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differentiated tissue [3] and evidence of shoot differentiation from leaf segments was previously reported [25]. Advantages of avoiding a callus formation phase are that tissue culture time can be shortened and that somaclonal variation associated with callus induction can be diminished [35, 36]. In vitro propagation procedures of D. purpurea plants via direct organogenesis have been published in the last decade, often combined with other strategies to enhance cardenolide production [36–39]. The use of temporary immersion system (TIS) cultures for Digitalis biomass (Fig. 2a, b) and cardenolide production was first reported by Pérez-Alonso et al. [37]. In this work, shoot regeneration from seedlings was obtained on MS medium containing thiamine HCl (1 mg l 1), BA (1 mg l 1), IAA (0.1 mg l 1), myo-inositol (100 mg l 1), sucrose (30 g l 1), and gelrite (3 g l 1). The effect of immersion frequencies on the biomass of D. purpurea was studied using two glass vessels (1 l capacity each), one with shoot cultures and the other with liquid culture medium as a reservoir. Biomass accumulation increased during the 28 days of culture. Best results of fresh weight, dry weight, number of shoots per explant, and lower hyperhydricity were obtained with immersions every 4 h (six times per day). Digitoxin and digoxin contents were determined in all treatments, but lanatoside C was never detected. The immersion frequencies influenced digoxin concentration variation, but not digitoxin content. Finally, glycoside production in TIS is recommended as a suitable alternative for D. purpurea organ culture [37]. In 2013 Patil et al. [38] published a compelling work on in vitro propagation and cardenolide production in shoot cultures under different growth regulator concentrations, elicitation, and precursor feeding treatments. Shoot regeneration was performed from nodes, internodes, and leaf segments cultured in MS medium with various concentrations of BA, kinetin, and thidiazuron (TDZ). Best results of shoot regeneration frequency (85.7%) and the number of shoots per explant (12.7) were obtained for nodes cultured with 7.5 μM BA. The combination of auxins (IAA, NAA, 2,4-D) with BA, kinetin, and TDZ inhibited the shoot regeneration in all the explants. Liquid and solid media were further evaluated for shoot maintenance, and the liquid medium was further recommended (average production of 18 shoots per explant). Rooting of shoots, mean number of roots per shoot, and root length were increased with 15 μM IAA. Then, the acclimatization of the plantlets was achieved with 80% of survival and without morphological abnormalities. These in vitro culture conditions were applied to produce biomass and cardenolides in D. purpurea. Furthermore, the growth of shoots and cardenolide content were studied at different incubation periods, and optimal values were obtained after 28 days of incubation (digitoxin 53.6 μg gDW 1, digoxin 28.8 μg gDW 1). Besides, the effects of growth regulators on both parameters were analyzed. Shoots grown in a medium without growth regulators produced significantly lower amounts of digitoxin (21.6 μg gDW 1) and digoxin (15.9 μg gDW 1), while the incorporation of 5 μM of an auxin (IAA or NAA) in the medium augmented digitoxin and digoxin accumulation between 1.55- and 2-folds [38]. Moreover, elicitation and precursor feeding considerably increased the cardenolide content up to 9.1- and 11.9-fold (discussed in Sect. 6). Another direct regeneration method for in vitro propagation and Agrobacteriummediated transformation of D. purpurea was reported one year later by Li et al. [39].

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In this work, leaf segments were used as initial explants, and MS medium with combinations of growth regulators TDZ and NAA was evaluated, but cardenolide content was not analyzed. Shoot regeneration was achieved in 100% of the explants with 1 mg l 1 TDZ and 0.1 mg l 1 NAA, with more than six shoots per explant. These conditions were applied to transform D. purpurea leaf segments. Kanamycinresistant shoots were easily rooted in one-half MS medium without growth regulators, and plantlets were next transferred to pots. Finally, these authors proposed a simple direct shoot regeneration system to propagate D. purpurea plants in 7–10 weeks [39]. Recently, Pérez-Alonso et al. [36] developed an efficient direct shoot organogenesis method to obtain in vitro plants of D. purpurea from leaf segments (Fig. 2c, d).

Fig. 2 Shoot multiplication by temporary immersion system (a, b) and direct organogenesis from leaf segments cultured on MS semisolid medium containing 0.54 μM NAA and 13.2 μM 6-BAP (c, d). Bars: (a, b), 1 cm, (c, d) 50 mm

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In this work variable combinations of NAA and BA were studied. Furthermore, genetic stability of the regenerated plants and mother plant was analyzed using random amplified polymorphic DNA (RAPD). Besides, leaf powder from regenerated plantlets was analyzed to fulfill the quality specifications of the British Pharmacopoeia, and cardenolide content was also compared. Shoot regeneration was obtained in 98.5% of the explants with an average number of shoots per leaf segments of 18.9 on MS medium containing 0.54 μM NAA and 13.2 μM BA. Genetic stability and cardenolide content of micropropagated D. purpurea plants were similar to the mother plant (digitoxin 202.7 μg gDW 1, digoxin 22.6 μg gDW 1). The protocol described in this work might be combined with other biotechnological approaches to increase cardenolide production in vitro for industrial purposes. Recently, the influence of light quality on plant growth and cardenolide production in D. purpurea cultured in plant factory was determined [40]. In this work different combinations of red and blue light-emitting diodes (LEDs) promoted biomass (8:2 combination) and cardenolide production (2:8 combination). These studies should be further applied to plants cultured in vitro, to optimize the culture conditions for uniform plant growth and cardenolide production.

5

Genetic Stability

Irrespective of whether in vitro culture is applied for germplasm conservation, plant propagation, or secondary metabolite production, micropropagated plants should preserve the identity and genetic integrity of mother plants [41]. However, somaclonal variation might be caused by several factors including the growth regulators, the type and duration of culture, and the genotype and pre-existing variations like chimeric tissues, among others (reviewed by [42]). These variations can be detected by molecular, morphological, physiological, and biochemical methods. Nevertheless, molecular techniques enable detection in earlier stages even when genetic instability is not yet phenotypically expressed [42, 43]. The first report of DNA stability determination in the Digitalis genus was performed in Digitalis obscura L. shoots by RAPD [44]. In this work, 15 wildgrowing plants were collected in 3 different regions of Spain. Cardenolide production in the field (digitoxin 505–1139 μg gDW 1), genotype effects on survival, and shoot proliferation were correlated with the highly polymorphic RAPD patterns obtained. Afterward, long-term shoot cultures in the higher BA concentration showed a fasciated and hyperhydric phenotype. Two years later, stabilized shoot cultures of the elite genotype (T4) without visible abnormalities were transferred to soil. Finally, RAPD markers allowed to confirm the identity of the donor plant and those plants derived from in vitro culture. Besides, similar amounts of cardenolides were produced [44]. Genetic stability was also evaluated by RAPD in shoot tips of D. minor after several subcultures and cryopreservations [45]. The band patterns of in vitro cultured plants were different from the original wild-growing plant. The authors

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Fig. 3 Gel electrophoresis of a RAPD pattern bands obtained with primers 2 and 6 as reported [36]. M: Digitalis purpurea mother plant, 1–12: Randomly selected plants regenerated by direct shoot organogenesis, MW: GeneRuler 100 bp Plus DNA Ladder (Thermo Scientific), W: Contamination control

demonstrated somaclonal variation was generated by long-term culture but not because of cryopreservation. RAPD markers were also successfully applied to establish phylogenetic relationships between six species within the Digitalis genus [46]. Moreover, this PCR-based molecular method allowed to study populations of Digitalis minor L [47]. Besides, genetic polymorphism in two populations of Digitalis grandiflora Mill. was examined by RAPD and inter-simple sequence repeat (ISSR) methods [48]. These and other newer methods are suggested for genetic studies with diverse applications [49]. To our knowledge, Pérez-Alonso et al. [36] developed the only study on the genetic stability of D. purpurea micropropagated plants (Fig. 3). This work and the results previously compiled in this section demonstrated that genetic stability is another factor to consider in the assessment of an efficient propagation protocol in Digitalis spp.

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Biotechnological Approaches for Biomass and Cardenolide Production

Regardless of all these efforts, cardenolide production by Digitalis plants cultivated in vitro has been considerably lower than in field-grown plants [3]. Therefore, in the last decade, the in vitro culture of D. purpurea has been combined with other biotechnological strategies, like temporary immersion systems (TIS), elicitation, and genetic engineering [50].

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Large-scale shoot culture of D. purpurea in TIS is a suitable strategy for biomass and cardenolide production. Pérez-Alonso et al. [37, 50] proposed a reproducible protocol with this aim and recommended the combination of this method with elicitation. To our knowledge, elicitation and precursor feeding are the most effective methods to enhance cardenolide production on micropropagated D. purpurea. A very exhaustive study on this topic was performed by Patil et al. [38]. In this work different concentrations of abiotic (salicylic acid, mannitol, sorbitol, PEG 600, NaCl, KCl) and biotic (chitin powder, yeast extract, the mycelial mass of Aspergillus niger, Helminthosporium sp., Alternaria sp.) elicitors were evaluated. Biomass (expressed as dry weight) was significantly lower in all the treatments with elicitors. Nevertheless, digitoxin and digoxin accumulation was increased with salicylic acid, mannitol, sorbitol, Helminthosporium sp., and Alternaria sp. The concentrations of elicitors that yielded higher cardenolide content are summarized in Table 1. PEG 6000 and NaCl enhanced digoxin content, but not digitoxin. Considerable enhancement of cardenolide content was obtained by using KCl (digitoxin 7.7-fold and digoxin 8.6fold) and Helminthosporium sp. mycelial mass (digitoxin nearly 250 μg gDW 1 and digoxin below 60 μg gDW 1). Another strategy evaluated by Patil et al. [38] was the addition of precursors of cardenolide metabolism to the culture medium. Cholesterol and squalene slightly enhanced digitoxin and digoxin content, but progesterone at concentrations of 200 and 300 mg l 1 induced the highest increase in accumulation of these two cardenolides (9.1- and 11.9-fold) reported so far. Unfortunately, precursor feeding is expensive and not commercially viable for cardenolide production in vitro [3]. Other elicitors were studied by Pérez-Alonso et al. [51]. Biomass and cardenolide content were enhanced by SilioPlant (0.01 g l 1) and ChitoPlant (Table 1). The addition of methyl jasmonate affected all the parameters evaluated. Genetic engineering is another alternative to enhance cardenolide biosynthesis [52]. With this goal, two Agrobacterium-mediated transformation protocols of D. purpurea leaf segments have been developed, one using direct organogenesis [39] and the other by indirect organogenesis regeneration [31]. The protocol proposed by Li et al. [39] yielded 0.62 kanamycin-resistant shoots per initial explant in a short time. However, escapes of the selection process and chimeric plants might be obtained. On the other hand, Pérez-Alonso et al. [31] achieved an efficiency of 6.91 transgenic lines per initial leaf segment selected with geneticin. This protocol is considered as a reliable and fast tool for metabolic engineering in D. purpurea [7, 53]. Sales et al. [52] introduced a gene coding for the catalytic domain of a key enzyme in the mevalonic acid biosynthetic pathway in D. minor via A. tumefaciens genetic transformation. Its expression increased the accumulation of sterols and cardenolides. This research demonstrated that it is possible to modify the secondary metabolism and to enhance the production of cardenolides through genetic engineering. But no cardenolide pathway modification through genetic engineering has been reported until today. The combination of the biotechnological approaches described in this chapter will open the possibility to establish an economically viable process for cardenolide production in vitro.

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Table 1 Effect of biotechnological approaches on biomass and cardenolide accumulation in shoot cultures of Digitalis purpurea L. Strategy Temporary immersion system (TIS) Elicitation

Reference PérezAlonso et al. [37, 50] Patil et al. [38]

Precursor feeding

Elicitation

7

PérezAlonso et al. [51]

Treatment Immersions during 2 min every 4 h Control Salicylic acid 50 μM Salicylic acid 200 μM Mannitol 200 mM Mannitol 300 mM Sorbitol 100 mM Sorbitol 200 mM PEG 6000 5 mM NaCl 100 mM KCl 80 mM KCl 200 mM Progesterone 200 mg l 1 Progesterone 300 mg l 1 Cholesterol 200 mg l 1 Cholesterol 300 mg l 1 Squalene 4 mM Control ChitoPlant 0.1 g l 1 SilioPlant 0.01 g l 1

Biomass DW (g) 5.82  0.46

Digitoxin (μg gDW 1) 28.8  1.94

Digoxin (μg gDW 1) 20.6  1.29

1.69  0.10 0.67  0.11

53.6  3.1 117.6  2.7

18.0  1.0 23.0  0.9

0.49  0.19

28.4  1.2

134.3  1.6

0.49  0.20

98.9  0.9

87.0  1.1

0.42  0.10

140.1  2.2

62.5  1.1

0.71  0.16

157.5  1.2

134.3  1.7

0.37  0.12

186.1  1.7

114.1  1.2

0.64  0.05

38.2  2.4

54.5  1.7

0.10  0.03 0.66  0.10 0.24  0.12 0.38  0.06

22.1  1.2 413.6  2.1 106.3  1.1 492.6  1.8

32.7  1.3 102.2  1.3 156.4  1.5 184.4  1.5

0.31  0.04

216.4  1.5

214.2  1.2

0.30  0.01

136.9  1.6

54.2  1.2

0.30  0.03

112.4  1.3

78.2  1.9

0.30  0.01

81.4  1.3

31.9  1.4

0.4  0.01 0.55  0.01

27.0  0.5 66.0  0.3

3.0  0.01 9.0  0.02

0.48  0.01

187  0.8

10  0.01

Conclusions

In vitro culture of D. purpurea has been widely studied, but it is yet not feasible to develop an economically viable alternative process using biotechnology. Digitoxin and digoxin accumulation in vitro is still lower than that produced by plants grown in

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natural conditions. Therefore, the growing parameters in vitro should be selected to promote biomass formation and a higher yield of cardenolides. For this, plant genotype, shoot differentiation, culture time, medium composition (like macroand micronutrients, growth regulators, vitamins), and light photoperiod and wavelength should be carefully chosen. Then the influence of these conditions on plant growth and genetic stability should be further evaluated in balance with the yield of desired cardenolides for pharmaceutical purposes. Finally, the optimized in vitro culture conditions might be combined with other biotechnological approaches for large-scale uniform production. Elicitation and precursor feeding are the most effective strategies developed until today, but both affect biomass production and imply additional costs. Thus, the enhancement of cardenolide production in vitro by genetic engineering is the most promising alternative to obtain elite D. purpurea lines. The combination of in vitro culture techniques and one or more biotechnological strategies is needed to fulfill the requirements for pharmaceutical industries in modern times.

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Bioreactor-Grown Shoot Cultures for the Secondary Metabolite Production Agata Krol, Adam Kokotkiewicz, Agnieszka Szopa, Halina Maria Ekiert, and Maria Luczkiewicz

Contents 1 2 3 4

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Materials and Methods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Secondary Metabolite Production in Bioreactor-Grown Shoot Cultures . . . . . . . . . . . . . . . . . . Types of Bioreactors Used for In Vitro Shoot Cultures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1 Mechanically Agitated Bioreactors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2 Pneumatically Agitated Bioreactors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.3 Temporary and Continuous Immersion Systems . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.4 Gas Phase Bioreactors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5 Comparative Studies on Bioreactor Performance . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6 Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

189 190 190 194 195 197 212 213 238 240 240

Abstract

In vitro shoot cultures have long been investigated as a potential source of added value chemicals. Similarly to cell cultures, they can be employed for the production of compounds of interest such as drugs, antioxidants, and flavorings. Since in vitro shoots retain tissue differentiation of the parent plant, they are often capable of biosynthesizing secondary metabolites not found in unorganized cell suspensions. However, large-scale cultivation of shoot cultures is challenging and A. Krol · A. Kokotkiewicz · M. Luczkiewicz (*) Department of Pharmacognosy, Faculty of Pharmacy, Medical University of Gdańsk, Gdańsk, Poland e-mail: [email protected]; [email protected]; [email protected]; [email protected] A. Szopa · H. M. Ekiert Department of Pharmaceutical Botany, Faculty of Pharmacy, Jagiellonian University, Medical College, Kraków, Poland e-mail: [email protected]; [email protected]; [email protected] © Springer Nature Switzerland AG 2021 K. G. Ramawat et al. (eds.), Plant Cell and Tissue Differentiation and Secondary Metabolites, Reference Series in Phytochemistry, https://doi.org/10.1007/978-3-030-30185-9_34

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requires specialized bioreactor systems. In the current work, reports concerning secondary metabolite production in bioreactor-grown shoot cultures were reviewed with respect to the examined compounds, types of bioreactors used, and comparative studies involving different fermenter systems. The aim of the chapter is to compile the results of experimental papers, with the emphasis on providing important experimental details and outcomes of the studies, including productivities of bioreactor-grown shoot cultures with respect to secondary metabolites. Keywords

Bioreactors · Shoot cultures · In vitro culture · Secondary metabolites · Largescale plant tissue culture List of Abbreviations

2,4-D 2iP ABA ALB BA BB BCB BFB CIB GA3 GPB HB IAA IBA ITR KIN MC MS NAA PGR RB SAB SH STB T TDZ TIB TS

2,4-Dichlorophenoxyacetic acid 6-(γ,γ-Dimethylallylamino)purine Abscisic acid Airlift Bioreactor 6-Benzylaminopurine Balloon Bioreactor Bubble Column Bioreactor Bubble-Free Bioreactor Continuous Immersion Bioreactor Gibberellic acid Gas Phase Bioreactor Hydraulic Bioreactor Indole-3-acetic acid Indole-3-butyric acid Immersion Time Ratio Kinetin Misting cycle Murashige-Skoog Naphthaleneacetic acid Plant Growth Regulators Roller Bioreactor Simple Aeration Bioreactor Schenk and Hildebrandt Stirred-Tank Bioreactor Temperature Thidiazuron Temporary Immersion Bioreactor Timespan

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Bioreactor-Grown Shoot Cultures for the Secondary Metabolite Production

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Introduction

For several decades, plant in vitro cultures have been investigated as a potential source of secondary metabolites. In particular, they can be employed for the production of valuable plant substances such as drugs, dyes, natural pesticides, and antioxidants. As compared to wild-grown or cultivates plants, in vitro systems offer several advantages which have been extensively discussed in numerous review papers. Plant cells or organs grown in vitro can be used for the production of desired chemicals year-round, regardless of the season, climate zone, weather disturbances, and other environmental factors such as pests and soil quality. Both chemical environment and physical conditions can be strictly controlled, thus enabling to optimize the process for high and stable metabolite yields [1–8]. Plant in vitro cultures are especially useful in the case of high-value products whose natural resources are limited and threatened by overexploitation: the anticancer drug paclitaxel, red dye shikonin, and ginseng biomass are well-known and widely quoted examples of such substances [6, 9–14]. From a technical standpoint, nondifferentiated cell cultures are the most suitable for secondary metabolite production. The major advantage of plant cell suspensions is that they can be adapted for large-scale bioreactor cultivation with relative ease which is crucial for industrial production of a desired chemical. With some modifications, conventional stirred-tank bioreactor (STB) systems can be adjusted for plant cells and subsequently employed for high-volume batch processes [6, 7, 15, 16]. For example, the previously mentioned paclitaxel is produced on industrial scale using cell suspensions of Taxus sp. [10, 13, 17, 18]. Despite obvious advantages, the aforementioned type of culture also has its limitations. The most important ones are genetic instability and the inability to biosynthesize metabolites whose accumulation requires cytodifferentiation. The examples of such products include artemisinin [19], hypericins [20], and volatile constituents [21], stored in specialized secretory structures not present in nondifferentiated cells. In order to overcome these problems, attempts were made to establish large-scale in vitro systems based on plant organs which could be employed for the production wider range of secondary metabolites. These include adventitious and hairy roots, as well as shoots, microshoots, and plantlet cultures. The secondary metabolism of bioreactor-grown adventitious [22–24] and hairy roots [25–27] have been reviewed in several articles, however, analogous papers concerning in vitro shoots are scarce [28, 29]. The aim of the chapter is to compile the literature data concerning low molecular mass secondary metabolites production in bioreactor cultures of shoots, microshoots, and plantlets. For clarity and conciseness, the collected data are presented in tables which included information of the examined plant species, biomass type, bioreactor construction, experimental details, as well as concentration and productivities of the investigated compounds.

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Materials and Methods

The experimental papers were collected using Scopus database, with “bioreactor”/ “fermenter” + “shoots”/“microshoots”/“plantlets” used as search terms in “Article Title, Abstract, Keywords.” Of the articles published in the years 1984–2019, only the papers concerning secondary metabolite production were included in the review whereas the ones describing solely micropropagation procedures were omitted. The cultures of somatic embryos were not included unless they were used to establish plantlet cultures further investigated for the accumulation of secondary metabolites. Articles dealing with recombinant protein production using in vitro shoots and plantlet cultures are also not in the scope of the current work. Besides experimental papers, the so far published review papers concerning bioreactor systems, in vitro and in vivo production of specific groups of secondary metabolites and plant in vitro cultures in general, were included. The information concerning concentrations and productivities of secondary metabolites was extracted directly from text, tables, and graphs or calculated using the available experimental data, such as growth medium volume, fresh or dry weight of biomass, and duration of experiment. Xyscan software (version 4.3.0, Thomas S. Ullrich) was used to extract the data from graphs.

3

Secondary Metabolite Production in Bioreactor-Grown Shoot Cultures

The interest in the production of specific metabolites using in vitro culture methods stems from several factors. Most importantly, the compound of interest needs to be of high value and/or its natural resources limited, so that the in vitro-grown biomass can serve as commercially viable source [1, 2, 5, 30, 31]. As mentioned earlier, cell suspensions are preferred in this regard because of easier scale-up [6, 7, 15, 16, 31], but are often genetically unstable and/or fail to accumulate substantial amounts of desired substance [4, 7, 31, 32]. On the other hand, in vitro cultures of microshoots, shoots, and plantlets (further collectively referred to as “shoot cultures”) retain (at least some of them) tissue differentiation of their parent plants and thus are usually capable to accumulate a variety of secondary metabolites [1, 31]. Consequently, attempts were made to employ the aforementioned cultures for bioreactor-scale production of low molecular weight constituents. Figure 1 depicts the number of papers, dealing with bioreactor-grown shoot cultures evaluated for the presence of compounds representing different chemical classes, published between 1984 and 2019. Clearly, plant phenolics was one of the most extensively investigated groups of secondary metabolites which is likely because of their abundance in plants, as well as the fact that several compounds of this group exhibit antioxidative, therapeutic, and health-promoting properties [33]. In several cases, general spectrophotometric assays were used to determine total phenolic or total flavonoid content of bioreactorgrown shoot cultures (9 and 9 reports, respectively, Fig. 1). These parameters do not allow to calculate the productivity of in vitro systems with respect to specific constituents but are sometimes used to assess the quality of the plant material

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12

Number of papers

10 8 6 4 2

Alkaloids Benzophenones Diterpenes Triterpenes Flavonoids Phenolic glycosides Phenolic acids Lignans Xanthones Coumarines Phenyloetanoids Sesquiterpene lactones Cardiac glycosides Naphthodianthrone Phloroglucinol derivative Essential oil Secoirydoids Iridoids Total flavonoids Total phenolics

0

Fig. 1 The number of papers concerning the accumulation of different classes of secondary metabolites in bioreactor-grown shoot cultures, published between 1984 and 2019. The data was collected using Scopus database

obtained by micropropagation [34–37]. Other studies focused on the accumulation of selected phenolic metabolites like isoflavones [38, 39], flavones [40], xanthones [41], phenolic acids [42–45], lignans [46], and hypericins [20] (15 reports in total). The production of isoflavones, investigated chiefly because of their phytoestrogenic and potential anticancer properties, was studied in shoot cultures of legume plants such as Genista tinctoria [38], Pueraria tuberosa [39], and Cyclopia genistoides [41]. Bioactive flavones, responsible for therapeutic properties of skullcap (Scutellaria sp.), constitute another group of phenolics examined in bioreactor-grown biomasses [40]. There are also several studies concerning the accumulation of lignan constituents in shoot cultures. Within this group, anticancer aryltetralin lignans like podophyllotoxin are of particular interest. However, bioreactor-grown shoots have so far not been evaluated for their production, even despite the fact that experiments on suspension cultures of Podophyllum hexandrum and Linum flavum proved that loss of cell differentiation negatively affects lignan biosynthesis [47]. On the other hand, there are numerous reports dealing with accumulation of dibenzocyclooctadiene lignans in bioreactor cultures of shoots. This topic was extensively studied by Szopa and coworkers who evaluated in vitro shoot cultures of Schisandra chinensis and its cultivar “Sadova No. 1,” grown in different types of bioreactors, for the production of said constituents, along with flavonoids and phenolic acids [46, 48].

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As mentioned before, the accumulation of several bioactive metabolites in plants is directly related to the presence of specialized structures like glands and secretory canals. The example of such constituents is hypericins, stored in secretory glands present in leaves, stems, and petals of Hypericum perforatum and related species [20]. Since the nonorganized cultures of the above plants yielded only low amounts of naphthodianthrones, bioreactor-grown Hypericum shoots were evaluated as their alternative source [20, 49–51]. Another example is artemisinine, a sesquiterpene lactone being the essential drug in malaria treatment. Similarly to hypericins, the accumulation of artemisinin was shown to be cytodifferentiation-dependent: the compound is localized in glandular trichomes on the leaf surface [19]. Consequently, research on organ cultures of Artemisia annua has been undertaken. Studies in this field were conducted by Liu and co-workers who examined artemisinin production in A. annua shoots grown in different bioreactor systems [52–56]. Besides hypericins and artemisinin, secretory structures are sink organs for a variety of volatile constituents [57]. Due to lack of tissue differentiation, accumulation of essential oils in callus and suspension cultures is usually limited (or completely inhibited) and their composition significantly altered as compared to differentiated cultures [21, 58]. So far, there are relatively few reports dealing with essential oil accumulation in bioreactor-grown shoots. The volatiles were produced by the biomass; however, qualitative differences in essential oil composition (as compared to parent plant) were still observed in several cases. For instance, instead of menthol, the fermenter-grown transformed shoot cultures of Mentha piperita accumulated pulegone (a precursor in menthol biosynthesis) as a major volatile compound [59]. Another example is bioreactor shoot cultures of Rhododendron tomentosum which failed to produce ledol and palustrol (i.e., major volatiles of the parent plant) and accumulated essential oil with ledene oxide as a dominant constituent [60, 61]. These data clearly indicate that even in differentiated shoot cultures, terpene and sesquiterpene pathways are not fully developed. As presented in Fig. 1, several papers were published concerning the accumulation of alkaloids in bioreactor-grown shoots. However, studies on the production of anticancer compounds of this group are scarce. As mentioned earlier, diterpene alkaloids are industrially produced using cell suspensions of Taxus sp. [10, 13, 17, 18], and thus there was no strong rationale to develop plant organ cultures for paclitaxel manufacture. On the other hand, attempts to establish plant cell-based in vitro systems for the production of dimeric indole alkaloids were unsuccessful. Cell cultures of Catharanthus roseus, grown in bioreactors up to 5000 l volume, were shown to accumulate monoindole alkaloids including ajmalicine, serpentine, and catharanthine, but failed to biosynthesize bisindole derivatives vinblastine and vincristine [1, 11, 62, 63]. In vitro shoot cultures of C. roseus were demonstrated to produce dimeric indole alkaloids [63, 64 ] but low yields hampered the scale-up studies. So far, there is only one report dealing with alkaloid production in bioreactor-grown shoots of C. roseus. The culture, maintained in a mist-type bioreactor, accumulated monoindole alkaloids vindoline and catharanthine. However, the production of bisindole alkaloids was not examined in the study [65]. As far as other anticancer agents are concerned, there is a single report on the accumulation of camptothecin in bioreactor cultures of Camptotheca acuminata shoots [66].

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Among other alkaloids, galanthamine gained much interest because of its abilityacetylcholinesterase (AChE)-inhibiting properties, responsible for therapeutic effects in Alzheimer’s disease [67]. Since cell differentiation favored accumulation of galanthamine, studies focused on evaluating shoot cultures of Leucojum aestivum for the production of said metabolite. Different bioreactor systems were tested [68–71] and the results of these experiments were summarized in the review paper Berkov and co-workers [67]. Another alkaloid which could be potentially used in the treatment of neurological diseases is securinine, the indolizidine-class compound present in East-Asian plant Securinega suffruticosa. In vitro shoot cultures of this species were established and subsequently adapted for bioreactor cultivation [72]. Studies on tropane alkaloids production using in vitro systems are also worth noting. Monocultures of shoots and shooty teratomas of solanaceous species proved to be a poor source of tropane alkaloids [73] which is due to the fact that hyoscyamine biosynthesis is localized in roots, while shoots serve as its pool organ and place where further steps of tropane pathway take place [74]. Nevertheless, bioreactorgrown co-cultures of hairy roots and shooty teratomas of plants from nightshade family were capable of producing alkaloids and proved to be useful for biotransformation studies [74, 75]. Cardiac glycosides comprise another group of bioactive compounds whose production in in vitro systems has been extensively studied. The major observation was that loss of tissue differentiation is followed with substantial loss of biosynthetic capability [76, 77]. Cell suspension cultures of Digitalis lanata (major commercial source of cardiac glycosides) were successfully scaled-up using pilot size fermenters. However, these cultures failed to produce therapeutically relevant compounds and could only be used for biotransformation reactions [77]. The decline in the use of digitalis glycosides in congestive heart failure [78] was reflected in diminishing number of papers dealing with large scale cell cultures of Digitalis. Nevertheless, cardiac glycosides gained interest as potential anticancer agents and studies on their production using in vitro systems, including shoot cultures, have been continued. This topic was comprehensively reviewed by Verma and co-workers [76]. In the studies concerning in vitro shoot cultures of foxglove, attempts were made to employ temporary immersion systems for the production of digitoxin and digoxin [79, 80]. From a practical standpoint, the productivity with respect to a desired metabolite is a key factor determining the usefulness of an in vitro system. Given the high cost of cell culture maintenance, substantial increase in secondary metabolite concentration is necessary to keep the production commercially viable. As far as bioreactor cultures of shoots are concerned, the experiments yielded rather unsatisfactory results. In Table 1, raw materials of selected plants were compared with bioreactor-grown shoot cultures of the respective species in terms of secondary metabolite content. In most cases, the achieved concentrations of compounds of interest were lower or comparable to wild grown or cultivated plants. The notable exception to this rule is in vitro shoot cultures of Genista tinctoria which accumulated over 4-times more genistin in comparison with intact plant material [38]. However, the reported concentrations were still lower than in cell suspensions of the above plant where genistin amounts of up to 94 mg g 1 were reported [81]. In some cases, as, for

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Table 1 Comparison of secondary metabolite levels in bioreactor-grown shoots and intact plant materials of selected plant species Plant name Artemisia annua Camptotheca acuminata Centaurium erythraea Cyclopia genistoides Digitalis lanata Digitalis purpurea Echinacea purpurea Genista tinctoria Hypericum perforatum Leucojum aestivum Pueraria tuberosa Schisandra chinensis Scutellaria baicalensis Securinega suffruticosa Stevia rebaudiana a

Concentration (mg g In vitro shootsa 2.8–3.6 (GPB) [52, 54, 82] 0.1–2.6 (TIB) [66]

Compound Artemisinin (sesquiterpene lactone) Camptothecin (quinolizidine alkaloid) Gentiopicroside 8.0–22.0 (GPB) [84] (secoiridoidglucoside) Mangiferin (xanthone 10.3–26.2 (TIB) [41] C-glucoside) Digoxin (cardiac glycoside) 0.0–3.1  10 2 (TIB) [80] Digitoxin (cardiac 2.5  10 2–3.3  10 (TIB) [79] glycoside) Cichoric acid (phenolic 15.3–21.0 (LR) [42] acid) Genistin (isoflavone 69.4 (SAB) [38] glucoside) Hypericin 1.1  10 2–3.5  10 (BB) [49, 50] (naphthodianthrone) Galanthamine 0.6  10 1–5.2  10 (Amaryllidaceae alkaloid) (TIB) [68, 69] Puerarin (isoflavone 0.3–1.5 (SAB) [39] C-glucoside) Schisandrin 6.1  10 1–1.2 (dibenzocyclooctadiene (TIB) [46] lignan) Baicalin (flavone glycoside) 2.7 (TIB) [40] Securinine (indolizidine alkaloid) Stevioside (diterpene glycoside)

5.0–5.3 (SAB) [72] 6.7  10 1–6.5 (RB) [98, 99]

1

DW) Intact plant material 0.1–15.0 (herb) [19, 54, 56] 0.2–5.0 (leaves) [31, 83] 1.0–6.2 (herb) [85] 36.0–59.4 (leaves) [86]

2

2

1

8.6  10 2–2.0 (leaves) [87, 88] 2.1  10 1–8.1  10 1 (leaves) [89, 90] 4.9–21.4 (shoots) [42] 1.4–20.5 (roots) [42] 14.5 (herb) [91] 1.1  10 1–1.1 (herb) [20] 1.0–5.0 (leaves) [67] 0.0–100.8 (tubers) [92] 0.3 (leaves) [46] 1.3–34.0 (fruits) [46, 93, 94] 70.1–86.7 (root) [95] 1.2 (herb) [96] 2.0–9.0 (herb) [97] 91.0 (leaves) [100]

Bioreactor nomenclature according to the list of abbreviations

example S. baicalensis and S. chinensis, the contents of secondary metabolites in shoot cultures exceed those recorded in aerial parts of the parent plants, but are nevertheless noticeably lower as compared to raw materials like roots or fruits, serving as a major source of the desired compounds (Table 1).

4

Types of Bioreactors Used for In Vitro Shoot Cultures

The studies conducted during the few last decades demonstrated that in vitro shoot cultures can be successfully grown in a variety of bioreactor types. The stimulus for developing systems dedicated for shoot cultures was mainly from the

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micropropagation industry which was interested in large-scale production of genetically uniform, high quality plantlets [3, 101, 102]. Given the above, the majority of papers in this field concerned in vitro propagation of plants. However, bioreactorgrown shoots have also been investigated as a potential source of secondary metabolites, and this aspect of their use was reviewed in the current work. Since the construction details of bioreactors dedicated for differentiated plant cultures have been reviewed in several papers [3, 6, 28, 101–105], only general characteristics of the respective systems were provided, followed by compilation of data concerning secondary metabolite production. In Fig. 2, the data concerning the number of reports, dealing with secondary metabolite production in shoot cultures grown in various types of bioreactors, was presented, in order to illustrate the popularity of the respective systems.

4.1

Mechanically Agitated Bioreactors

Mechanically agitated bioreactors are widely used in industrial biotechnology [28, 102]. Within this group, the stirred-tank bioreactor (STB) is the most commonly used. Bioreactors of this type have been employed for years in industrial 30

Number of papers

25

20

15

10

5

0 STB

AL + SAB + BB

TIB + CIB

GPB

Fig. 2 The number of papers concerning the accumulation of secondary metabolites in shoot cultures grown in different types of bioreactors, published between 1984 and 2019. The data was collected using Scopus database. Bioreactor nomenclature according to list of abbreviations

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microbiology and are hence popular and reliable systems [3, 7]. After necessary modifications, involving the selection of impeller suitable for shear-sensitive plant cells, these bioreactors can be used for large-scale (up to 75,000 l), high-density (>15 g dry weight per liter) suspension cultures [7, 15, 101, 102, 106]. However, as far as shoot cultures are concerned, these advantages are largely irrelevant because of scale limitations and inherent properties of differentiated biomass. While STBs can be equipped with suitable inoculation system [107] and mesh barrier preventing the shoots from being damaged by the impeller [28], the reactor size cannot be substantially increased due to light requirements of the culture which are difficult to meet in the case of densely packed biomass. As a result, simpler and cheaper bioreactor designs, such as pneumatically agitated vessels and temporary immersion systems, are preferred for the cultivation of in vitro shoots [3, 101–103]. As presented in Table 2, STBs were rarely employed in studies concerning secondary metabolite production in shoot cultures. Besides conventional STBs, other types of mechanically agitated bioreactors were used for shoot cultivation. One of them is the Roller Bioreactor (RB), examined for steviosides production in Stevia rebaudiana plantlets [98, 99]. The apparatus, consisting of multiple low-volume (0.32 l) vessels placed on a rotating shaft, is similar to “Auxophyton,” previously devised by Steward and co-workers [103, 108]. Another example is the “Lab Rocker” (LR) system (Caisson Laboratories, Smithfield, USA), consisting of polycarbonate growth vessels ( 3) [101]. In order to deal with the excessive foaming, the cone-shaped balloon bioreactors (BB, alternatively termed as “baloon type bubble bioreactors” or BTBB) were devised [3, 102]. The airlift (ALB)systems, on the other hand, are equipped with a draft tube designed to reduce shear stress exerted on cells [28, 101]. Over the last two decades, air-driven bioreactors of SAB, BCB, and BB types were often employed in secondary metabolite studies, conducted on shoots, plantlets, and somatic embryos. Growth containers of up to 5 l volume were the most commonly used. Depending on the application, the bioreactors can be equipped with baskets [38, 46, 72], rafts [39] or meshes [37, 54] for biomass immobilization. In the case of G. tinctoria shoots-hairy roots co-culture, two concentric baskets were used to separate the two biomasses [32, 38]. In other instances, the installed supports were meant to reduce mechanical stress and hyperhydricity, a common issue with fully submerged shoot cultures [3, 102, 103, 114]. The low-cost and disposable versions of pneumatically agitated systems were also developed: the examples include the aerated Growtek bioreactor (Tarsons, India) [39] and LifeReactor (Osmotek, Israel) [114], respectively. In Table 3, the results of experiments on secondary metabolism of shoot cultures maintained in air-driven bioreactors were summarized.

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Table 2 Secondary metabolite production in shoot cultures grown in mechanically agitated bioreactors Type of stirrer and constructions details

Medium

Experiment parameters Photo- Light period intensitya Tempera- Timespan ture (TS) Air flow rate Volume Sucrose PGR [h light/ [μmol [vvm] [ml] Salts [gL 1] [mg L 1] h dark] m 2 s 1]a (T) [ C] [days]

Magnetic stirrer

1000

MS

30

–

24/0

7.8

25

28

0.11

Magnetic stirrer; the dual reactor system with mesh installed in the center of the vessel to support the root culture Magnetic stirrer and turbine impeller

3000+ 3000

MS

30

–

24/0 for 7.8 shoots

25

28

0.037–0.067

2000 + 2000

MS

–

–

24/0

21–28

25

29

0.2

Magnetic stirrer

1000

MS

–

–

24/0

21

25

28

0.11

Magnetic stirrer

3500

MS

–

–

24/0

21

25

28

0.029–0.057

Two flat-blade 2500 impellers having 4 blades with rotational speed 100 rpm

MS

–

BA 1.0; IAA 1.0

24/0

78.3

28

15

1.4

“Southern Sun Liquid Lab Rocker” growth system with rocking motion of the system [30 s ON/ 132 s OFF] Impeller with rotational speed 10 rpm, equipped with

100

MS

30

–

16/8

40–85

NS

12

NS

4800

B5

30

2,4-D 2.0

NS

NS

NS

23

NS

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Plant culture

Type

Species

Co-culture of A. belladonna hairy roots and shooty teratomas

Atropa belladonnaand a Duboisia leichhardtii X D. myoporoides hybrid Co-culture of Atropa belladonna A. belladonna and a Duboisialeichhairy roots and hardtii X shooty D. myoporoides teratomas hybrid

Metabolite Hyoscyamine Scopolamine

Hyoscyamine Scopolamine

Productivity Max. content [mg L 1 [mgg 1 DW] d 1] Parameters studied 8.00  10 in shoots 2.20  10 in shoots

2

2.20  10 in shoots 1.50  10 in shoots

1

1

1

3.14  10 in shoots 8.64  10 in shoots

2

8.51  10 in shoots 5.80  10 in shoots

2

Co-culture of Atropa belladonna transformed roots and Duboisia hybrid shooty teratomas Atropa belladonna shooty teratomas

Atropa belladonna Hyoscyamine and Duboisia hybrid Scopolamine

ND 4.80 in shooty teratomas

– 6.62  10 in shooty teratomas

Atropa belladonna

Scopolamine

ND

–

Atropa belladonna shooty teratomas

Atropa belladonna

Scopolamine

ND

–

Shoot

Digitalis purpurea

Digitoxin

4.18  10

Plantlets

Echinacea purpurea Cichoric acid

2.19  101 in 9th day

NA

Shoot

Rutagraveolens

NA

2.00

Furanocoumarins

2

4.29  10

2

2

4

2

Ref

Effects of different [75] bioreactor systems on in vitro biomass production and accumulation of scopolamine Effects of different [75] bioreactor systems on in vitro biomass production and accumulation of scopolamine Effects of interspecies and intergenus organ co-culture on the production of plant secondary metabolites Effects of different bioreactor systems on morphology of teratomas and scopolamine content Effects of different bioreactor systems on morphology of teratomas and scopolamine content Effect of mechanical agitation on the growth of cells and digitoxin content Effects of GA3, paclobutrazol, and uniconazole on secondary metabolite production

[74]

[73]

[73]

[110]

[42]

Effects of different [111] bioreactor systems on furanocoumarin production

(continued)

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Table 2 (continued) Medium Type of stirrer and constructions details

Experiment parameters Photo- Light period intensitya Tempera- Timespan ture [h light/ [μmol (TS) Air flow rate Volume Sucrose PGR [vvm] [ml] Salts [gL 1] [mg L 1] h dark] m 2 s 1]a (T) [ C] [days]

temperature, oxygen and pH probes The roller 4 bioreactor – the flow of nutrient medium inside the vessel with 4 rpm rotation velocity of disks The roller 4 bioreactor – the flow of nutrient medium inside the vessel with rotation velocity of disks 4 rev/min.

MS

30

–

NS

28

25  1

35

NS

MS

30

–

NS

28

25  1

35

NS

NA data not available, NS data not specified, ND not detected a Unit conversion in accordance with Sager and McFarlane [112]

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Plant culture

Type

Species

Metabolite

Shoot

Stevia rebaudiana

Stevioside Rebaudioside A Rebaudioside C

Shoot

Stevia rebaudiana

Stevioside Rebaudioside A Rebaudioside C

Productivity Max. content [mg L 1 1 [mgg DW] d 1] Parameters studied

4.27  10

1

1.66  10

1

7.98  10

2

1

5.74  10

1

1

1.08  10

1

1

1.08  10

1

6.42 in leaves 2.50 in leaves 1.20 in leaves 6.37  10 in leaves 2.53  10 in leaves 1.20  10 in leaves

Ref

Effects of medium [98] composition on growth and accumulation of steviol glycosides in the roller bioreactor Effects of medium [98, 99] composition on growth and accumulation of steviol glycosides in the roller bioreactor

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Table 3 Secondary metabolite production in shoot cultures grown in pneumatically agitated bioreactors Medium

Experiment parameters

Type of Volume system [ml] Salts

Photoperiod [h light/ Sucrose PGR 1 1 [g L ] [mg L ] h dark]

Light intensitya Temper- Timespan Air flow (TS) rate Air sparger [μmol ature [vvm] constructions m 2 s 1]a (T) [ C] [days]

AL

2000

MS

30

NAA 0.05; BA 0.5

16/8

42

25  1

25

1

Vertical immersed tube with holes, fixed in the center of the bioreactor

AL

5000

MS

30

–

16/8

40

22

56

0.1

NS

SAB

NS

MS

30

–

16/8

75

23

28

NS

Collapsible vessel (“LifeReactor”) attached to an air pump via a filter disc

SAB

400

MS

–

IBA 0.05; TDZ 0.25

16/8

50

25 0.2

28

0,1–0,3b “Growtek” bioreactor with aeration provided by a needle inserted through side tube of bioreactor

SAB

150

MS

30

BA 0.25

16/8

100

22  1

28

NS

“Growtek” bioreactor

SAB

500

MS

30

BA 2.0; NAA 0.15

16/8

30

24  2

56

0.1

NS

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Plant culture

Type

Species

Metabolite

Max. content [mgg 1 DW]

Productivity [mg L 1 d 1]

Shoot

Artemisia annua

Artemisinin

1.80  10

5.81  10

Eleutheroside B

5.6  10 2 for plantlets

NA

Eleutheroside E

3.83  10 1 for plantlets

NA

Eleutheroside E1 3.80  10 1 for plantlets

NA

Embryo Eleutherococcus genic chiisanensis cells

Plantlets Scutellaria baicalensis

Shoot

Pueraria tuberosa

1

Baicalin

1.58  10

3

NA

Baicalein

4.40  10

4

NA

Wogonin

3.20  10

5

NA

Total isoflavonoid 1.89 for 20 v/v aeration

NA

Puerarin

1.48 for 20 v/v aeration

NA

Daidzein

1.26  10 for 20 v/v aeration

NA

Genistein

NA 3.26  10 2 with no aeration

Genistin

2.51  10 for 20 v/v aeration

1

1

Ocimum basilicum Rosmarinic acid

5.12  1010

1.10

Shoot

Leucojum aestivum

5.20  10

4.08  10

1

Parameters studied

Ref

Effect of different bioreactor systems on biomass production and accumulation of artemisinin

[54]

Plantlets production [115]

Effect of different bioreactor systems on biomass production and accumulation of bioactive metabolites

[40]

Effect of aeration [39] volume on biomass growth and isoflavonoid accumulation

NA

Shoot

Galanthamine

1

Effects of different [43] types of bioreactors on biomass production and rosmarinic acid content 2

Effect of the type and size of bioreactors on growth and galanthamine content

[69]

(continued)

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Table 3 (continued) Medium

Experiment parameters

Type of Volume system [ml] Salts

Photoperiod [h light/ Sucrose PGR [g L 1] [mg L 1] h dark]

Light intensitya Temper- Timespan Air flow (TS) rate Air sparger [μmol ature [vvm] constructions m 2 s 1]a (T) [ C] [days]

SAB

450

MS

–

NAA 1.0; KIN 0.1

16/8

NS

25  2

15; 18; 30

0.08

Submerged horizontal sparger at the bottom of the bioreactor

SAB

NS

MS

–

BA 2.5; IAA 0.01

16/8

36

26 0.5

40

0.1

“Growtek” bioreactor with aeration provided by a needle inserted through side tube of bioreactor

SAB

500

MS

–

BA 3.0; NAA 1.0

16/8

60

24  1

30; 60

1.0

Aeration provided by a sparger placed at the bottom of the bioreactor vessel

SAB

500

MS

–

BA 3.0; NAA 1.0

16/8

90  2

24  2

30; 60

1.0

Aeration provided by a sparger placed at the bottom of the bioreactor vessel and internal rack for biomass immobilization

SAB

600

MS

–

BA 0.1; 2iP 3.0; NAA 0.3

16/8

88  8

24  1

70

1.33

NA; bioreactor with the immobilization basket placed above the bottom of the growth vessel

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Plant culture

Max. content [mgg 1 DW]

Productivity [mg L 1 d 1]

Camphor

3.48c for ethephon elicitation

3.37  10 1 for ethephon elicitation

1,8-cineol

7.50  10–1c for gibberellic acid elicitation

1.26  10 1 for gibberellic acid elicitation

β-caryophyllene

6.50  10–1c for ethephon elicitation

6.42  10 2 for ethephon elicitation

Type

Species

Metabolite

Shoot

Artemisia annua

Parameters studied

Ref

Effect of gibberellic [116] acid and ethephon on terpenoids accumulation

Shoot

Bacopa monnieri

Total phenolic content

3.4  101d

NA

Effects of different bioreactor systems on biomass production and accumulation of phenolic compounds

[34]

Shoot

Schisandra chinensis

Schisandrin

5.51  10 1 for TS 30 d

1.47  10 1 for TS 30 d

[46]

Deoxyschisandrin 4.32  10 1 for TS 30 d

1.15  10 1 for TS 30 d

Gomisin A

3.86  10 1 for TS 30 d

1.03  10 1 for TS 30 d

Effects of different bioreactor systems on biomass production and accumulation of schisandra lignans

Total phenolics acid

3.16  10 1 for TS 30 d

8.35  10 2 for TS 30 d

[44]

Chlorogenic acid 2.46  10 2 for TS 30 d

6.63  10 3 for TS 30 d

Gallic acid

8.50  10 2 for TS 30 d

2.29  10 2 for TS 30 d

Protocatechuic acid

8.38  10 2 for TS 30 d

2.26  10 2 for TS 30 d

Effects of different bioreactor systems and cultivation mode on biomass production and accumulation of phenolic acids and flavonoids

Salicylic acid

5.09  10 2 for TS 30 d

1.37  10 2 for TS 30 d

Total flavonoids

2.20  10 1 for TS 30 d

5.93  10 2 for TS 30 d

Quercitrin

1.83  10 1 for TS 30 d

4.94  10 2 for TS 30 d

Securinine

3.29 for supplementation with lysine hydrochloride on 20th day

1.39 for medium supplementation with lysine hydrochloride on 20th day

Shoot

Shoot

Schisandra chinensis

Securinega suffruticosa

Allosecurinine

Effect of medium supplementation with biosynthesis precursors (lysine hydrochloride) on 3.44 for 1.33 for medium biomass yield and supplementation supplementation indolizidine alkaloids content with lysine with lysine hydrochloride hydrochloride on on 20th day 20th day

[72]

(continued)

206

A. Krol et al.

Table 3 (continued) Medium

Experiment parameters

Type of Volume system [ml] Salts

Photoperiod [h light/ Sucrose PGR [g L 1] [mg L 1] h dark]

Light intensitya Temper- Timespan Air flow (TS) rate Air sparger [μmol ature [vvm] constructions m 2 s 1]a (T) [ C] [days]

BCB

1500

MS

–

BA 0.1

16/8

8.3

23

19

1.3

Aeration from the bottom of the bioreactor tank

BCB

1500

MS

–

BA 0.1

16/8

17

23  1

NS

1.3

NS

BB

1000

MS

–

–

16/8

60

24  1

56

0.1

NS; bioreactor with net

BB

1000

MS

–

–

16/8

60

24  1

56

0.1

NS

BB

750

Hypo- 30 nex I

–

16/8

45–50

25  2

84

0.03; 0.06; 0.09; 0.12

NS

BB

NS

MS

–

16/8

75

23

28

NS

Sparger positioned at the bottom of the container and connected to an air pump

30

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207

Plant culture

Type

Species

Metabolite

Max. content [mgg 1 DW]

Productivity [mg L 1 d 1]

Shoot

Hypericum perforatum

Hypericin

7.74  10

1.87  10

Shoot

Shoot

Shoot

Hypericum perforatum

Rosa rugosa

Rosa rugosa

Plantlets Anoectochilus formasanus

Plantlets Scutellaria baicalensis

2

Hyperforin

3.99

Pseudohypericin

2.23  10

1

Hypericin

7.71  10

2

Hyperforin

3.67

Pseudohypericin

3.03  10

Total phenolics

3.90  101

9.93

Total flavonoids

1.31  101

3.34

Total phenolics

3.92  101d

17.0d

Total flavonoids

1.33  10

5.75e

1

3.36 6.52  10

NA NA

1

1e

NA

Total phenolics

1.03  101d

NA

Total flavonoids

2.00  10–1e

NA

Baicalin

1.41  10

3

NA

Baicalein

3.90  10

4

NA

Wogonin

4.72  10

5

NA

1

Parameters studied

Ref

Effects of different culture systems on accumulation of biomass and concentrations of secondary metabolites

[117]

Effects of different culture systems on accumulation of biomass and concentrations of secondary metabolites

[117]

Effects of different bioreactor systems on biomass production and accumulation of total phenolics and flavonoids

[37]

Effects of different bioreactor systems on biomass production and accumulation of total phenolics and flavonoids

[37]

Effect of aeration [118] volume and inoculum density on biomass growth Effect of different [40] bioreactor system on in vitro biomass production and accumulation of bioactive metabolites

(continued)

208

A. Krol et al.

Table 3 (continued) Medium

Experiment parameters

Type of Volume system [ml] Salts

Photoperiod [h light/ Sucrose PGR [g L 1] [mg L 1] h dark]

Light intensitya Temper- Timespan Air flow (TS) rate Air sparger [μmol ature [vvm] constructions m 2 s 1]a (T) [ C] [days]

BB

500

MS

–

BA 3.0; NAA 1.0

16/8

90  2

24  2

30; 60

1.0

Aeration provided by a sparger placed at the bottom of the bioreactor vessel

BB

500

MS

–

BA 3.0; NAA 1.0

16/8

60

24  1

30; 60

1.0

Aeration provided by Sparger placed at the bottom of the bioreactor vessel

BB

1000

MS

60

–

16/8

110

18; 22; 26

35

0.15; 0.30; 0.45

Aeration provided by ceramic Sparger placed at the bottom of the bioreactor vessel; bioreactor with internal sections

BB

NS

MS

30

BA 2.0; NAA 0.15

16/8

30

24  2

56

0.3

NS

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Bioreactor-Grown Shoot Cultures for the Secondary Metabolite Production

209

Plant culture

Type

Species

Metabolite

Max. content [mgg 1 DW]

Shoot

Schisandra chinensis

Total phenolics acid

4.67  10 TS 30 d

1

for 1.07  10 TS 30 d

1

Salicylic acid

5.91  10 TS 60 d

2

for 1.52  10 TS 60 d

2

for

Total flavonoids

2.08  10 TS 30 d

1

for 4.23  10 TS 30 d

2

for

Quercitrin

1.84  10 TS 30 d

1

for 4.22  10 TS 30 d

2

for

1

for 1.53  10 TS 30 d

1

Productivity [mg L 1 d 1]

Parameters studied

Ref

for Effects of different bioreactor systems Chlorogenic acid 4.65  10 2 for 1.07  10 2 for and cultivation mode on biomass TS 30 d TS 30 d production and the 1 2 Gallic acid 1.00  10 for 2.30  10 for accumulation of TS 30 d TS 30 d phenolic acids and 1 2 Protocatechuic 1.63  10 for 3.74  10 for flavonoids TS 30 d TS 30 d acid

[44]

Shoot

Schisandra chinensis

Schisandrin

6.73  10 TS 30 d

for Effects of different bioreactor systems Deoxyschisandrin 4.28  10 1 for 9.71  10 2 for on biomass production and TS 30 d TS 30 d accumulation of Gomisin A 3.49  10 1 for 7.90  10 2 for schisandra lignans TS 30 d TS 30 d

Shoot

Leucojum aestivum

Galanthamine

1.7 for 0.30 vvm 4.86  10 2 for Studies on biomass [70] production and and 22  C 0.30 vvm and accumulation of 22  C galanthamine in the “modified bubblecolumn bioreactor”

Shoot

Leucojum aestivum

Galanthamine

6.80  10

1

3.29  10

2

[46]

Scale-up of biomass [69] and galanthamine production

(continued)

210

A. Krol et al.

Table 3 (continued) Medium

Experiment parameters

Type of Volume system [ml] Salts

Photoperiod [h light/ Sucrose PGR [g L 1] [mg L 1] h dark]

Light intensitya Temper- Timespan Air flow (TS) rate Air sparger [μmol ature [vvm] constructions m 2 s 1]a (T) [ C] [days]

BB

1000

SH

30

IBA 5.0; ABA 10.0

16/8

80  8

25  2

60

0.8

Aeration provided by a silicon hose from the bottom of the growth vessel; two stainless steel baskets with the smaller one placed concentrically inside the larger basket

Bb

2000

MS

30

GA3 4.0

16/8

35

25

42

0.1

NS

BB

1000

MS

30

TDZ 1.1

16/8

35

23

25

0.1

Sparger placed at the bottom of the container

BB

1000

MS

15; 30; 45; 60

–

16/8

35

23

25

NS

Sparger positioned at the bottom of container

NA data not available, NS data not specified a Unit conversion in accordance with Sager and McFarlane [112] b Expressed as a v/v c Expressed as a mg% FW d Expressed as a gallic acid equivalents [mg GAE g 1 DWor mg GAE L 1 d 1] e Expressed as mg of (+)-catechin equivalent [mg g 1 DW or mg L 1 d 1]

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211

Plant culture

Max. content [mgg 1 DW]

Productivity [mg L 1 d 1]

Genistin

6.94  101 for TS 40 d

1.38  103

Genistein

6.74  10 TS 26 d

Daidzein

3.29 for TS 50 d 7.99  101

Daidzin

1.65  101 for TS 54 d

4.35  102

Total Eleutherosides

1.48  10

1

NA

2

NA

Type

Species

Metabolite

Coculture of shoots and hairy roots

Genista tinctoria

Somatic Eleutherococcus embryos sessiliflorus

1

Chlorogenic acid 3.40  10

Plantlets Hypericum perforatum

Shoot

Hypericum perforatum

for 7.94

1.84  10

Total phenolics

1.67  101

Hypericin

3.36  10

2

4.51  10

3

Pseudohypericin

5.24  10

1

7.01  10

2

Hyperforin

4.04

5.44  10

1

Hypericin

3.18  10 2 for 1.5  10 2 for 30 g/l sucrose 30 g/l sucrose

Pseudohypericin Hyperforin

Ref

Effect of prototype [38] basket-bubble bioreactor on biomass production and accumulation of isoflavones

Total flavonoids

1

Parameters studied

NA NA

Effect of different bioreactor systems on growth and eleutherosides production

[119]

Effects of different culture systems on hypericin, pseudohypericin, and hyperforin contents

[50]

Effects of CO2 and [49] sucrose enrichment 5.08  10 1 for 1.50  10 1 for on biomass production and 45 g/l sucrose 45 g/l sucrose secondary 5.35  10 1 for 2.97  10 1 for metabolites 15 g/l sucrose 15 g/l sucrose accumulation

212

4.3

A. Krol et al.

Temporary and Continuous Immersion Systems

Temporary immersion bioreactors (TIBs) were designed with the aim of overcoming problems associated with submerged cultures, such as anoxia, abnormal morphology including hyperhydricity, and low survivability of the explants. The discussed systems share the advantages of agar cultures, like lack of mechanical stress and good gas exchange, and liquid cultures which include better nutrients availability, easier handling, and scalability. Temporary immersion systems have been developed since 1980s and gradually became the most popular bioreactor type employed in micropropagation studies [28, 103, 105, 120, 121]. They have been extensively reviewed by Etienne and Berthouly [103] and, more recently, by Watt [121] and Georgiev et al. [105]. In the basic configuration, often referred to as the “twin flask” system, the TIB consists of two containers, one serving as medium reservoir and the other one being the growth vessel which is usually equipped with some sort of support for the shoots. The medium is periodically moved between the two vessels, usually by air pump/s which also provide gas exchange for the culture. The twin flask system offers several advantages: it is simple, scalable up to 10–20 l volume, and customizable. Given the above, this type of TIB is one of the most commonly used, and its various iterations have been successfully employed for micropropagation [103, 105, 120, 121] and secondary metabolite studies (Table 4). Over the years, TIBs have become simpler and easier to handle, and numerous systems are commercially available at the moment. The currently used bioreactors are usually made of transparent plastics and include two compartments (i.e., medium reservoir and growth container) in a single vessel. The systems are pneumatically driven which minimizes equipment and maintenance costs, whereas the single-vessel design simplifies handling and saves shelf space [103, 105]. The most popular and successful of TIBs is RITA ® (Vitropic, France), presented in 1995 [105, 120, 121] and still available on the market after the 25th anniversary of its introduction. Despite relatively low volume (usually ca. 0.2 l of growth medium), the system shares all advantages of temporary immersion culture and thus has been often used in studies on secondary metabolism of plants (Table 4). The more recently introduced [122] Plantform system (Plant Form AB, Sweden) has also been employed in experiments on secondary metabolite accumulation in shoot cultures [44–46, 48, 60]. As in the case of pneumatically agitated bioreactors, attempts were made to develop the low-cost, disposable TIBs for economically viable production of plant material. The example of such a system is the “Box-in-Bag” (BIB) bioreactor which shares the advantages of rigid culture vessels and bag bioreactors [32, 105, 120]. Similarly to the twin flask system, the BIB consists of two containers: the 10 l disposable growth vessel and reservoir accommodating 5 l of growth medium. The flat design and vertical distribution of the plant material ensure good access to light which is crucial for the production of high quality plantlets. The “Box-in-Bag” bioreactor offers scale-up possibilities and is intended to improve the effectiveness of micropropagation. So far, it has been employed in industrial propagation of Coffea sp., but no studies on secondary metabolite content in BIB-grown shoots have been conducted [32, 105, 120]. Another type of disposable bioreactor is the

8

Bioreactor-Grown Shoot Cultures for the Secondary Metabolite Production

213

previously mentioned LifeReactor which can be converted to twin flask-type temporary immersion system by simply connecting two separate bags [120]. In terms of plantlets quality and micropropagation effectiveness, TIBs are considered superior to continuous immersion bioreactors (CIBs). Nevertheless, in some instances, continuously submerged cultures showed better growth [43] as compared to shoots grown in TIBs. Some of the custom-made designs included periodic aeration of the continuously submerged culture and thus can be classified as hybrid pneumatically agitated/continuous immersion systems (CIB-SAB or CIB-BB, depending on bioreactor configuration). In experiments on Leucojum aestivum shoot cultures, these bioreactors yielded higher amounts of secondary metabolites as compared to TIB [69]. The commercially available CIBs, like Growtek, are often equipped with supports [43] to prevent the shoots from drowning. However, research has shown that the lack of thereof does not necessarily negatively affect system productivity [69]. Experiments on Ruta graveolens in vitro shoots demonstrated that CIB-grown culture can be scaled up to 5 l without detrimental effects on biomass yield and furanocoumarin content [123]. In order to prevent hypoxia, CIBs can be provided with silicone tubing for bubble-free aeration (BFB bioreactor); however, this solution has been rarely used for in vitro cultures of shoots [40]. The results of studies concerning secondary metabolite production in TIB- and CIB-grown shoot cultures are compiled in Table 4. In many cases, the experiments conducted focused on optimizing immersion frequency which is a crucial parameter affecting productivity of the system. Other strategies aimed at increasing secondary metabolite content, such as elicitation, have also been employed.

4.4

Gas Phase Bioreactors

Similarly to TIBs, the gas phase bioreactors (GPBs) were developed to overcome problems related to continuously submerged organ cultures like poor oxygen transfer, abnormal morphology, and (in some cases) mechanical stress. However, while the former were designed specifically for shoot cultures [103, 105], the gas phase systems were hairy and adventitious root cultures in mind [25, 104, 130]. Nevertheless, the assets of GPBs such as good gas exchange and complete lack of mechanical stress proved to be useful for shoot cultivation, providing good growth and higher secondary metabolite production as compared to other systems [54, 60]. In GPBs, the medium is applied in dispersed form onto the immobilized biomass, placed on a support in a ventilated growth container. Technically, depending on the droplet size, the gas phase systems can be divided into mist (0.01–10 μm) and spray (10–103 μm) bioreactors [104]. However, the size of droplets is rarely provided in experimental papers focused on secondary metabolism of shoot cultures. Therefore, in the current study, the systems were classified according to the mode of medium application: the dispersion can be achieved by using a hydraulic nozzle [46, 60, 131], atomizer nozzle [59, 65], or ultrasonic mist generator [52, 54]. The nomenclature of the systems varies, and similar systems are often referred to as mist, sprinkle, spray, or droplet bioreactors. The details of the so far conducted experiments concerning secondary metabolite production in GPB-grown shoots are presented in Table 5.

214

A. Krol et al.

Table 4 Secondary metabolite production in shoot cultures grown in temporary and continuous immersion bioreactors Medium

Experiment parameters

Volume [ml] Salts

Photo period [h light/ Sucrose PGR [g L 1] [mg L 1] h dark]

Light intensitya Temper- Timespan Immersion time [μmol ature (TS) ratio (ITR) m 2 s 1]a (T) [ Cz] [days] [min/h]

RITA

250

MS

30

BA 0.5

16/8

60

25  1

28; 56

1/3; 1/6

RITA

200

B5

30

–

16/8

77

25  2

4; 9; 21

5/24

RITA

200

B5

30

–

16/8

103

25  2

21

30/3; 30/12; 30/24; 5/3; 5/12; 5/24

RITA

500

MS

30

TDZ 1.10

16/8

35

23

25

5/3

RITA

500

MS

30

TDZ 1.10

16/8

35

23

25

5/30

Bioreactor type

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215

Plant culture Immobilization and Constructions details

Type

Species

Metabolite

–

Shoot

Camptotheca acuminata

–

Shoot

Castilleja tenuiflora

–

Shoot

Castilleja tenuiflora

Max. content [mgg 1 DW]

Productivity Parameters [mg L 1 d 1] studied

Ref

Camptothecin

2.60 for ITR 1/6 and TS 28 d

NA

[66]

Total phenolics

2.95  101bfor 1.40b for N N 1.32 mM 1.32 mM and TS 21 d and TS 21 d

Verbascoside

2.83  102 for 1.35  101 for N 1.32 N 0.66 mM mM and and TS 21 d TS 21 d

Isoverbascoside

9.00  101 for 4.64 for N N 1.32 mM 1.32 mM and TS 21 d and TS 21 d

Aucubin

1.25  101 for 6.90  10 1 for N 25.74 N 0.00 mM mM and and TS 21 d TS 21 d

Total phenolics

3.06  101a for NA ITR 30/24

Total flavonoids 4.58  10–2c for ITR 30/24

Plantlets Hypericum perforatum

Temporary Plantlets Hypericum root zone perforatum immersion

Effects of different culture systems on camptothecin content

Effects of [36] different immersion cycles on morphogenesis and total phenolics and total flavonoids content

NA

3.39  10

2

2.33  10

3

Pseudohypericin 5.41  10

1

3.75  10

2

Hyperforin

2.20  10

3

1.52  10

1

Hypericin

4.47  10

2

3.57  10

3

Pseudohypericin 8.80  10

1

6.94  10

2

3.29  10

3

2.56  10

1

Hypericin

Hyperforin

Effects of [124] nitrogen deficiency and starvation on morphogenesis and secondary metabolism

Effects of [50] different culture systems on the hypericin, pseudohypericin, and hyperforin contents Effects of different culture systems on the hypericin,

[50]

(continued)

216

A. Krol et al.

Table 4 (continued) Medium

Experiment parameters

Bioreactor type

Volume [ml] Salts

Photo period [h light/ Sucrose PGR [g L 1] [mg L 1] h dark]

RITA

200

MS

60

–

16/8

NS

18; 22; 26; 30

35

15/6; 15/8; 15/10; 15/12

RITA

200

MS

60

–

16/8

NS

18; 22; 26

35

15/4; 15/6; 15/7; 15/10; 15/12

RITA

150

MS

30

IBA 0.25

16/8

100

22  1

28

1/12

RITA

200

MS

30

NAA 1.15; BA 2.0

16/8

110

26

35

15/6; 15/12; 15/24

RITA

200

SH

–

2iP 5.0

16/8

40

24  2

28

5/1.5

Light intensitya Temper- Timespan Immersion time [μmol ature (TS) ratio (ITR) m 2 s 1]a (T) [ Cz] [days] [min/h]

8

Bioreactor-Grown Shoot Cultures for the Secondary Metabolite Production

217

Plant culture Immobilization and Constructions details

Type

Species

Metabolite

Max. content [mgg 1 DW]

Productivity Parameters [mg L 1 d 1] studied

Ref

pseudohypericin, and hyperforin contents Shoot

Shoot

Leucojum aestivum

Leucojum aestivum

3.79  10–2 for ITR 15/8 and T 26

Galanthamine

1.06  10 1 for ITR 15/8 and T 26

Lycorine

7.83  10 4 for ITR 15/10 and T 26

Norgalanthamine

2.60  10 1 for ITR 15/8 and T 22

Galanthamine

NA – immersion time ratio and temperature affected biomass production and alkaloids accumulation

Hamayne Lycorine

Effects of [68] immersion frequency and 2.43  10 4 temperature for ITR 15/10 on biomass and alkaloids and T 26 production 2 5.54  10 for ITR 15/8 and T 22 Effects of [71] immersion frequency and temperature on production of alkaloids compounds

Shoot

Ocimum basilicum

Rosmarinic acid 1.55  102

Shoot

Pancratium maritimum

Haemanthamine

9.00  10 1 for ITR 15/12

Lycorine

8.00  10 1 for ITR 15/12

1.03  10 1 Effects of for ITR 15/12 immersion 9.45  10 2 frequency on for ITR 15/12 biomass and alkaloids content

5.00  10–3d for total essential oil

3.57  10–3d for total essential oil

Shoot

Rhododendron Essential oil tomentosum p-cymene Alloaromadendrene Shyobunone Ledene oxide (II)

1.07

Effect of [43] different types of bioreactors on biomass production and rosmarinic acid content [105]

Effects of [60] culture conditions and bioreactor construction on the growth and volatile oil production

(continued)

218

A. Krol et al.

Table 4 (continued) Medium

Experiment parameters

Volume [ml] Salts

Photo period [h light/ Sucrose PGR [g L 1] [mg L 1] h dark]

Light intensitya Temper- Timespan Immersion time [μmol ature (TS) ratio (ITR) m 2 s 1]a (T) [ Cz] [days] [min/h]

RITA

200

SH

–

2iP 5.0

16/8

40

24  2

28

5/1.5

RITA

NS

MS

30

–

16/8

75

23

28

5/3

RITA

200

MS

–

BA 3.0; NAA 1.0

16/8

88  8

24  1

30; 60

5/1.5

RITA

200

MS

–

BA 3.0; NAA 1.0

16/8

90  2

24  1

30; 60

NS

Bioreactor type

8

Bioreactor-Grown Shoot Cultures for the Secondary Metabolite Production

219

Plant culture Immobilization and Constructions details

Max. content [mgg 1 DW]

Productivity Parameters [mg L 1 d 1] studied

5.67  10–3d for total essential oil with aphid extract elicitation

3.93  10–3d for total essential oil with aphid extract elicitation

Effects of biotic and abiotic elicitation on the production of essential oil

[61]

Baicalin

2.75  10

3

NA

2.10  10

3

NA

Wogonin

1.05  10

4

NA

Effect of different bioreactor system on biomass production and accumulation of bioactive metabolites

[40]

Baicalein

Schisandrin

7.86  10 1 for TS 30 d

1.94  10 1 for TS 30 d

[46]

Deoxyschisandrin

5.03  10 1 for TS 30 d

1.24  10 1 for TS 30 d

Gomisin A

5.27  10 1 for TS 30 d

1.30  10 1 for TS 30 d

Effects of different bioreactor systems on biomass production and accumulation of schisandra lignans

Total phenolics

3.27  10 1 for TS 30 d

8.15  10 2 for TS 30 d

[44]

Chlorogenic acid 2.32  10 2 for TS 30 d

5.79  10 3 for TS 30 d

Gallic acid

8.87  10 2 for TS 30 d

2.21  10 2 for TS 30 d

Protocatechuic acid

9.87  10 2 for TS 30 d

2.46  10 2 for TS 30 d

Salicylic acid

5.27  10 2 for TS 30 d

2.46  10 2 for TS 30 d

Total flavonoids 2.13  10 1 for TS 30 d

5.31  10 2 for TS 30 d

Effects of different bioreactor systems and cultivation mode on biomass production and accumulation of phenolic acids and flavonoids

1.85  10 1 for TS 30 d

4.62  10 2 for TS 30 d

Type

Species

Shoot

Rhododendron Essential oil tomentosum p-cymene Alloaromadendrene Shyobunone Ledene oxide (II)

Plantlets Scutellaria baicalensis

Shoot

Shoot

Schisandra chinensis

Schisandra chinensis

Metabolite

Quercitrin

Ref

(continued)

220

A. Krol et al.

Table 4 (continued) Medium

Experiment parameters

Volume [ml] Salts

Photo period [h light/ Sucrose PGR [g L 1] [mg L 1] h dark]

Light intensitya Temper- Timespan Immersion time [μmol ature (TS) ratio (ITR) m 2 s 1]a (T) [ Cz] [days] [min/h]

RITA

200

MS; ½MS

30

BA 1.5; NAA 0.5

16/8

15

23  2

3; 6; 12; 18

3/6

RITA

300

½MS

30

–

16/8

32.5

25  2

60

15/24

Plantform NS

SH

–

2iP 5.0

16/8

40

24  2

28

5/1.5

Plantform 500

MS

–

BA 3.0; NAA 1.0

16/8

88  8

24  1

30; 60

5/1.5

Bioreactor type

8

Bioreactor-Grown Shoot Cultures for the Secondary Metabolite Production

221

Plant culture Immobilization and Constructions details

Type

Species

Metabolite

–

Shoot

Thapsia garganica

Thapsigargin

3.37 for ½ MS NA and MeJa elicitation

Nortrilobolide

2.15  101 for NA ½ MS and MeJa elicitation

Swertiamarin

1.31  101 for NA fructose addition

Gentiopicrin

2.18  101 for NA fructose addition

Sweroside

1.22  101 for NA fructose addition

Shoot

Zeltnera beyrichii

Max. content [mgg 1 DW]

Productivity Parameters [mg L 1 d 1] studied

Ref

Effect of [125] biotic and abiotic elicitation and reduced nutrient supply on the production of thapsigargins Experiments [126] on the accumulation of xanthones and secoiridoid glycosides in bioreactor grown shoots

Shoot

Rhododendron Essential oil 1.91  10–3d for total tomentosum p-cymene essential oil Alloaromadendrene Shyobunone Ledene oxide (II)

NA

Effects of [60] culture conditions and bioreactor construction on the growth and volatile oil production

Shoot

Schisandra chinensis

2.45  10 1 for TS 30 d

Effects of different bioreactor systems on biomass production and accumulation of schisandra lignans

Schisandrin

1.18 for TS 30 d

[46]

(continued)

222

A. Krol et al.

Table 4 (continued) Medium

Bioreactor type

Experiment parameters

Volume [ml] Salts

Photo period [h light/ Sucrose PGR [g L 1] [mg L 1] h dark]

Light intensitya Temper- Timespan Immersion time [μmol ature (TS) ratio (ITR) m 2 s 1]a (T) [ Cz] [days] [min/h]

Plantform 500

MS

–

BA 3.0; NAA 1.0

16/8

90  2

24  1

30; 60

NS

Plantform 500

MS

–

BA 3.0; NAA 1.0

24/0

12.6

25  2

30

5/1.5

Silicone NS tubing bioreactor

MS

30

–

16/8

75

23

28

60/1

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Bioreactor-Grown Shoot Cultures for the Secondary Metabolite Production

223

Plant culture Immobilization and Constructions details

Type

Shoot

Shoot

Species

Schisandra chinensis

Schisandra chinensis cultivar Sadova

Bubble free Plantlets Scutellaria aeration of baicalensis the medium through silicone tube

Metabolite

Max. content [mgg 1 DW]

Productivity Parameters [mg L 1 d 1] studied

Deoxyschisandrin

7.77  10 1 for TS 30 d

1.60  10 1 for TS 30 d

Gomisin A

6.79  10 1 for TS 30 d

1.40  10 1 for TS 30 d

Total phenolics

3.46  10 1 for TS 30 d

7.18  10 2 for TS 30 d

Chlorogenic acid 2.98  10 2 for TS 30 d

6.19  10 3 for TS 30 d

Gallic acid

7.45  10 2 for TS 30 d

1.53  10 2 for TS 30 d

Protocatechuic acid

1.09  10 2 for TS 30 d

2.26  10 2 for TS 30 d

Salicylic acid

5.84  10 2 for TS 30 d

1.21  10 2 for TS 30 d

Total flavonoids 2.05  10 1 for TS 30 d

4.25  10 2 for TS 30 d

Quercitrin

1.75  10 1 for TS 30 d

3.64  10 2 for TS 30 d

Schisandrin

1.44 for yeast extract elicitation

NA

Angeloylgomisin Q

2.23  10 for yeast extract elicitation

1

NA

Gomisin A

4.26  10 for yeast extract elicitation

1

NA

Angeloylgomisin H

4.51  10 for yeast extract elicitation

1

NA

Baicalin

1.53  10

3

NA

Baicalein

4.00  10

4

NA

Wogonin

6.36  10

5

NA

Ref

Effects of [44] different bioreactor systems and cultivation mode on biomass production and accumulation of phenolic acids and flavonoids

Effect of yeast extract on secondary metabolite production

[48]

Effect of [40] different bioreactor system on biomass production and accumulation of bioactive metabolites

(continued)

224

A. Krol et al.

Table 4 (continued) Medium

Experiment parameters

Volume [ml] Salts

Photo period [h light/ Sucrose PGR [g L 1] [mg L 1] h dark]

Light intensitya Temper- Timespan Immersion time [μmol ature (TS) ratio (ITR) m 2 s 1]a (T) [ Cz] [days] [min/h]

Custommade

NS

SH

–

2iP 5.0

16/8

40

24  2

28

5/90

Custommade

500

MS

30

BA 2.0; NAA 0.15

16/8

30

24  2

56

5/4

Custommade

200

MS

30

IAA 2.00; BA 4.00 6.00 8.00

24/0

42

25  2

28

5/3; 15/12

Custommade

250

MS

30

BA 1.0; IAA 0.1

16/8

125–150

27  2

28

2/2; 2/4; 2/6; 2/12

Custommade

250

MS

30

BA 1.0; IAA 0.1

16/8

125–150

27  2

28

2/4

Bioreactor type

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Bioreactor-Grown Shoot Cultures for the Secondary Metabolite Production

225

Plant culture Immobilization and Constructions details

Productivity Parameters [mg L 1 d 1] studied

Ref

Species

Shoot

Rhododendron Essential oil NA 2.92  10 3 for essential oil tomentosum p-cymene content Alloaromadendrene Shyobunone Ledene oxide (II)

Shoot

Leucojum aestivum

Galanthamine

6.00  10

Shoot

Gynura procumbens

Total flavonoid content

3.07  101 for ITR 15/12 and MS medium IAA 2.0 and BA 8.00

2.74  10 1 for ITR 15/12 and MS medium IAA 2.0 and BA 8.00

Effects of [127] growth regulators and immersion frequency on production of biomass and flavonoid compounds

Shoot

Digitalis purpurea

Digitoxin

3.25  10 2 for ITR 2/2

9.57  10 3 for ITR 2/2

Digoxin

2.06  10 2 for ITR 2/4

1.71  10 2 for ITR 2/4

Lanatoside C

ND

ND

The effect of [79] immersion frequencies on biomass and cardenolides accumulation

Lanatoside C

3.16  10 1 for Chitoplant elicitation

Digoxin

2.84  10 2 for Chitoplant elicitation

Digitoxin

ND

1.53  10 1 Effect of [80] for Chitoplant elicitation with elicitation Chitoplant, 1.71  10 2 Silioplant, and for Chitoplant methyl jasmonate on elicitation biomass and ND cardenolides accumulation

Shoot

Digitalis lanata

Metabolite

Max. content [mgg 1 DW]

Type

2

1.44  10

Effects of [60] culture conditions and bioreactor construction on the growth and volatile oil production 3

Effects of [69] different types of bioreactors and inoculum weight on the production of the biomass and galanthamine content

(continued)

226

A. Krol et al.

Table 4 (continued) Medium

Experiment parameters

Volume [ml] Salts

Photo period [h light/ Sucrose PGR [g L 1] [mg L 1] h dark]

Light intensitya Temper- Timespan Immersion time [μmol ature (TS) ratio (ITR) m 2 s 1]a (T) [ Cz] [days] [min/h]

Custommade

1000

MS

30

KIN 0.2; BA 0.3

20/4

NS

14  1

14

3/4

Custommade

2000

½ MS

20

NAA 1.0; GA3 1.0

16/8

50

26

56

5/4

Custommade

2000

MS

30

BA 0.3

16/8

50

26

56

5/4

Custommade

2000

MS

30

BA 1.0

16/8

50

26

56

5/4

Custommade

2000

MS

30

BA 1.0

16/8

50

26

56

5/4

Bioreactor type

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227

Plant culture Immobilization and Constructions details

Type

Species

Plantlets Deschampsia antarctica

Productivity Parameters [mg L 1 d 1] studied

Metabolite

Max. content [mgg 1 DW]

Shikimic acid

1.71 for UV-B NA

Vanillic acid

3.97  10 1 for UV-B + B

NA

Chlorogenic acid 6.49 for UV-B + B

NA

Scopoletin

7.81 for UV-B + B

NA

Rutin

1.40 for UV-B + B

NA

Gallic acid

2.01 for UV-B + B

NA

Isoquercitrin

3.15  10 1 for UV-B + B

NA

Ref

Effects of UV-B elicitation on phenolic metabolites production

[128]

Shoot

Fabiana imbricata

Oleanolic acid

1.00  10

2

NA

Effects of different culture systems on the oleanolic acid content

[111]

Shoot

Cymbopogon citratus

α- citral

3.50  10

1

NA

5.40  10

1

NA

Effects of different culture systems on the α- citral and β-citral contents

[111]

β-citral

Shoot

Hypericum perforatum

Hypericin

1.80  10

1

NA

Effects of different culture systems on the hypericin content

[111]

Shoot

Lavandula officinalis

Rosmarinic acid

5.70

NA

Effects of different culture systems on the rosmarinic acid and essential oil contents

[111]

(continued)

228

A. Krol et al.

Table 4 (continued) Medium

Experiment parameters

Volume [ml] Salts

Photo period [h light/ Sucrose PGR [g L 1] [mg L 1] h dark]

Light intensitya Temper- Timespan Immersion time [μmol ature (TS) ratio (ITR) m 2 s 1]a (T) [ Cz] [days] [min/h]

Custommade

2500

MS

30

BA 0.3

16/8

50

26

42; 56

5/4

Custommade

300

SH

15

IBA 1.0

24/0

88

24  1

32; 60

45/0.75

Custommade

2000

MS

30

GA3 4.0

16/8

35

25

42

30/6

Custom- 200; made (the 400 “‘DualVessel System”)

MS

30

BA 30

16/8

60

25  1

28; 56

1/3; 1/6

Bioreactor type

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229

Plant culture Immobilization and Constructions details

Max. content [mgg 1 DW]

Productivity Parameters [mg L 1 d 1] studied

9.7 for 5 L vessel and TS 42 db

NA

Total flavonoids 5.10 for reduced ITRc

NA

Type

Species

Metabolite

Shoot

Cymbopogon citratus

Total phenolics

Caffeic acid

1.65  101 for NA 10 L vessel and TS 42 d

Chlorogenic acid 1.94  102 for NA reduced ITR

Shoot

Cyclopia genistoides

Somatic Eleutheroembryos coccus sessiliflorus

Neochlorogenic acid

1.19  102for reduced ITR

p-hydroxybenzoic acid

9.90  101 for NA 10 L vessel and TS 42 d

Mangiferin

2.62  101 for 5.48 for TS TS 60 d 60 d

Isomangiferin

7.55 for TS 60 d

1.58 for TS 60 d

Iriflophenone 6.50 for TS 3-C-b-glucoside 32 d

3.04 for TS 32 d

Total Eleutherosides

NA

NA 1.48  10 1 for germinated embryo

NA Chlorogenic acid 3.40  10 2 for germinated embryo Total flavonoids 1.84  101 for NA germinated embryo

Shoot

Camptotheca acuminata

Ref

Effects of [129] gassing with CO2-enriched air, reduced immersion frequency, vessel size and culture time on total phenolic and flavonoid content

Total phenolics

1.67  101 for NA germinated embryo

Camptothecin

2.12 for ITR 1/3 and TS 56 d

NA

Effects of different liquid culture systems on accumulation of phenolic secondary metabolites

[41]

Effect of [119] different bioreactor systems on biomass growth and eleutherosides production

Effects of different culture systems on camptothecin and 10-hydro camptothecin contents

[66]

(continued)

230

A. Krol et al.

Table 4 (continued) Medium

Experiment parameters

Photo period [h light/ Sucrose PGR [g L 1] [mg L 1] h dark]

Light intensitya Temper- Timespan Immersion time [μmol ature (TS) ratio (ITR) m 2 s 1]a (T) [ Cz] [days] [min/h]

MS

–

–

16/8

60

24  1

56

NS

MS

–

–

16/8

60

24  1

56

NS

Custom- NS made, largemouth glass culture vessel 5 L

MS

–

–

16/8

40

24  2

90

60/1

Custom- NS made, glass culture vessel 2 L

MS

–

–

16/8

40

24  2

90

60/1

CIB-BB

NS

MS

30

BA 2.0; NAA 0.15

16/8

30

24  2

56

60/1

CIB-SAB 500

MS

30

BA 2.0; NAA 0.15

16/8

30

24  2

56

60/1

Bioreactor type

Volume [ml] Salts

Custom- 1000 made, balloon1000 type bioreactor

NA data not available, NS data not specified, ND not detected a Unit conversion in accordance with Sager and McFarlane [112] b Expressed as a gallic acid equivalents [mg GAE g 1 DW or mg GAE L 1 d 1] c Expressed as a quercetin equivalents [mg CE g 1 DW or mg CE L 1 d 1] d Expressed as [μlg 1 DW or μlL 1 d 1] e Expressed as mg of (+)-catechin equivalent [mg g 1 DW or mg L 1 d 1]

8

Bioreactor-Grown Shoot Cultures for the Secondary Metabolite Production

231

Plant culture Immobilization and Constructions details

Type

Species

Metabolite

Max. content [mgg 1 DW]

Productivity Parameters [mg L 1 d 1] studied

Shoot

Rosa rugosa

Total phenolics

2.38  101b

3.14 b

Total flavonoids 8.00e Bioreactor Shoot with net

Rosa rugosa

1.04 e

Total phenolics

2.58  10

1b

Total flavonoid

1.10  10

1e

9.64  10–1b 3.93  10–1e

Shoot

Ruta graveolens

Furanocoumarins 2.25  101 for NA Bergapten total furanoPsoralen coumarins Xanthotoxin

Shoot

Ruta graveolens

Furanocoumarins 8.67 for total Bergapten furanoPsoralen coumarins Xanthotoxin

Gassing Shoot four times per day for 5 min

Leucojum aestivum

Galanthamine

7.20  10

Bioreactor Shoot aerated 12 times daily for 5 min

Leucojum aestivum

Galanthamine

2.40

1

Ref

Effects of [37] different bioreactor systems on biomass production and the production of total phenolics and flavonoids Scale-up in liquid culture system

[123]

[69]

NA

2.89  10

2

Scale-up of biomass and galanthamine production

2.15  10

1

Effects of [69] different types of bioreactors and inoculum weight biomass production and galanthamine content

232

A. Krol et al.

Table 5 Secondary metabolite production in shoot cultures grown in gas phase bioreactors Medium Immobilization and constructions details Ultrasonic mist generator; bioreactor with a stainless steel mesh support Ultrasonic mist generator; bioreactor with clay stones support Ultrasonic mist generator; bioreactor with three-tier stainless mesh support Hydraulic “‘Multiplate radius-flow bioreactor”; constant medium circulation without misting, three-tier plate support Ultrasonic mist generator; “‘Modified inner-loop ultrasonic nutrient mist bioreactor” with three-tier stainless steel mesh support Bioreactor with hydraulic nozzle; medium circulation supplied with peristaltic pump

Experiment parameters

Volume [ml] Salts

Photoperiod Sucrose PGR [h light/ [g L 1] [mg L 1] h dark]

Light intensitya Temper- Timespan ature [μmol (TS) m 2 s 1]a (T) [ C] [days]

Misting cycle (MC)ON/ OFF [min/min]

1000

Gam- 30 borgs B5

–

16/8

100

25  2

28

10/50

1000

Gam- 30 borgs B5

–

16/8

100

25  2

28

10/50

400

MS

30

NAA 0.05; BA 0.5

16/8

42

25  1

25

3/90

400

MS

30

NAA 0.05; BA 0.5

16/8

42

25  1

25

–

400

MS

30

NAA 0.05; BA 0.5

12/12

42

25  1

5 10 15 20 25 30

3/30; 3/60; 3/90; 3/120; 3/150; 3/180

NS

MS

30

IAA 0.1; BA 1.0

24/0

40

26  2

21; 28

40/1.5

8

Bioreactor-Grown Shoot Cultures for the Secondary Metabolite Production

233

Plant culture

Liquid flow rate [ml/min] Type

Species

Metabolite

Productivity Max. content [mg L 1 Parameters [mgg 1 DW] d 1] studied

2.0

Shoot

Artemisia annua

Artemisinin

3.00  10–1b NA

2.0

Shoot

Artemisia annua

Artemisinin

3.60  10–1b NA

NS

Shoot

Artemisia annua

Artemisinin

3.60  10–1b 1.89

2.0

Shoot

Artemisia annua

Artemisinin

3.20  10–1b 1.38

NS

Shoot

Artemisia annua

Artemisinin

[52] 3.20  10–1b 1.88 for TS Effect of for TS 25 d 25 d and MC misting cycle on growth and MC 3/90 3/90 and artemisinin accumulation

112

Shoot

Centaurium erythraea

Gentiopicroside Sweroside

2.20  101 for TS 21 d 8.00 for TS 21 d 6.00 for TS 28d

Swertiamarin

9.67 for TS 21 d 3.48 for TS 21 d 2.5 for TS 28 d

Ref

[82] Effect of different mist bioreactor systems on biomass production and accumulation of artemisinin [54] Effect of different bioreactor systems on in vitro biomass production and accumulation of artemisinin

Effect of mist [84] bioreactor system on the production of biomass and biologically active secoiridoid glucosides

(continued)

234

A. Krol et al.

Table 5 (continued) Medium Immobilization and constructions details

Experiment parameters

Volume [ml] Salts

Photoperiod Sucrose PGR [h light/ [g L 1] [mg L 1] h dark]

Light intensitya Temper- Timespan ature [μmol (TS) m 2 s 1]a (T) [ C] [days]

Misting cycle (MC)ON/ OFF [min/min]

MS

20

–

NS

NS

25

60

60/60

MS

40

–

NS

NS

25

60

60/60

MS

–

BA 1.0; IAA 0.1

16/8

40

26  2

28

0.67/3

Bioreactor with NS hydraulic nozzle, medium circulation by peristaltic pump

SH

–

2iP 2.0

16/8

40

24  2

28

5/85

Bioreactor with 1000 hydraulic nozzle, medium circulation by peristaltic pump

MS

–

IAA 0.1; BA 0.45

16/8

40

26  2

21

0.67/0.83

500 Bioreactor supplied with hydraulic nozzle, medium circulation by peristaltic pump

MS

–

BA 3.0; NAA 1.0

16/8

60

24  1

30; 60

0.5/10

12,000 Atomizer mist generator; glass fermenters with two stainless steel meshes 12,000 Atomizer mist generator; glass fermenters with two stainless steel meshes Bioreactor with 1000 hydraulic nozzle, medium circulation by peristaltic pump

8

Bioreactor-Grown Shoot Cultures for the Secondary Metabolite Production

235

Plant culture

Liquid flow rate [ml/min] Type 200

200

90

Species

Metabolite

Productivity Max. content [mg L 1 Parameters [mgg 1 DW] d 1] studied

Transformed Mentha shoot citrata

Linalool Linalyl acetate

3.80  10 6.91  10

Transformed Mentha shoot piperita

Pulegone Menthofuran

2.34  10 6.78  10

Shoot

Total phenolic content Total flavonoid content

5.87  101c

NA

1.94  101d

NA

Rehmannia glutinosa

3 2

1 2

5.28  10 2.43  10

1.13 3.21  10

3 1

1

100

Shoot

Rhododendron Essential oil tomentosum p-cymene Alloaromadendrene Shyobunone Ledene oxide (II)

5.30  10 3 (total essential oil content)

NA

105

Shoot

Salvia officinalis

Rosmarinic acid Carnosic acid Carnosol

2.62  101

2.81

1.74 1.34

1.87  10 1.44  10

Schisandrin

8.52  10 1 for TS 30 d 5.79  10 1 for TS 30 d 5.25  10 1 for TS 30 d

2.19  10 1 for TS 30 d 1.49  10 1 for TS 30 d 1.35  10 1 for TS 30 d

100

Shoot

Schisandra chinensis

Deoxyschisandrin Gomisin A

1 1

Ref

[59] Effects of temperature, light intensity and medium composition on the growth and the production of monoterpenes Effects of culture conditions and bioreactor construction on the growth and volatile oil production Effects of culture conditions and bioreactor construction on the growth and volatile oil production Effect of the nutrient sprinkle bioreactor system on rosmarinic acid and diterpenes accumulation Effects of different bioreactor systems on biomass production and accumulation of schisandra lignans

[35]

[60]

[131]

[46]

(continued)

236

A. Krol et al.

Table 5 (continued) Medium Immobilization and constructions details

Experiment parameters

Volume [ml] Salts

Photoperiod Sucrose PGR [h light/ [g L 1] [mg L 1] h dark]

Light intensitya Temper- Timespan ature [μmol (TS) m 2 s 1]a (T) [ C] [days]

Misting cycle (MC)ON/ OFF [min/min]

500 Bioreactor supplied with hydraulic nozzle, medium circulation by peristaltic pump

MS

–

BA 3.0; NAA 1.0

16/8

90  2

24  2

30; 60

0.5/10

NS Bioreactor supplied with hydraulic nozzle, medium circulation by peristaltic pump

MS

30

IAA 0.1; BA 0.5

16/8

40

26  2

35

0.67/2

Ultrasonic mist generator

MS

30

–

12/12

40

25  1

25

3/90

400

NA data not available, NS data not specified Unit conversion in accordance with Sager and McFarlane [112] Expressed in % c Expressed as a gallic acid equivalents [mg GAE g 1 DW or mg GAE L 1 d 1] d Expressed as a quercetin equivalents [mg CE g 1 DW or mg CE L 1 d 1] a

b

8

Bioreactor-Grown Shoot Cultures for the Secondary Metabolite Production

237

Plant culture

Liquid flow rate [ml/min] Type 100

Shoot

Productivity Max. content [mg L 1 Parameters [mgg 1 DW] d 1] studied

Species

Metabolite

Schisandra chinensis

Total phenolics acid Chlorogenic acid Gallic acid

3.96  10 1 for TS 30 d

Artemisinin

3.5  10–1b

90

Shoot

Scutellaria alpina

500

Shoot

Artemisia annua

3.75  10 2 for TS 30 d 8.87  10 2 for TS 30 d Protocatechuic 1.22  10 1 acid for TS 30 d Salicylic acid 6.74  10 2 for TS 30 d Total 2.90  10 1 flavonoids for TS 30 d Quercitrin 2.10  10 1 for TS 30 d Baicalin 1.52  101 Verbascoside 6.95 Wogonoside 4.08 Luteolin 5.15 Luteolin-71.45 glucoside

Ref

[44] 1.02  10 1 Effects of for TS 30 d different bioreactor 9.70  10 3 systems and for TS 30 d cultivation 2.29  10 2 mode on for TS 30 d biomass production 3.16  10 2 and for TS 30 d accumulation 1.74  10 2 of phenolic for TS 30 d acids and 7.51  10 2 flavonoids for TS 30 d 5.43  10 2 for TS 30 d [132] 7.11  10 1 Effects of 3.23  10 1 liquid 1.91  10 1 systems for shoot 2.41  10 1 multiplica2 6.86  10 tion, secondary metabolite production and plant regeneration [53] 1.88 Effect of novel mist bioreactor system on biomass production and accumulation of artemisinin

238

A. Krol et al.

The major disadvantage of GPBs is that they are relatively more complex as compared to pneumatically driven or temporary immersion systems, and thus more prone to defects. However, the mist bioreactors have been improved over the years by using more reliable ultrasonic transducers [104]. Contrary to TIBs, the gas phase systems employed in the studies are usually custom made. The low-cost GPBs, such as the 50 l LCMB (Low-Cost Mist Bioreactor) by Rootec, Germany [6, 25, 31], have been designed, but they have not been used for shoot cultivation. As in the case of other types of bioreactors, the scale-up of gas phase systems is problematic because of poor illumination of the biomass. This phenomenon was already noticeable in relatively small (14 l) fermenters used to grow transformed shoots of Mentha sp.: the limited access to light resulted in altered composition of the essential oil, as compared to small-scale studies [59]. Another problem associated with GPBs is an inefficient use of the reactor volume: in most instances, the growing biomass was distributed on a single shelf [35, 46, 60, 84, 131, 132]. The multitier systems can be used but their inoculation is time consuming [52, 54]. The hybrid bubble column-gas phase bioreactors enable more effective use of reactor’s space. First, the system is run in the BCB mode and the inoculum is evenly dispersed onto multiple anchor points during the BCB-phase. The medium is drained afterwards and the bioreactor is switched into the GPB mode. The pilot-scale (500 l) system of this type was originally devised by Wilson for the cultivation of Datura stramonium hairy roots [6, 25, 28] and more recently applied in smaller scale for transformed root cultures of Hyoscyamus niger [133]. Given that the shoot inoculum often tends to float on the medium surface, the above-described system would likely not work for shoot cultures. In order to reduce production cost and save floor space, Fei and Weathers [134, 135] came up with the design of vertically oriented, low-cost GPB. The inoculum (Daucus carota cells or Artemisia annua leaf fragments) was immobilized on polylysine-coated polypropylene strips hanging inside the bioreactor. The forming plantlets were then grown using the misting system [134, 135]. On rare occasions, the non-misting systems with hydraulic medium circulation (hydraulic bioreactor, HB) were applied. The example is the “multiplate radius-flow” bioreactor employed for the cultivation of Artemistia annua shoots [54].

5

Comparative Studies on Bioreactor Performance

The studies on secondary metabolite production in shoots often employed different types of bioreactors, in order to select the system providing the highest yields of the desired compounds. The examples include experiments on shoot cultures of Artemisia annua [54] Scutellaria baicalensis [40], Hypericum perforatum [50], Rosa rugosa [37], Leucojum aestivum [69], Schisandra chinensis [44, 46] and Rhododendron tomentosum (Ledum palustre) [60]. In the above mentioned studies, in vitro shoot cultures were maintained using different reactor configurations and examined for biomass growth and secondary metabolite content. The results of comparative research are summarized in Table 6.

8

Bioreactor-Grown Shoot Cultures for the Secondary Metabolite Production

239

Table 6 Comparative studies on secondary metabolite production in shoot cultures grown indifferent types of bioreactors Plant name Artemisia annua Hypericum perforatum Hypericum perforatum Leucojum aestivum Ocimum basilicum Rhododendron tomentosum Rosa rugosa Rosa rugosa Schisandra chinensis

Compound Artemisinin Hypericin Hyperforin Galanthamine Rosmarinic acid Essential oil Total phenolics Total flavonoids Total lignans

Parameter Productivity (mg l 1 d 1) Productivity (μg l 1 d 1) Productivity (μg l 1 d 1) Productivity (μg l 1 d 1) Productivity (mg l 1 d 1) Productivity (μl l 1 d 1) Productivity (mg l 1 d 1) Productivity (mg l 1 d 1) Productivity (mg l 1 d 1)

Schisandra chinensis

Total phenolic acids

Productivity (mg l 1 d 1)

Schisandra chinensis

Total flavonoids

Productivity (mg l 1 d 1)

Scutellaria baicalensis

Total flavones

Content (μg g 1)

Relative performance of the tested systemsa GPB (1.89) > HB (1.38) > ALB (0.58) BB (4.51) > TIB (2.33–3.57) BB (543.9) > TIB (151.8–256.1) BB (40.81) > CIB-SAB (6.86–18.86) > TIB (1.44) SAB (0.88–1.07) > TIB (0.82–1.05) GPB (3.67) > TIB (1.16–3.57) BB (9.93–17.00) > TIB (0.96–3.14) BB (3.34–5.75) > TIB (0.39–1.04) TIB (0.94–1.20) > GPB (1.03–1.06) > SAB (0.44–0.91) > BB (0.53–0.60) BB (0.10–0.11) > GPB (0.08–0.10) > TIB (0.07–0.08) > SAB (0.05–0.08) GBP (0.05–0.08) > SAB (0.03–0.06) > BB (0.05) > TIB (0.04–0.05) TIB (4.96) > SAB (2.05) > CIB (2.00) > BB (1.85)

Reference [54] [50] [50] [69] [43] [60] [37] [37] [46]

[44]

[44]

[40]

a

Reported/calculated values in parentheses, bioreactor nomenclature according to list of abbreviations

Pneumatically agitated bioreactors and temporary immersion systems have been the most commonly used, followed by gas-phase bioreactors. In most cases, the available data enabled to calculate the volumetric productivity of the system (Table 6). The conducted experiments provided mixed results, and none of the bioreactor types can be considered as unequivocally superior in terms of secondary metabolite accumulation. Moreover, general conclusions cannot be drawn since the comparative studies on shoot cultures of a specific plant species are all single reports which have not been reproduced by other research groups. Another problem with data interpretation is caused by the fact that the tested systems were not necessarily run using the optimal settings. Parameters such as aeration rate in pneumatically mixed bioreactors and immersion or spraying frequency in temporary immersion and

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gas phase installations are crucial for system’s performance [3, 28, 102–105]. However, in comparative studies these parameters are usually not optimized for individual bioreactor types. Given the above, in order to maximize and scale-up secondary metabolite production in shoot cultures of a specific plant, the suitable bioreactor still has to be selected empirically.

6

Conclusions

The collected data indicate that bioreactor-grown shoots can be employed as a source of variety of secondary metabolites including alkaloids, phenolics, and volatiles. Studies demonstrated that in vitro shoot cultures can be successfully maintained in different types of bioreactors; however, scaling-up of these systems presents substantial challenge. Light requirements and poor growth of shoots in fully submerged, dense cultures limit practical size of a single unit to 10–20 l and impose horizontal distribution of the biomass. Given this, production scale-up can be more effectively achieved by employing multiple small-size vessels. This approach also requires single units to be cheap, stackable, and easy to operate which favors temporary immersion systems. Besides pneumatically agitated and gas phase bioreactors, TIBs have been often employed in secondary metabolite studies. Several systems of this type are available commercially and research on developing low-cost, disposable TIBs is ongoing. As far as secondary metabolite yields are concerned, bioreactor-grown shoot cultures accumulate relatively low amounts of compounds. So far, in vitro shoots are not commercially viable as a source of chemicals. However, they offer a stable platform for the production of certain constituents which can be exploited in the case of shortage of natural resources. Acknowledgments The study was supported by the project POWR.03.02.00-00-I014/17-00 co-financed by the European Union through the European Social Fund under the Operational Programme Knowledge Education Development 2014–2020

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Production of Specific Flavonoids and Verbascoside in Shoot Cultures of Scutellaria baicalensis Beata Kawka, Inga Kwiecień, and Halina Maria Ekiert

Contents 1 2 3 4 5 6

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Chemical Composition of the Baikal Skullcap Root . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Traditional Applications of the Baikal Skullcap . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Biological Activity of the Root Extracts and Isolated Compounds . . . . . . . . . . . . . . . . . . . . . . Biogenesis of the Studied Groups of Compounds and Their Characteristic . . . . . . . . . . . . . Review of the Research on In Vitro Cultures of Scutellaria baicalensis . . . . . . . . . . . . . . . . . 6.1 Micropropagation of Scutellaria baicalensis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.2 Genetic Transformation of Scutellaria baicalensis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7 Accumulation of Secondary Metabolites in In Vitro Cultures of Scutellaria baicalensis: Studies from Our Laboratory . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8 Stationary In Vitro Cultures of Scutellaria baicalensis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.1 Cultures Grown on Murashige and Skoog Medium . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.2 Cultures Grown on Linsmaier and Skoog Medium . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9 Agitated In Vitro Cultures of Scutellaria baicalensis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9.1 Cultures Grown in Murashige and Skoog Medium . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9.2 Cultures Grown in Linsmaier and Skoog Medium . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10 Administering of Biosynthetic Precursors of Phenolic Compounds . . . . . . . . . . . . . . . . . . . . . . 10.1 Stationary Cultures of Scutellaria baicalensis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10.2 Agitated Cultures of Scutellaria baicalensis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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B. Kawka · I. Kwiecień (*) Chair and Department of Pharmaceutical Botany, Jagiellonian University, Medical College, Kraków, Poland e-mail: [email protected]; [email protected] H. M. Ekiert Department of Pharmaceutical Botany, Faculty of Pharmacy, Jagiellonian University, Medical College, Kraków, Poland e-mail: [email protected]; [email protected] © Springer Nature Switzerland AG 2021 K. G. Ramawat et al. (eds.), Plant Cell and Tissue Differentiation and Secondary Metabolites, Reference Series in Phytochemistry, https://doi.org/10.1007/978-3-030-30185-9_7

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11 Elicitation of In Vitro Cultures of Scutellaria baicalensis: Agitated Cultures . . . . . . . . . . . 12 Combined Strategies: Simultaneous Addition of Elicitor and Biosynthetic Precursors – Agitated Cultures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 13 In Vitro Cultures of Scutellaria baicalensis in Bioreactors: Preliminary Research . . . . . . 14 Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Abstract

Scutellaria baicalensis (Baikal skullcap) has been used for centuries in traditional Chinese medicine (TCM). The root of the Baikal skullcap (Scutellariae baicalensis radix) is also a recognized raw material in European therapeutic practice. It is mentioned in European Pharmacopoeia since 2010 and also in the WHO monograph. This raw material is a rich source of specyfic flavonoids, especially from the group of flavones, such as baicalein, baicalin, wogonin, and wogonoside. These compounds show a number of valuable biological activities. The aim of the presented research was to propose in vitro cultures of Scutellaria baicalensis as an alternative to the raw material obtained from cultivated plants for providing apart from flavonoids also phenylpropanoid glycosides and phenolic acids. Standard biotechnological treatments, such as a selection of the composition of the basal medium, selection of the concentration and mutual proportions of plant growth regulators, and different types of culture cultivation (solid, agitated and bioreactor’s cultures), were used. In addition, elicitation and feeding with biosynthetic precursors were tested. The optimization of culturing conditions led to increased accumulation of species-specific flavonoids. Additionally in vitro cultures proved to be a particularly rich source of the phenylpropanoid glycoside - verbascoside. Keywords

Baikal skullcap · Scutellaria baicalensis · In vitro cultures · Scutellaria-specific flavonoids · Verbascoside · Precursors feeding · Elicitors Abbreviations

BAP DW HPLC LS MS NAA PGRs TCM WHO

6-Benzylaminopurine Dry weight High-Performance Liquid Chromatography Linsmaier and Skoog Murashige and Skoog α-Naphthaleneacetic acid Plant Growth Regulators Traditional Chinese Medicine World Health Organization

9

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Introduction

The best-known species of the genus Scutellaria is Scutellaria baicalensis Georgi (Baikal skullcap). This species has its natural habitats in the regions of Eastern Russia, Mongolia, and China. It grows even at an altitude of 8000 m because it is resistant to temperatures down to 15  C. It is a perennial plant species [1]. The Latin generic name comes from the shape of the calyx, which resembles a bowl or tray, and was first used by the botanist Jean Bauhin [2] (Fig. 1a, b). In China, where the Baikal skullcap has been used for millennia in traditional medicine, it is called “Huang-Qin” [3], while in Japanese Kampo medicine, it is called Ogon or Ougon [4]. It is a plant species valued and widely used in Asia, both on its own and in mixtures with other plants. The raw material used for medicinal purposes is the root – Scutellariae baicalensis radix [5]. The mention of the skullcap root first appeared in Chinese Shennong’s Classic of Materia Medica around 100 BC [6]. The root as a medicinal raw material was also described in Bencao Gangmu (Compendium of Materia Medica) published in China in 1593, during the Ming dynasty [7]. Monographs of this raw material can now be found in many pharmacopoeias, including the Chinese (Pharmacopoeia of the People’s Republic of China) [8], Japanese [9], and Korean [10] ones, and also in The International Pharmacopoeia published by the World Health Organization (WHO) [1]. Since 2010, its monograph has also been included in the Supplement 7.1 of European Pharmacopoeia [11]. According to the requirements of the European Pharmacopoeia, the root should be harvested from plants at least 3 or 4 years old. The baicalin content of the dried plant raw material should not be less than 9.0% (Fig. 1c) [5]. Because of its qualities, Scutellaria baicalensis is grown throughout Asia [12]. In other countries, this species is grown on experimental plantations and in botanical gardens [6, 13, 14]. For the medicinal purpose in Europe, roots are imported most of all from China.

Fig. 1 Scutellaria baicalensis shoots (a), flowers (b), and dry roots (raw material) (c)

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Due to the fact that this species is a highly valued medicinal resource almost all over the world, biotechnological studies of S. baicalensis have long been conducted using all available methods. The aim of our research is to achieve endogenic accumulation of bioactive metabolites at a level that will allow to propose in vitro cultures of this species as an alternative to the raw plant material for providing those compounds.

2

Chemical Composition of the Baikal Skullcap Root

The Baikal skullcap root is a flavonoid-rich raw material. It contains, according to various references, from more than 40–125 compounds from this group, mainly flavones but also flavonols, flavanones, chalcones, and biflavonoids [15–17]. Some of them, like apigenin or luteolin, are popular in the plant kingdom, but others are very rare and specific only for Scutellaria genus. However, only a few of them occur in the root of the plant in a concentration high enough to be responsible for pharmacological activity. The most important active compounds are specific for Scutellaria genus flavonoids, e.g., baicalein, wogonin, oroxylin A, scutellarein, and chrysin along with their glycosidic forms: baicalin (baicalein 7-O-glucuronide), wogonoside (wogonin 7-O-glucuronide), and 7-O-glucuronide of oroxylin A (Structures 1) [17]. The main group of flavonoid glycosides specific for the genus Scutellaria, found in S. baicalensis, are glucuronides. Structures 1 The chemical structure of Scutellaria baicalensis specific flavonoids (aglycones)

O HO

O

CH3

HO

O

HO OH

O

OH

wogonin

baicalein HO

O

HO H3C

OH

O

O

O

O OH

chrysin

O

oroxylin A OH

HO

O

HO OH

O

scutellarein

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In addition to flavonoids, the raw material also contains iridoid glycosides, diterpenoids, phenylpropanoid glycosides like verbascoside or leucosceptoside A, and other groups of compounds, such as essential oil, tannins, phenolic acids, steroids, and polysaccharides [3]. The concentration of flavonoid compounds in the raw material is highly variable and can range from 1% to more than 20%, with that of baicalin being the highest. The largest amounts of flavonoid compounds are found in the raw material of Chinese origin [18]. The skullcap leaves have a composition similar to that of the root, but in terms of active compounds, the content is lower and less varied [19].

3

Traditional Applications of the Baikal Skullcap

In Asia, Scutellaria baicalensis has been used for their anti-inflammatory, sedative, antiviral, anticoagulant, and antioxidant properties [4, 16]. In China, where it has been used for over 2000 years in traditional Chinese medicine (TCM), and in Japan (Japanese traditional Kampo medicine), the Baikal skullcap is used on its own or as an ingredient of complex herbal mixtures [19]. It is used to treat attacks of asthma, allergies, inflammations, and food poisoning. In Nepal, compresses of skullcap leaves are used to relieve swelling and heal wounds after insect bites [20].

4

Biological Activity of the Root Extracts and Isolated Compounds

Scutellariae baicalensis radix has a proven strong antioxidant effect as a result of the ability to scavenge free radicals and the ability to chelate metal ions. This effect is related to a number of biological activities of the raw material, including antiinflammatory, antibacterial, antifungal, and antiviral activities [21]. The compound that is responsible for the anti-inflammatory effect of the Baikal skullcap is primarily baicalin. It interacts with inflammation mediators by inhibiting the formation of proinflammatory cytokines and chemokines and by inhibiting the proliferation of T lymphocytes [22]. The anti-free radical activity of skullcap flavones positively affects the circulatory system by lowering blood pressure, inhibiting arterial calcification and platelet aggregation, and consequently, preventing atherosclerosis [1]. Extracts from the Baikal skullcap are used in neurodegenerative diseases such as Alzheimer’s disease and Parkinson’s disease [23, 24]. The neuroprotective effect of this raw material is multidirectional and results from antiapoptotic, anti-inflammatory, and antioxidant activities. Both in vitro and in vivo studies have also demonstrated hepatoprotective and choleretic effects of skullcap flavones. Baicalin, baicalein, and wogonin protected the liver against fibrosis as a result of an infection caused by the hepatitis B and C viruses [25]. Antioxidant properties and those that stimulate the immune system also translate into the adaptogenic effect of the Baikal skullcap [26].

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Currently, the Baikal skullcap is credited with anticancer properties. This activity is associated with antioxidant and anti-inflammatory effects and cell cycle inhibition. Due to the stimulating effect of skullcap root extracts on innate human immunity, they may be important in the treatment of leukemias. In vivo tests have also demonstrated the ability of the skullcap flavones to inhibit the growth of solid tumors, without inducing systemic toxicity [27, 28]. The therapeutic practice also makes use of the Baikal skullcap leaves, from which mostly an infusion is prepared. It has mainly a sedative effect and thus can be used both in the treatment of neurosis and insomnia [29]. The compound responsible for the sedative and anxiolytic effects of the skullcap is probably wogonin, which acts via the GABA A receptor, joining the binding site for benzodiazepines [30, 31].

5

Biogenesis of the Studied Groups of Compounds and Their Characteristic

Our research focused on three groups of metabolites: flavonoids, phenolic acids, and phenylpropanoid glycosides. They are compounds with a phenolic structure, whose biosynthetic pathways are biogenetically related to one another. Flavonoids are polyphenolic compounds widely distributed in plants. The biosynthesis of flavonoids is a complex process involving two metabolic pathways: the malonic acid pathway, in which malonyl-CoA is formed from glycolysis-derived acetyl-CoA followed by the formation of one of the rings, and the shikimic acid pathway through which shikimic acid is obtained and then the aromatic amino acids – phenylalanine or tyrosine. The key stage in flavonoid biosynthesis is the condensation of 4-coumaroyl-CoA with three molecules of malonyl-CoA catalyzed by chalcone synthase. The product of the reaction is an unstable 15-carbon chalcone acting as the skeleton of flavonoids [32]. Subsequently, in the presence of chalcone isomerase, it undergoes stereospecific cyclization to (2S)-flavanone – naringenin. Flavanones are the starting compounds for the synthesis of other flavonoid groups [33]. Apart from popular in the plant kingdom flavonoids, in the root of the Baikal skullcap, the most abundant are specific for Scutellaria genus. They are lipophilic flavones with triple hydroxylation of the ring (baicalein and wogonin) and their glycosidic linkages with glucuronic acid (baicalin and wogonoside) [14]. Their biosynthesis proceeds differently, using cinnamoyl-CoA instead of 4-coumaroylCoA as a substrate, leading to the formation of an intermediate product, which is pinocembrin [34]. In the human body, flavonoids exhibit diverse pharmacological activity. Their most important properties include antioxidant, chemopreventive, cardioprotective, hepatoprotective, vasoprotective, platelet anti-aggregation, hypotonic, spasmolytic, diuretic, anti-inflammatory, and antiallergic effects [35]. Flavonoids are components of numerous preparations used as auxiliaries in the treatment of heart and blood vessel diseases, liver diseases, and alleviation of menopausal symptoms (isoflavones) [36]. Baikal skullcap-specific flavonoids have been proved to possess, most of all, anti oxidative, anti-inflammatory, antibacterial, and hepatoprotective properties [1].

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Phenolic acids belong to secondary metabolites commonly found in plants. Depending on the structure of the side chain, phenolic acids are divided into derivatives of benzoic acid, cinnamic acid, and phenylacetic acid. Especially common among plants are cinnamic acid derivatives such as caffeic acid, ferulic acid, 4-coumaric acid, and sinapic acid. The best-known derivatives of benzoic acid include 4-hydroxybenzoic, salicylic, gallic, protocatechuic, vanillic, and syringic acids [37]. The third group of phenolic acids are represented by 3-hydroxyphenylacetic acid and 3,4-dihydroxyphenylacetic acid. Phenolic acids occur in plants mainly in bound form, as esters or glycosides and structural elements of lignins or hydrolyzing tannins. Phenolic acids can be formed via several biosynthetic pathways: the shikimic acid pathway, the chorismic acid pathway, the polyketide pathway, and through deamination of aromatic amino acids. The basic route for the biosynthesis of phenolic acids is the biogenetic pathway leading through shikimic acid and chorismic acid. The latter is a precursor of aromatic amino acids – tryptophan, tyrosine, and phenylalanine (Scheme 1) [38]. Phenylalanine is the basic precursor of hydroxycinnamic acids. The conversion of this aromatic amino acid to hydroxycinnamic acid derivatives takes place in three stages (the so-called phenylpropanoid pathway). Another precursor of phenolic acids, but less often, is tyrosine. Like phenylalanine, it is formed from chorismic acid (Scheme 1) [38]. Phenolic acids are characterized by a broad spectrum of biological activity. They exhibit, for example, antibacterial, antifungal, antiviral, anti-inflammatory, antipyretic, immunostimulatory, hepatoprotective, and choleretic effects. As natural antioxidants, they are responsible for the removal of free radicals, inhibition of the formation of reactive oxygen or nitrogen species, and chelation of metal ions. They are counted among chemopreventive compounds and those protecting against the development of coronary heart disease and type 2 diabetes [39]. Phenylpropanoid glycosides are composed of phenolic compounds linked to several sugar molecules. The best known of them is verbascoside. Because phenolic acids (mainly caffeic acid) are a part of these glycosides, the biosynthesis also takes place via the phenylpropanoid pathway (Scheme 1) [40]. These compounds are very powerful antioxidants; they also have anti-inflammatory, analgesic, immunosuppressive, immunomodulatory, antineoplastic, hepatoprotective, and antimicrobial, especially antiviral effects [41].

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Review of the Research on In Vitro Cultures of Scutellaria baicalensis

6.1

Micropropagation of Scutellaria baicalensis

Plants growing in the soil are very sensitive to environmental conditions, which results in changes in their phytochemical profile. In addition to phenotypic variability, genetic diversity within a species is also an important factor. The inhomogeneity of the plant raw material obtained from natural habitats and cultivated crops is a

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Shikimic acid Shikimic acid pathway

Chorismic acid

Trp

Phe

Tyr TAT

PAL TAL

4-hydroxyphenylpyruvic acid Cinnamic acid

4-Coumaric acid

HPPR

4-hydroxyphenylacetic acid

4CL 4-Coumaroyl-CoA C4H

Phenolic acids

Phenolic acids CHS

+ malonyl-CoA

Chalcone CHI

Flavonoids Scheme 1 Biosynthetic pathways of phenolic metabolites in plants. C4H cinnamate-4-hydroxylase, CHI chalcone isomerase, CHS chalcone synthase, 4CL 4-coumaroyl:CoA-ligase, HPPR 4hydroxyphenylpyruvate reductase, PAL phenylalanine ammonialyase, Phe phenylalanine, TAL tyrosine ammonialyase, TAT tyrosine aminotransferase, Tyr tyrosine, Trp tryptophan

major problem because it results in a lower quality of the raw material. Therefore, protocols for micropropagation of S. baicalensis are being developed that allow obtaining homogeneous material, identical to the parent plant, and creation of collections of highly productive plants [42]. The research on the micropropagation of in vitro cultures of Scutellaria baicalensis has been focused on the impact of various plant growth regulators (PGRs) and culture

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conditions (lighting, temperature). The most effective production of lateral buds was obtained on Murashige and Skoog (MS) medium with the addition of α-naphthaleneacetic acid (NAA) and kinetin. To increase the rate of multiplication, a two-step procedure was used. First was obtained induction of callus with organogenetic potential, followed by shoot regeneration using 6-benzylaminopurine (BAP) and then rooting on half-strength MS [43]. Thidiazuron also proved effective in the induction of shoot regeneration from intact seedlings, leaves, and petioles [44].

6.2

Genetic Transformation of Scutellaria baicalensis

Numerous studies have been conducted on the genetic transformation of the Baikal skullcap in order to obtain hairy root cultures, due to the fact that it is a species accumulating biologically active metabolites in the roots [45–47]. Regarding the efficiency of the transformation process, the selection of the appropriate strain of Agrobacterium rhizogenes has proved to be very important. The choice of explant and site of infection are also important. The researchers then focus their attention on the selection of the growth medium and production medium. It has been shown, for example, that hairy roots grown in the Schenk and Hilde-brandt medium supplemented with 4% and 6% sucrose showed, after optimization, a 20- to 30-fold increase in the biomass of hairy roots after 4 weeks of culture [48]. All the cultures of transformed roots showed greater biomass growth than those of untransformed roots and accumulated significant amounts of flavonoids characteristic of the skullcap roots. The amount of baicalin was 14.1–30.0% of the biomass dry weight (DW). Generally, it was higher than the maximum amount in untransformed roots ~18%. The amount of wogonin varied from 0.08% to 0.18% depending on the line of hairy root culture, but in each case, it was higher than in untransformed roots ~0.07% [49]. It has also been shown that the presence of auxins increased the production of flavonoids in hairy root cultures of Scutellaria baicalensis. Indolylacetic acid was the most conducive to the accumulation of baicalin and baicalein, while the highest level of wogonin was observed in the presence of indolylbutyric acid [50]. Hairy root cultures have also been subjected to elicitation procedures. The accumulation of baicalin in transformed roots was increased by exposure to various elicitors such as methyl jasmonate, salicylic acid, and various fungal extracts. The accumulation of baicalin in elicited cultures was from 1.5 to 3 times higher than in control cultures [51, 52].

7

Accumulation of Secondary Metabolites in In Vitro Cultures of Scutellaria baicalensis: Studies from Our Laboratory

In our research, we focused on growing untransformed in vitro cultures of Scutellaria baicalensis as a more economical model. By using different strategies for process optimization, efforts were made to achieve increased accumulation of

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secondary metabolites from selected groups such as flavonoids, phenolic acids, and phenylpropanoid glycosides. During the analyses of methanolic extracts from the biomass of in vitro cultures, 32 flavonoids, including 9 specific to the genus Scutellaria; 25 phenolic acids and their parent compounds benzoic acid, cinnamic acid, and phenylacetic acid; and also verbascoside and isoverbascoside from the phenylpropanoid glycoside group were used as reference substances. A total of 62 compounds were estimated in methanolic extracts by HPLC.

8

Stationary In Vitro Cultures of Scutellaria baicalensis

8.1

Cultures Grown on Murashige and Skoog Medium

Shoot cultures (Fig. 2a) were grown on six variants of the gelled Murashige and Skoog (MS) medium (which was used for establishing of cultures), containing different amounts of cytokinin – 6-benzylaminopurine (BAP) and auxin – α-naphthaleneacetic acid (NAA) in various proportions (BAP/NAA [mg
L 1] – 1.0/1.0, 2.0/2.0, 3.0/1.0, 0.5/ 2.0, 1.0/0.5, 5.0/0.5). Methanolic extracts from biomass harvested after a 4-week growth period were used to estimate the amounts of metabolites by HPLC. Among the tested variants of the media, the highest biomass increase (almost eightfold) was obtained on a medium containing 1.0 mg
L 1 BAP and 1.0 mg
L 1 NAA. Somewhat smaller increases (4.5-fold) were found on 2 media (1.0 mg
L 1 BAP and 0.5 mg
L 1 NAA, and 0.5 mg
L 1 BAP and 2.0 mg
L 1 NAA). The variant containing 5.0 mg
L 1 BAP and 0.5 mg
L 1 NAA proved to be a medium that was totally unfavorable to the growth of S. baicalensis cultures. In the extracts analyzed, five flavonoids from among the metabolites characteristic of the genus Scutellaria were found: baicalein, baicalin, scutellarin, wogonin, and wogonoside. In addition, the presence of verbascoside was confirmed (Table 1). We do not detect any of the analyzed phenolic acids. The total flavonoid content varied, depending on the variant of the medium, from 266.1 to 428.6 mg
100 g 1 DW. Baicalin was the quantitatively dominant compound – max. 315.2 mg
100 g 1 DW. A compound accumulated in particularly high amounts was verbascoside – max. 830.9 mg
100 g 1 DW (Table 1). High amounts of flavonoids (over 350 mg
100 g 1 DW) were obtained in the biomass of shoots growing on three variants of MS medium: BAP/NAA [mg
L 1] 0.5/ 2.0, 2.0/2.0, and 1.0/0.5. The highest amounts of verbascoside were confirmed on the media containing 0.5/2.0 mg
L 1 BAP/NAA (596.1 mg
100 g 1 DW) and 1.0/ 0.5 mg
L 1 BAP/NAA (830.9 mg
100 g 1 DW) (Table 1). Because of the large increases in biomass, these two variants of the medium were proposed as the best “productive” and, at the same time, “growth-promoting” media [53]. Comparing the obtained results with the amounts of compounds estimated in the root plants growing in the M. Koczwara Garden of Medicinal Plants, Jagiellonian University, Medical College, Cracow, Poland, and acquired from commercial sources, the total amounts of flavonoids in the biomass from in vitro cultures are from 2.5 to 6.0 times lower. In the plant raw material, only small amounts of verbascoside (up to 75 mg
100 g 1 DW) and the presence of 3-coumaric and hydrocaffeic acid were

9

Production of Specific Flavonoids and Verbascoside in Shoot Cultures of. . .

259

Fig. 2 Scutellaria baicalensis in vitro cultures: stationary shoot culture (a), agitated shoot culture (b), culture in PlantForm bioreactor (c), and culture in RITA bioreactor (d)

found. Comparing the antioxidant potential of extracts from in vitro biomass and raw material, no significant differences were found, suggesting that shoot cultures of the Baikal skullcap are a promising source of antioxidants.

8.2

Cultures Grown on Linsmaier and Skoog Medium

Media less reach in nutrition ingredients, like Linsmaier and Skoog (LS) medium, may promote the growth of in vitro cultures, so we decided to test it. Shoot cultures were grown on five variants of the LS medium, differing in the concentration of PGRs – BAP and NAA, in the same proportions as in the case of the MS medium (BAP/NAA [mg
L 1] – 1.0/1.0, 2.0/2.0, 3.0/1.0, 0.5/2.0, 1.0/0.5). Because the MS variant containing 5.0 mg
L 1 BAP and 0.5 mg
L 1 NAA did not promote biomass growth, its use was abandoned. The increases in biomass obtained during an analogous (4-week) growth period were very high (from 9.1- to tenfold) on three LS variants (1.0/1.0, 0.5/2.0, 1.0/0.5 – BAP/NAA [mg
L 1]).

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Table 1 Content of phenolic compounds in in vitro cultures of Scutellaria baicalensis grown on two types of basal gelled media with different concentrations of PGRs Metabolite [mg 100 g DW 1] Baicalein Baicalin Scutellarin Wogonin Wogonoside Total flavonoids Verbascoside 3,4-dihydroxyphenylacetic acid

Plant growth regulators – BAP/NAA [mg L 1] 1.0/1.0 2.0/2.0 3.0/1.0 0.5/2.0 MS LS MS LS MS LS MS 10.4 3.0 10.1 1.0 1.0 2.2 16.8 185.2 303.7 230.2 353.5 198.0 273.4 315.2 35.6 55.0 30.9 71.7 35.9 62.3 24.2 5.6 140.7 1.4 25.3 1.0 161.2 16.8 29.3 96.5 99.6 56.4 33.4 88.3 55.6 266.1 598.9 372.2 507.9 269.3 587.4 428.6

LS 0.6 316.4 51.8 18.1 57.3 444.2

1.0/0.5 MS 15.8 293.7 10.0 15.8 21.8 357.1

LS 2.1 327.1 60.4 89.9 159.5 639.0

324.9 671.4 336.5 862.5 390.9 502.0 – 16.7 – 31.5 – 38.5

1003.7 16.5

830.9 –

838.7 19.7

596.1 –



Not detected

The methanolic extracts were found to contain five flavonoids: baicalein, baicalin, scutellarin, wogonin, and wogonoside. In addition, the presence of 3,4dihydroxyphenylacetic acid and verbascoside was confirmed (Table 1). The total flavonoid content ranged from 444.2 to 639.0 mg
100 g 1 DW, depending on the tested variant of LS medium. The main metabolites in this group were baicalin (max. 353.5 mg
100 g 1 DW) and wogonoside (max. 159.5 mg
100 g 1 DW). On the LS medium, the biomass was confirmed to contain small quantities (below 40 mg
100 g 1 DW) of 3,4-dihydroxyphenylacetic acid, along with very high amounts of verbascoside, from 502.2 to 1003.7 mg
100 g 1 DW, with the highest found on the medium variant containing 0.5/2.0 mg
L 1 BAP/NAA (Table 1). This variant was also very good at promoting the growth of S. baicalensis biomass [54]. Comparing the results obtained on the same variants of MS and LS media, the LS medium (despite the fact that it is poorer in organic components than the MS medium) can be nominated as definitely more conducive to the accumulation of flavonoids and verbascoside in the biomass of in vitro cultures. The total amounts of flavonoids were up to 2.25 times higher and those of verbascoside up to 2.56 times higher than on the MS medium [55].

9

Agitated In Vitro Cultures of Scutellaria baicalensis

9.1

Cultures Grown in Murashige and Skoog Medium

Agitated cultures of Scutellaria baicalensis (Fig. 2b) were grown in five variants of MS medium enriched with BAP and NAA in various proportions (BAP/NAA [mg
L 1] – 1.0/1.0, 2.0/2.0, 3.0/1.0, 0.5/2.0, 1.0/0.5) in 4-week cycles. They were the same MS variants that were tested in stationary cultures.

9

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261

Table 2 Content of phenolic compounds in agitated in vitro cultures of Scutellaria baicalensis grown in two types of basal media with different concentrations of PGRs Metabolite [mg
100 g DW 1] Baicalein Baicalin Scutellarin Wogonin Wogonoside Total flavonoids Verbascoside 3,4dihydroxyphenylacetic acid

Plant growth regulators – BAP/NAA [mg
L 1.0/1.0 2.0/2.0 3.0/1.0 MS LS MS LS MS LS 2.7 – 8.1 4.3 4.1 – 151.8 106.7 173.0 171.9 152.6 130.6 24.7 39.7 27.9 30.8 21.2 26.9 7.3 5.3 2.0 1.8 2.0 0.6 95.3 44.3 122.6 45.5 92.6 22.5 287.3 200.3 329.7 250.0 271.1 180.6

1

] 0.5/2.0 MS 10.0 239.4 34.4 56.0 101.4 441.2

810.5 745.3 365.8 532.1 348.3 620.1 537.8 2.8 51.2 4.9 35.0 3.2 16.4 3.3

LS 1.6 217.1 40.4 2.5 59.6 321.2

1.0/0.5 MS 5.2 259.0 27.2 4.7 114.0 410.1

LS 1.9 174.0 82.0 6.0 94.4 358.3

1098.0 26.6

560.7 5.4

1115.4 39.9



Not detected

Large increases in biomass, from 13.4- to 17.3-fold, were obtained on the MS variants tested. Very large increases in biomass (over 15-fold) were confirmed on three variants of the medium enriched with BAP/NAA [mg
L 1] – 1.0/1.0, 3.0/1.0, and 0.5/2.0. The highest increases were obtained on the last mentioned MS variant. In the extracts tested, the same flavonoids as in stationary cultures among characteristic of the genus Scutellaria, were found baicalein, baicalin, scutellarin, wogonin, and wogonoside, and also the phenylpropanoid glycoside – verbascoside. In addition, the presence of 3,4-dihydroxyphenylacetic acid was confirmed (Table 2). The total flavonoid content ranged from 271.1 to 441.2 mg
100 g 1 DW. The compounds that were accumulated in the largest amounts in the biomass grown were baicalin (max. 259.0 mg
100 g 1 DW) and wogonoside (max. 122.6 mg
100 g 1 DW). The maximum verbascoside content was 810.5 mg
100 g 1 DW (Table 2). The medium variants that were the most conducive to the accumulation of the estimated compounds were the same variants as in the case of stationary cultures (0.5/2.0, 2.0/2.0, and 1.0/0.5 BAP/NAA [mg
L 1]) [56]. The total amounts of flavonoids and verbascoside were comparable with those obtained on gelled MS medium. However, the obtained increases in biomass were from 2 to 4.1 times higher in agitated cultures than in stationary cultures.

9.2

Cultures Grown in Linsmaier and Skoog Medium

Agitated cultures of S. baicalensis were grown for a period of 4 weeks on variants of LS medium that were exactly the same as those of MS medium (five variants). The tested variants of LS medium also obtained large increases in biomass, from 8.1- to 14.6-fold. The highest increases in biomass (about 14-fold) were confirmed on two medium variants enriched with BAP/NAA [mg
L 1] – 1.0/1.0 and 1.0/0.5. LS

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“growth-promoting” media variants are also the best “growth-promoting” variants of the MS medium. The analyzed extracts of the biomass from all the medium variants were found to contain four flavonoids, such as baicalin, scutellarin, wogonin, and wogonoside, as well as 3,4-dihydroxyphenylacetic acid and verbascoside. The presence of baicalein in small quantities was confirmed only on three medium variants (Table 2). The total flavonoid content on the tested LS variants was slightly lower than on the MS medium and varied in the range from 180.6 to 358.3 mg
100 g 1 DW. The highest baicalin content was found on the LS medium supplemented with 0.5 mg
L 1 BAP and 2.0 mg
L 1 NAA (217.1 mg
100 g 1 DW). The medium of this composition and another variant with 1.0/0.5 mg
L 1 BAP/NAA were particularly conducive to the accumulation of verbascoside, 1098.0 and 1115.4 mg
100 g 1 DW, respectively (Table 2). These are medium variants that also promote increases in biomass (over 12-fold), so they can serve as “universal” media. A comparison of the results obtained in agitated and stationary cultures maintained on LS media indicates that agitated cultures are characterized by greater increases in biomass, with a similar or higher level of accumulation of the estimated compounds.

10

Administering of Biosynthetic Precursors of Phenolic Compounds

10.1

Stationary Cultures of Scutellaria baicalensis

Shoot cultures of Scutellaria baicalensis were grown on LS medium containing 1.0 mg
L 1 BAP and 0.5 mg
L 1 NAA, selected on the basis of previous experiments as the best “productive” and, at the same time, “growth-promoting” medium for this species. To serve as biosynthetic precursors of phenolic compounds, phenylalanine and tyrosine were administered at a concentration of 1 g
L 1 to the culture medium at the time of establishing an experiment (point “0”). The cultures were grown for a period of 4 weeks. Precursor doses were tested in our laboratory previously. The cultures whose medium had been administered phenylalanine showed 1.45 times larger increases in biomass than the control cultures, whereas the cultures supplemented with tyrosine grew by 10% less well than control. In extracts from the biomass of cultures which had been administered phenylalanine or tyrosine, the same compounds were found as in the biomass from the control cultures, i.e., baicalin, scutellarin, wogonin and wogonoside, and also 3,4dihydroxyphenylacetic acid and verbascoside. The exception was baicalein, the presence of which was confirmed only in extracts from the biomass of cultures supplemented with phenylalanine (Table 3). The total flavonoid content of the biomass from cultures supplemented with phenylalanine was 373.7 mg
100 g 1 DW and from those with tyrosine 548.5 mg
100 g 1 DW, which was, respectively, 1.3- and 1.9-fold higher than in the

9

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263

Table 3 Content of phenolic compounds in stationary in vitro cultures of Scutellaria baicalensis grown on LS medium supplemented with 1.0 mg
L 1 BAP and 0.5 mg
L 1 NAA after administering of biosynthetic precursors – phenylalanine and tyrosine [1 g
L 1] Metabolite [mg
100 g DW 1] Baicalein Baicalin Scutellarin Wogonin Wogonoside Total flavonoids Verbascoside 3,4-dihydroxy-phenylacetic acid

Control – 245.1 17.2 7.4 18.3 288.0 582.6 30.6

Phenylalanine 14.2 286.5 35.3 11.3 26.4 373.7 787.3 41.3

Tyrosine – 390.8 30.7 5.8 121.2 548.5 1172.1 59.3



Not detected

control cultures grown in parallel. Among the flavonoids, the dominant metabolite was, as in the other experiments, baicalin. The verbascoside content of the biomass from cultures supplemented with phenylalanine was 787.3 mg
100 g 1 DW and from those with tyrosine 1172.1 mg
100 g 1 DW, which was 1.35 and 2.0 times higher than in the control cultures (Table 3).

10.2

Agitated Cultures of Scutellaria baicalensis

Because of the large increases in biomass obtained in agitated cultures and the reports in the literature about better absorption of components from a liquid medium, an experiment was conducted with agitated cultures. The agitated cultures of the Baikal skullcap were grown in LS medium containing 1.0 mg
L 1 BAP and 0.5 mg
L 1 NAA. To serve as biosynthetic precursors of phenolic compounds, phenylalanine and tyrosine were administered. The cultures were grown for 2 weeks on the medium without precursors, and then they were added to each flask in appropriate amounts to obtain their final concentrations of 1.0, 1.5, 2.0, and 2.5 g
L 1. The cultures were then continued for a period of 3 or 7 days. The cultures whose medium had been administered phenylalanine showed from 12.0- to 17.2-fold increases in growth on day 3 and from 14.8- to 18.0-fold increases 7 days after the administration of the precursor. By comparison, the cultures supplemented with tyrosine showed from 13.9- to 15.6-fold increases in growth on day 3 and from 14.6- to 17.6-fold increases 7 days after the administration of the precursor. In both cases, the results were comparable with the increases in biomass of the control cultures, which means that both amino acids, in the entire range of the concentrations applied, do not negatively affect the growth of cultures. The increases in biomass after 7 days were up to 1.45 times higher for phenylalanine and up to 1.25 times higher for tyrosine in comparisons to after 3 days. Extracts from the biomass of cultures supplemented with phenylalanine were found to contain baicalein, baicalin, scutellarin, wogonin and wogonoside, and also

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Table 4 The maximal amounts of estimated metabolites in agitated in vitro cultures of Scutellaria baicalensis grown in LS medium supplemented with 1.0 mg
L 1 BAP and 0.5 mg
L 1NAA after administering different concentrations of biosynthetic precursors (phenylalanine and tyrosine) and elicitor (methyl jasmonate) collected after 3 and 7 days Metabolite [mg
100g DW -1] Baicalein Baicalin Chrysin Scutellarin Wogonin Wogonoside Total flavonoids Verbascoside 3,4dihydroxyphenylacetic acid

Control 2.8b 119.0a 0.1b 8.7b 4.2b 46.6a 174.1a

Phenylalanine 6.7a 217.1b –* 34.8b 35.6a 71.2b 302.9b

Tyrosine – 236.7a 15.2b 36.9a 5.6b 69.2b 341.8a

Methyl jasmonate – 217.5a 1.6a 37.8b 1.5a 69.2b 310.3a

330.8a 39.1b

1291.2b 214.5a

2369.4a 128.8a

1906.5b 52.1a

Phenylalanine + methyl jasmonate 6.8a 214.2a 30.1a 32.9b 17.3a 60.8b 303.2a

Tyrosine + methyl jasmonate 18.6b 372.3b 40.0b 96.2b 13.2b 198.9b 729.4b

2675.5b 196.8a

3022.5b 153.5b



Not detected, a3rd day, b7th day

3,4-dihydroxyphenylacetic acid and verbascoside, while extracts from the biomass of cultures supplemented with tyrosine contained baicalin, chrysin, scutellarin, wogonin and wogonoside, and also 3,4-dihydroxyphenylacetic acid and verbascoside (Table 4). The total amounts of the estimated flavonoids after administering phenylalanine ranged from 121.9 to 302.9 mg
100 g 1 DW and after the administration of tyrosine from 172.0 to 341.8 mg
100 g 1 DW (Table 4). Baicalin was the dominant compound of this group in all the extracts. Higher levels of flavonoids were obtained in the cultures grown for 7 days after the addition of phenylalanine and after 3 days in the case of tyrosine. The concentration of the precursors did not have a significant effect on the results obtained. The verbascoside content of extracts was very high. After the administration of phenylalanine, it was max. 480.0 mg
100 g 1 DW on day 3 and 1291.2 mg
100 g 1 DW on day 7. By contrast, after the administration of tyrosine, it was max. 2369.4 on day 3 and 892.3 mg
100 g 1 DW on day 7. A very interesting result, after the administration of either precursor, was the obtaining of three to six times higher levels of 3,4-dihydroxyphenylacetic acid (max. 214.5 mg
100 g 1 DW) than in the control and in all previous experiments (Table 4). This is direct evidence of cellular uptake of aromatic amino acids from the liquid medium and their incorporation into the biosynthetic pathways of phenolic metabolites.

9

11

Production of Specific Flavonoids and Verbascoside in Shoot Cultures of. . .

265

Elicitation of In Vitro Cultures of Scutellaria baicalensis: Agitated Cultures

Agitated cultures were grown in the same variant of LS medium (1.0 mg
L 1 BAP and 0.5 mg
L 1 NAA) that was used for the administration of precursors. Methyl jasmonate was administered as the elicitor. The cultures were grown for 2 weeks in the medium without additives, and then methyl jasmonate was added to each flask in such amounts as to obtain the final concentrations of 10, 50, and 100 μM (doses were tested in our laboratory previously). The cultures were then continued for 3 or 7 days. The increases in biomass ranged from 12.7 to 14.8 times on day 3 and from 13.4 to 15.9 times 7 days after administering methyl jasmonate. The applied concentrations of the elicitor did not affect biomass growth. Extracts from the biomass of cultures elicited for 3 days were found to contain baicalin, chrysin, scutellarin, wogonin and wogonoside, and also 3,4dihydroxyphenylacetic acid and verbascoside. After 7 days of elicitation, the presence of chrysin and wogonin was not confirmed (Table 4). Following the addition of methyl jasmonate, the total amounts of the estimated flavonoids ranged from 125.2 to 310.3 mg
100 g 1 DW after 3 days of culture and from 146.1 to 306.6 mg
100 g 1 DW after 7 days. Higher levels of flavonoids were obtained after using lower concentrations of the elicitor (Table 4). Baicalin was the dominant compound. The verbascoside content of extracts was very high. On day 3, after the administration of methyl jasmonate at a concentration of 10 μM, it was 1350.8 mg
100 g 1 DW. By comparison, on day 7, all the concentrations of the elicitor resulted in verbascoside amounts above 1 g
100 g 1 DW (max. 1906.5 mg
100 g 1 DW; concentration 50 μM). The amounts of 3,4-dihydroxyphenylacetic acid were maintained at the level of the control values (Table 4) [57].

12

Combined Strategies: Simultaneous Addition of Elicitor and Biosynthetic Precursors – Agitated Cultures

We decided to combine both strategies and administer the precursor together with the elicitor. Agitated cultures were grown in LS medium analogous to that in the previous experiments (1.0 mg
L 1 BAP and 0.5 mg
L 1 NAA). The cultures were grown for 2 weeks in the medium without additives, and then each flask received methyl jasmonate at a final concentration of 50 μM and one of the precursors (phenylalanine or tyrosine) at the final concentrations of 1.0, 1.5, 2.0, and 2.5 g
L 1. The cultures were then grown for 3 or 7 days. The increases in the biomass of cultures with phenylalanine and methyl jasmonate in the medium were from 10.6- to 15.6-fold on day 3 and from 12.0- to 17.2-fold 7 days after the administration of the precursor. By comparison, the cultures supplemented with tyrosine together with methyl jasmonate showed from 13.0- to 14.1-fold increases on day 3 and from 11.0- to 14.1-fold increases 7 days after the

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administration of compounds. In both cases, the concentration of the precursor had no effect on biomass growth increments. The administration of the precursors together with the elicitor decreased the biomass growth increments in comparison with the administration of the precursors alone in the cultures grown for 7 days. The compounds present in extracts from the biomass of cultures with the addition of phenylalanine and the elicitor included baicalin, chrysin, scutellarin, wogonin and wogonoside, and also 3,4-dihydroxyphenylacetic acid and verbascoside. After 7 days of experiment, the presence of baicalein was not confirmed. The same qualitative composition was confirmed in extracts from the biomass of cultures that had received tyrosine together with methyl jasmonate (Table 4). The maximum total amounts of flavonoids after the administration of phenylalanine with the elicitor were 303.2 (day 3) and 281.1 (day 7) mg
100 g 1 DW, and after the administration of tyrosine with the elicitor, they were 341.7 (day 3) and 729.4 (day 7) mg
100 g 1 DW (Table 4). In all the extracts, baicalin was the dominant compound in this group (max. 372.3 mg
100 g 1 DW). For both precursors combined with elicitor, the results obtained on day 3 were higher than after the administration of phenylalanine and tyrosine on their own (Table 4). The verbascoside content of extracts was also very high. The maximum verbascoside content obtained on day 7 was 2676.5 mg
100 g 1 DW after the administration of phenylalanine with methyl jasmonate and 3022.5 mg
100 g 1 DW after the administration of tyrosine with methyl jasmonate. Combining the precursors with the elicitor resulted in an 8.8- and 10.0-fold increase in verbascoside content relative to the control and a 3.5- and 4.1-fold increase in comparison with the administration of the precursors on their own. In the case of 3,4dihydroxyphenylacetic acid, higher amounts (max. 196.8 mg
100 g 1 DW) were also obtained, compared with the control samples (Table 4).

13

In Vitro Cultures of Scutellaria baicalensis in Bioreactors: Preliminary Research

Cultures of S. baicalensis were grown in two types of commercially available temporary immersion bioreactors – PlantForm (PlantForm, Sweden) (Fig. 2c) and RITA (VITROPIC, France) (Fig. 2d). In PlantForm bioreactors, variants of LS medium containing 0.5 mg
L 1 BAP and 2.0 mg
L 1 NAA and 1.0 mg
L 1 BAP and 1.0 mg
L 1 NAA were tested. In RITA bioreactors, cultures were grown in LS medium containing 0.5 mg
L 1 BAP and 2.0 mg
L 1 NAA. In both types of bioreactors, biomass was maintained for a period of 30 days. In PlantForm bioreactors, higher increases in dry biomass were recorded on the 1.0/1.0 variant (13.2-fold) than on the 0.5/2.0 variant (10.9-fold). In RITA bioreactors, 9.01-fold increases in biomass were found. All the extracts from the biomass grown in PlantForm bioreactors were found to contain four flavonoids, such as baicalin, chrysin, scutellarin, and wogonoside, and additionally verbascoside. Amounts of chrysin were less than 0.1mg
100g DW. By

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267

Table 5 Content of phenolic compounds in in vitro cultures of Scutellaria baicalensis grown in LS medium supplemented with different concentration of PGRs in two types of bioreactors – PlantForm and RITA

Metabolite [mg
100 g DW 1] Baicalein Baicalin Chrysin Scutellarin Wogonin Wogonoside Total flavonoids Verbascoside 3,4-dihydroxy-phenylacetic acid

PlantForm BAP/NAA [mg
L 1] 0.5/2.0 1.0/1.0 – – 246.0 294.8 – – 50.1 65.0 – – 19.6 12.7 315.7 372.4 561.6 591.2 – –

RITA 0.5/2.0 87.0 624.0 113.7 189.8 75.9 35.1 1125.5 1978.6 124.4

The immersion frequency in both bioreactors was 5 min every 90 min Not detected  Less than 0.1 mg 100 g DW 1 

comparison, the biomass harvested from RITA bioreactors contained baicalein, baicalin, chrysin, scutellarin, wogonin, and wogonoside, as well as 3,4dihydroxyphenylacetic acid and verbascoside (Table 5). In extracts from the biomass grown in PlantForm bioreactors, the total flavonoid content was 315.7 mg
100 g 1 DW and 372.4 mg
100 g 1 DW and was lower than in stationary cultures and comparable to the content in agitated cultures. Baicalin was the quantitatively dominant compound. The verbascoside content, 561.6 and 591.2 mg
100 g 1 DW, was lower than in stationary and agitated cultures. None of the extracts contained 3,4-dihydroxyphenylacetic acid (Table 5). This model of bioreactor cultures requires optimization of the culturing process because, with very good increments in growth, satisfactory amounts of metabolites were not obtained, in comparison with the other types of cultures. In extracts from the biomass growing in RITA bioreactors, the average total flavonoid content was 1125.5 mg
100 g 1 DW and was higher than obtained in stationary and agitated cultures. The dominant compounds from this group were baicalin (624.0 mg
100 g 1 DW) and scutellarin (189.8 mg
100 g 1 DW). In biomass extracts, the verbascoside content (1978.6 mg
100 g 1 DW) was confirmed to be twice as high as in the agitated cultures and the amount of 3,4-dihydroxyphenylacetic acid (124.4 mg
100 g 1 DW) to be 2.5–3.2 times higher than in stationary and agitated cultures, respectively (Table 5). RITA bioreactors, despite somewhat smaller increases in biomass than in PlantForm bioreactors, appear to be, at this stage of research, a better research model because of the richer qualitative composition and high amounts of the metabolites of interest.

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Conclusions

By planning and implementing the successive stages of biotechnological experiments using in vitro cultures of Scutellaria baicalensis, it has been shown that they accumulate lower, but satisfactory, amounts of flavonoids than those present in the medicinal raw material from the plants growing in the open air. Compared with the soil-grown plant, the tested cultures accumulate much higher amounts of the phenylpropanoid glycoside, verbascoside. Whereas from the group of phenolic acids, only one acid – 3,4-dihydroxyphenylacetic acid – is accumulated in in vitro cutures. In all the types of cultures, flavonoids are predominantly accumulated in

Table 6 The maximal content of estimated metabolites in different types of Scutellaria baicalensis in vitro cultures Metabolite Baicalein

Maximal content [mg
100 g DW 1] 87.0

Baicalin

624.0

Chrysin

113.7

Scutellarin

189.8

Wogonin

161.2

Wogonoside

198.9 159.5

Total flavonoids

1125.5

Verbascoside

3022.5 2675.5

3,4dihydroxyphenylacetic acid

214.5 196.8

Type of culture RITA bioreactor culture RITA bioreactor culture RITA bioreactor culture RITA bioreactor culture Stationary culture Agitated culture Stationary culture RITA bioreactor culture Agitated culture Agitated culture Agitated culture Agitated culture

Basal medium and other conditions LS

LS

LS

LS

LS LS, 1.0 g
L 1 tyrosine +50 μM methyl jasmonate; 7th day LS LS

LS, 1.0 g
L 1 tyrosine +50 μM methyl jasmonate; 7th day LS, 1.0 g
L 1 phenylalanine +50 μM methyl jasmonate; 7th day LS, 1.5 g
L 1 phenylalanine; 3rd day LS, 2.5 g
L 1 phenylalanine +50 μM methyl jasmonate; 3rd day

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the form of glycosides (glucuronides characteristic of the genus Scutellaria), mainly baicalin, and also wogonoside and scutellarin. It has also been proved that stationary and agitated cultures grow better and accumulate more metabolites on variants of LS medium, compared with variants of MS medium, and that the quantitative proportions of tested plant growth regulators (BAP and NAA) in the tested variants are of great importance for both biomass growth and metabolite accumulation. It is possible to nominate variants that can serve as both “productive” and “growth-promoting” media. The increases in biomass in agitated cultures and bioreactor cultures are very high. Supplementation of culture media with biogenetic precursors of studied secondary metabolites such as phenylalanine and tyrosine clearly affects the accumulation of all the studied groups of metabolites in shoot cultures of Scutellaria baicalensis, while elicitation of the cultures is particularly conducive to the accumulation of verbascoside. As a result of implementing the combined strategy, simultaneous addition of elicitor and biosynthetic precursors, and process optimization, the amounts of verbascoside increased to a level of about 3 g
100 g 1 DW (Table 6). Scutellaria baicalensis cultures grown in RITA bioreactors are characterized by good biomass increments and high ability to accumulate metabolites, especially verbascoside (Table 6). Success has been achieved in confirming that untransformed in vitro cultures of Scutellaria baicalensis could be a source of the flavonoids characteristic of this species and that such cultures can also be proposed as a very rich source of the phenylpropanoid glycoside – verbascoside. The in vitro cultures of Scutellaria baicalensis are a good model for the investigation on the accumulation of bioactive secondary metabolites.

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Secondary Metabolites in Shoot Cultures of Hypericum

10

Ana Coste, Carmen Pop, Adela Halmagyi, and Anca Butiuc-Keul

Contents 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Pharmaceutical Value of Hypericum Species . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Biotechnological Approaches to Improve Secondary Metabolite Production in Shoot Cultures of Hypericum Species . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1 Plant Growth Regulators and Signaling Compounds . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2 Elicitation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.3 Precursor Feeding . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.4 Genetic Transformation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 Other Factors Affecting Secondary Metabolite Yield in Shoot Cultures of Hypericum Species . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1 Culture System . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2 Genotype and Ploidy Level . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.3 Developmental Stage . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5 Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

275 276 277 278 291 294 294 295 296 296 298 298 299

All authors made equal contribution in this work and are equally considered as first authors. A. Coste (*) · A. Halmagyi Department of Experimental Biology and Biochemistry, Institute of Biological Research Cluj-Napoca, Cluj-Napoca, Rom^ania e-mail: [email protected]; [email protected] C. Pop (*) Department of Drug Industry and Pharmaceutical Biotechnology, Faculty of Pharmacy, University of Medicine and Pharmacy “Iuliu Haţieganu” Cluj-Napoca, Cluj-Napoca, Rom^ania e-mail: [email protected] A. Butiuc-Keul Department of Molecular Biology and Biotechnology, Faculty of Biology and Geology, Babeş-Bolyai University, Cluj-Napoca, Rom^ania e-mail: [email protected] © Springer Nature Switzerland AG 2021 K. G. Ramawat et al. (eds.), Plant Cell and Tissue Differentiation and Secondary Metabolites, Reference Series in Phytochemistry, https://doi.org/10.1007/978-3-030-30185-9_9

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Abstract

The rich phytochemical profile of Hypericum species and the presence of unique compounds, such as hypericins and hyperforin, make them medicinally valuable worldwide. Among Hypericum spp., H. perforatum L. remains the most investigated and exploited species for specific compound production, while others have been surveyed mainly as a source of novel drugs. The main focus of Hypericum biotechnology is the manipulation of secondary metabolism and development of environmentally sustainable and economically viable culture systems toward the efficient production of target and novel secondary metabolites. Since specific compounds accumulate mainly in the aerial parts of Hypericum plants, large-scale shoot cultures are viewed as superior alternative for the production of these compounds under controlled conditions to their extraction from wild- or greenhouse-grown plants or their chemical synthesis. This chapter describes how media culture composition, culture conditions, elicitors, and other critical parameters influence the behavior of Hypericum spp. in shoot cultures and how optimization of these factors could allow improvements in secondary metabolite-related production and discovery of novel drugs. Keywords

Secondary metabolites · Shoot cultures · Hypericins · Elicitation · Hypericum spp. Abbreviations

MS LS B5 PGRs IBA IAA BAP Kin NAA TDZ JA SA MJA DHPJA Z PEG PEC CHI DEX FCM

Murashige and Skoog Linsmaier and Skoog Gamborg B-5 Plant growth regulators Indole-3-butyric acid Indole-3-acetic acid 6-Benzylaminopurine Kinetin 1-Naphthaleneacetic acid Thidiazuron Jasmonic acid Salicylic acid Methyl jasmonate 2,3-Dihydroxypropyl jasmonate Zeatin Polyethylene glycol Pectin Chitin Dextran Flow cytometry

10

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DW FW PI PB spp. H. A.

1

275

Dry weight Fresh weight Partial immersion Paper bridge Species Hypericum Agrobacterium

Introduction

The genus Hypericum (Hypericaceae) is divided into 36 sections including about 500 species of herbs, shrubs, and trees, which either are naturally occurring or have been introduced to every continent in the world, except Antarctica [1, 2]. Species of the genus have been reported to contain a wide range of biologically active compounds that are associated with valuable bioactivities, including neuroprotection [3] and antitumor properties [4]. These plants have been used as traditional medicinals in various parts of the world. Among them, H. perforatum L., also known as St. John’s wort, is one of the world’s top-selling antidepressant [5]. Increasing demand for H. perforatum raw material leads to the excessive exploitation of this species for the extraction of valuable secondary metabolites limiting natural resources and variability of phytochemicals in the wild- and field-grown plants [6, 7]. Since conventional methods are time-consuming and yield of fieldgrown plants is affected by physiological and environmental factors, effective biotechnological methods have been attempted for the improvement of specific secondary metabolite production under controlled conditions [8]. In vitro cell and organ cultures were mainly exploited for the production of target compounds such as naphthodianthrones (hypericins) and acylphloroglucinols (hyperforin), which contribute to the antidepressant activity of Hypericum plant extracts [9, 10]. The development of novel technologies of cell culture and molecular biology, such as cell line selection, cell immobilization, permeabilization, precursor feeding, product secretion, biotransformation, metabolic engineering, bioreactor engineering, synthetic biology, and elicitation, allow many ways to improve secondary metabolite production [11, 12]. However, it was observed that accumulation of hypericins in unorganized cell suspension cultures was inferior compared to organized structures such as seedlings and shoots [13]. This is due to the fact that the major compounds of the genus are known to accumulate mainly in specialized multicellular structures, including translucent glands, dark glands, or secretory canals, which are present in the aerial parts of the plants [14, 15]. Many studies have demonstrated that significant presence of dark glands in various tissues was correlated with the highest concentrations of hypericins [16]. Since most of the important secondary metabolites accumulate in the aerial parts of Hypericum spp. [17, 18] and cell cultures reveal low

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amounts of these compounds [19], large-scale shoot cultures may be a promising approach for their successful bioproduction [20]. Therefore, in this chapter, we discuss the available information on the secondary metabolite production in shoot cultures of Hypericum spp., underline the potential application of different elicitors, and discuss how elicitation-mediated changes may be exploited for novel drug discovery within this genus.

2

Pharmaceutical Value of Hypericum Species

Hypericum spp. are known worldwide for traditional and modern medicinal uses in rural and ethnic communities of different countries and have been reported as an antitumor, antidepressant, antiviral, antimicrobial, anti-inflammatory, and healing agents [9, 20–23]. The vital compounds imparting medicinal properties to Hypericum spp. include naphthodianthrones (e.g., hypericins and pseudohypericin), prenylated acylphloroglucinols (e.g., hyperforin and adhyperforin), xanthones, essential oils, phenolic acids, and a broad array of flavonoids, including flavonols, biflavones, and procyanidins, all of which synergistically contribute to their therapeutic effects [24]. One of the best chemically investigated representatives of the genus is H. perforatum. The use of this medicinal herb as an antidepressant is recognized worldwide, while the efficacy of dry extracts of aerial/flowering parts [Hyperici herba] has been demonstrated in several clinical trials [25]. Recently, the Herbal Medicinal Product Committee belonging to the European Medicines Agency accepted the attributes of this herb and recognized its “well-established” use [26, 27]. In addition to its antidepressant activity, H. perforatum preparations are also externally applied or orally administered for several other ailments, including digestive, urinary, respiratory, and cardiac diseases and gastric ulcer [28, 29]. Due to pronounced bioactivities of H. perforatum extracts, several other representatives of the genus have been surveyed mainly for target compounds or as a source of novel chemicals, with proven biological activity. Thus, Kitanov [30] evaluated the total amount of hypericins in 36 taxa of the genus Hypericum, belonging to 17 distinct sections. Most of these species contained hypericins, with the amount of pseudohypericin usually exceeding that of hypericin. Other studies revealed new pharmaceutically promising species of the genus: H. adenotrichum, H. triquetrifolium, H. montanum, H. montbretti, H. sinarium, H. scabrum, H. sampsonii, and H. barbatum, which were reported to contain even higher levels of hypericins than those registered for H. perforatum [31–34]. Hypericin derivatives were also reported in H. rumeliacum, H. humifusum, H. tetrapterum, and H. annulatum, while phloroglucinol derivatives were found to prevail in H. androsaemum, H. stellatum, and H. kouytchense, all of which are hypericin-lacking species [24]. Generally it was noticed that Hypericum spp. with a higher abundance of naphthodianthrones revealed fewer or no phloroglucinols and vice versa. Other species of the genus, such as H. bupleuroides, H. canariense, H. balearicum, and H. pulchrum, were reported to be rich in flavonoids and

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chlorogenic acid derivatives [24], while H. maculatum was reported to synthesize and accumulate hypericin, essential oils, and flavonoids in similar amounts found in H. perforatum [35–38]. A recent review of Stojanovič et al. [39] revealed the presence of 11 main bioactive compounds in 132 representatives of the Hypericum genus, while Velingkar et al. [23] and Mir et al. [40] reported 10 classes of bioactive compounds in H. perforatum. Moreover, recently several compounds were detected that have not previously been identified in the genus Hypericum. Crockett et al. [41] reported the isolation of a new phloroglucinol derivative, 1-[6hydroxy-2,4-dimethoxyphenyl]-2-methyl-1-propanone, from H. cistifolium and H. galioides, while Kimáková et al. [24] determined the presence of two new anthraquinones in 17 Hypericum spp. other than H. perforatum. In spite of the widespread medicinal values of this genus, only H. perforatum has been intensely characterized in terms of phytochemistry and pharmacology. Though exciting reports on the pharmacological activities of other Hypericum spp. have been published [23, 42], approximately 70% of this genus is yet to be explored. Hence, exploring other species of the genus Hypericum will lead to the identification of novel compounds, and drug leads with new or improved bioactivities.

3

Biotechnological Approaches to Improve Secondary Metabolite Production in Shoot Cultures of Hypericum Species

St. John’s wort has traditionally been supplied from both cultivated and wildharvested materials. However, the concentration of bioactive compounds can vary in plants depending upon the genotype, the type of cultivar, altitude, photoperiod, time of the year, different environmental and growing conditions, processing and preparation of sample materials, and harvesting time [16, 21, 43]. Thus, concerns about variability in quality, adulteration, and contamination, as well as the possibility of losses in biodiversity when collected from the wild [44], have raised the interest in Hypericum biotechnology for the efficient and viable bioproduction of target metabolites. Moreover, a number of known Hypericum constituents are structurally complex and either defiant or difficult to produce via chemosynthesis, or the cost of their synthesis outweighs their commercial availability [45]. Though each group of compounds present in Hypericum spp. is pharmacologically important, mainly three major Hypericum phytochemicals, hypericin, pseudohypericin, and hyperforin, are considered as the main bioactive compounds and considered responsible for the purported antidepressant effect of Hypericum spp. [5]. Hypericin levels were found to vary between 0.03 and 0.3% in leaves and 1 up to 14% in flower buds, while hyperforin content has been reported to reach 6.9% in flower buds, 8.5% in unripe fruits, and 1.5% in leaves [8]. These compounds, however, were proven to accumulate mainly in specialized secretory structures in the aerial parts of the plants [44], though recent research revealed the presence of hypericins in other organs and structures, completely lacking dark glands [46, 47]. Consequently many investigations have been directed at understanding and

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enhancing hypericin and hyperforin production in in vitro shoot cultures of mainly H. perforatum [18, 19, 48, 49], while other representatives have been surveyed for both hypericin production and a source of novel chemicals [50–55]. Various strategies such as optimization of culture conditions, medium composition, elicitation, precursor feeding and genetic transformation have been used to stimulate the production of secondary metabolites in shoot cultures of Hypericum spp. [56–62].

3.1

Plant Growth Regulators and Signaling Compounds

The achievements in the synthesis of secondary metabolites under the effect of PGRs and signaling compounds were intensively investigated in Hypericum spp. (Table 1) [61, 63–65, 87]. It is now well recognized that the secondary metabolite production maximization requires a comprehensive approach combining the selection of a proper combination/concentration of PGRs. A thorough study of Jamwal et al. [88] reviewed the plant growth regulator-mediated synthesis of secondary metabolites in medicinal plants including Hypericum. In H. perforatum the extracts obtained from PGRtreated samples generally showed improved bioactivities compared to the controls [89]. A supplementation of Murashige and Skoog (MS) medium [90] with 3 mg L 1 IBA and 1 mg L 1 Kin led to 0.119 μg mg 1 hypericin in H. perforatum shoots [66]. Santarem and Astarita [63] reported similar hypericin levels (0.45 μg g 1 FW) in H. perforatum shoot cultures (MS with 4.5 μM BAP and 0.05 μM NAA) to fieldgrown plants. Moreover, they have registered a ninefold increase in hypericin concentration versus callus cultures. An enhanced hypericin production (1.7-fold) over the control (MS without PGRs) and pseudohypericin (1.9-fold) was registered in H. maculatum shoots cultured on MS medium supplemented with 0.4 mg L 1 BAP [53]. H. hirsutum shoots cultured on medium with 0.4 mg L 1 2iP, 0.2 mg L 1, BAP 0.1 mg L 1, Kin and 0.05 mg L 1 NAA showed accumulation of hyperforin (1.54 μg g 1 DW) representing up to 6.16-fold over the controls. Surprisingly, a decreased concentration in hypericins and hyperforin in both species was achieved when 0.4 mg L 1 TDZ was added to the MS culture medium [53]. Same authors reported that modified MS medium (containing 10 mM NH4+ and 5 mM NO3 ) combined with PGRs resulted in approximately 2-fold increased production of hypericins compared to controls in H. maculatum, while H. hirsutum shoots on the same medium revealed a 6.16-fold increase in hyperforin production. Cytokinin supplementation in culture medium was an efficient promoter of secondary metabolite production in various Hypericum spp. Cytokinin (BAP, Z, and TDZ) supplements in the culture medium of H. sampsonii and H. perforatum had significantly different effects on the production of main secondary metabolites [31]. The authors reported increased pseudohypericin (2.95-fold) and hypericin (2.62-fold) production in H. perforatum under TDZ (0.45 μM) treatment, whereas no enhancement of hypericins and hyperforin production was elicited by TDZ in H. sampsonii. Interestingly, pseudohypericin and hypericin production decreased in

H. perforatum

H. perforatum

Species/ Cultivars H. perforatum

Culture medium composition Culture system [PGRs/elicitors-conc] Shoot cultures TDZ (0.45 μM) Gelled media MS + 1/2NH4NO3 + 1/2 KNO3 BAP/IBA (0.44/0.049 μM) (A) BAP (0.22/ 0.44 μM) + IBA (0.049 μM)  (Z) (0.23/0.46 μM) (B) MS hormone free (C) Shoot cultures TDZ (2.27 μM) Gelled media MS + 1/2NH4NO3 + 1/2 KNO3 BAP/IBA (0.44/0.049 μM) (A) BAP (0.22/ 0.44 μM) + IBA (0.049 μM)  (Z) (0.23/0.46 μM) (B) MS hormone free (C) Shoot cultures BAP/Z/IBA (0.22/0.23/ Gelled media 0.049 μM) MS + 1/2NH4NO3 + 1/2 KNO3 BAP/IBA (0.44/0.049 μM) (A) MS hormone free (C) Percent/fold enhancement 2.95-fold vs. control 2.62-fold vs. control

Hyperforin (17 mg g 1 DW)

25  2  C 16 h light/8 h dark photoperiod 72.35 μmmol m 2 s 1

Culture conditions photoperiod temperature 25  2  C 16 h light/8 h dark photoperiod 72.35 μmmol m 2 s 1

50-fold vs. H. 25  2  C sampsonii 16 h light/8 h dark 1.66-fold vs. control photoperiod 72.35 μmmol m 2 s 1

3.85-fold vs. control Pseudohypericin (5.4 mg g 1 DW) 2.9-fold vs. control Hypericin (0.290 mg g 1 DW)

Products/active constituentscontent Pseudohypericin (4.2 mg g 1 DW) Hypericin (0.260 mg g 1 DW)

Secondary Metabolites in Shoot Cultures of Hypericum (continued)

3 subcultures [31] on A for 25 days 20 days on C

6 days after [31] A, B, C 3 subcultures on A for 25 days 1 subculture on B 20 days on C

Subculturing interval References 6 days after [31] A, B, C 3 subcultures on A for 25 days 1 subculture on B 20 days on C

Table 1 Effect of different PGRs/elicitors on the production of various secondary metabolites in shoot cultures of Hypericum spp.

10 279

H. sampsonii

H. perforatum

H. sampsonii

Species/ Cultivars H. perforatum

Culture medium composition Culture system [PGRs/elicitors-conc] Shoot cultures BAP/IBA (0.44/ Gelled media 0.049 μM) MS + 1/2NH4NO3 + 1/2 KNO3 BAP/IBA (0.44/ 0.049 μM) (A) MS hormone free (C) Shoot cultures Z/IBA (0.46/0.049 μM) Gelled media MS + 1/2NH4NO3 + 1/2 KNO3 BAP/IBA (0.44/ 0.049 μM) (A) MS hormone free (C) Shoot cultures MJA (50 μM) Gelled media MS + 1/2NH4NO3 + 1/2 KNO3 BAP/IBA (0.44/ 0.049 μM) (A) BAP (0.22/ 0.44 μM)  (Z) (0.23/0.46 μM) (B) MS hormone free (C) Shoot cultures MJA (50 μM) Gelled media MS + 1/2NH4NO3 + 1/2 KNO3 BAP/IBA (0.44/ 0.049 μM) (A) BAP (0.22/ 0.44 μM) + IBA

Table 1 (continued)

1

Percent/fold enhancement 1.74-fold vs. H. sampsonii 3.05-fold vs. control 65-fold vs. H. sampsonii 1.08-fold vs. control

Culture conditions photoperiod temperature 25  2  C 16 h light/8 h dark photoperiod 72.35 μmmol m 2 s 1

Pseudohypericin Hypericin Hyperforin

1.35-fold vs. control 25  2  C 1.40-fold vs. control 16 h light/8 h dark 1.69-fold vs. control photoperiod 72.35 μmmol m 2 s 1

1.76-fold vs. control 25  2  C 16 h light/8 h dark 2-fold vs. control 1.65-fold vs. control photoperiod 72.35 μmmol m 2 s 1

3.50-fold vs. H. 25  2  C DW) perforatum 16 h light/8 h dark 1.85-fold vs. control photoperiod 72.35 μmmol m 2 s 1

Pseudohypericin Hypericin Hyperforin

Hypericin (0.203 mg g

Products/active constituentscontent Pseudohypericin (1.22 mg g 1 DW) Hyperforin (13 mg g 1 DW)

25 days after [31] A, B, C 3 subcultures on A for 25 days 1 subculture on B 20 days on C 25 days after [31] A, B, C 3 subcultures on A for 25 days 1 subculture

3 subcultures [31] on A for 25 days 20 days on C

Subculturing interval References 3 subcultures [31] on A for 25 days 20 days on C

280 A. Coste et al.

Shoot cultures Gelled media

Shoot cultures Gelled media

Shoot cultures

H. perforatum

H. sampsonii

H. hirsutum

(0.049 μM)  (Z) (0.23/0.46 μM) (B) MS hormone free (C) DHPJA (50 μM) MS + 1/2NH4NO3 + 1/ 2 KNO3 BAP/IBA (0.44/ 0.049 μM) (A) BAP (0.22/ 0.44 μM) + IBA (0.049 μM)  (Z) (0.23/0.46 μM) (B) MS hormone free (C) DHPJA (50 μM) MS + 1/2NH4NO3 + 1/ 2 KNO3 BAP/IBA (0.44/ 0.049 μM) (A) BAP (0.22/ 0.44 μM) + IBA (0.049 μM)  (Z) (0.23/0.46 μM) (B) MS hormone free (C) SA (50 μM) MS 30 g L 1 sucrose Hypericin Pseudohypericin

Pseudohypericin Hypericin Hyperforin

Pseudohypericin Hypericin Hyperforin

7.98-fold vs. control 25  2  C 13.58-fold vs. 16 h light/8 h dark control photoperiod 36 lmol s 1 m 2 light intensity

1.33-fold vs. control 25  2  C 1.75-fold vs. control 16 h light/8 h dark 1.79-fold vs. control photoperiod 72.35 μmmol m 2 s 1

1.05-fold vs. control 25  2  C 1.15-fold vs. control 16 h light/8 h dark 1.49-fold vs. control photoperiod 72.35 μmmol m 2 s 1

5 weeks

(continued)

[53]

25 days after [31] A, B, C 3 subcultures on A for 25 days 1 subculture on B for 12 weeks 20 days on C

25 days after [31] A, B, C 3 subcultures on A for 25 days 1 subculture on B 20 days on C

on B 20 days on C

10 Secondary Metabolites in Shoot Cultures of Hypericum 281

Shoot cultures Gelled media

Shoot cultures Gelled media

Shoot cultures Gelled media

Shoot cultures Gelled media

H. calycinum

H. rumeliacum

H. calycinum

No growth regulators MS vitamins supplementation 30 g L 1 sucrose No growth regulators MS vitamins supplementation 30 g L 1 sucrose

No growth regulators MS vitamins supplementation 30 g L 1 sucrose

BAP/NAA (4.5/ 0.05 μM) MS 30 g L 1 sucrose

Culture medium composition Culture system [PGRs/elicitors-conc] Shoot cultures SA (200 μM) MS 30 g L 1 sucrose

H. perforatum

Species/ Cultivars H. maculatum

Table 1 (continued)

1

FW)

Phenolic compounds (450 mg g 1 DW) Flavonoids (58 mg g 1 DW) Hypericins

Phenolic compounds (450 mg g 1 DW) Flavonoids (58 mg g 1 DW) Hypericins (0.045 mg g 1 DW)

Hypericin (0.45 μg g

Products/active constituentscontent Hypericin Pseudohypericin

1.18-fold vs. B5 1.16-fold vs. B5

4.5-fold vs. H. tetrapterum 1.12-fold vs. B5

2-fold vs. H. tetrapterum 1.93-fold vs. H. tetrapterum

Similar to fieldgrown plants 9-fold vs. callus

Percent/fold enhancement 2.2-fold vs. control 3.94-fold vs. control

25  2  C 16 h light/8 h dark photoperiod 60 μmol m 2 s 1 25  2  C 16 h light/8 h dark photoperiod 60 μmol m 2 s 1

Culture conditions photoperiod temperature 25  2  C 16 h light/8 h dark photoperiod 36 lmol s 1 m 2 light intensity 25  2  C 16 h light/8 h dark photoperiod 30 μmol m 2 s 1 25  2  C 16 h light/8 h dark photoperiod 60 μmol m 2 s 1

45 days

45 days

45 days

60 days

Subculturing interval 5 weeks

[64]

[64]

[64]

[63]

References [53]

282 A. Coste et al.

H. erectum

H. scabroides

H. perforatum

H. perforatum

H. perforatum

H. perforatum

H. perforatum

Hypericin (0.119 μg g 1)

Hypericin (0.119 μg g 1)

IBA/Kin (3.0/ 1.0 mg L 1) MS 30 g L 1 sucrose

IBA/Kin (3.0/ 1.0 mg L 1) MS 30 g L 1 sucrose

22  C 16 h light/8 h dark photoperiod

22  C 16 h light/8 h dark photoperiod

1.5-fold vs. control 2.5-fold vs. control

5–20 days

5–20 days

5–20 days

12 days + 5 days after amino acid feeding

12 days + 5 days after amino acid feeding

25  2  C 5 weeks 16 h light/8 h dark photoperiod 36 μmmol m 2s 1 4 weeks Constant light 25  C in the dark

39.66-fold vs. gelled 25  2  C media under dark 16 h light/8 h dark conditions at 25  C photoperiod

14.87-fold vs. liquid 25  2  C media under light 16 h light/8 h dark conditions at 25  C photoperiod

13.22-fold vs. gelled 25  2  C media under light 16 h light/8 h dark conditions at 15  C photoperiod

Polyphenols 1.87-fold vs. shoot (15.2 mg g 1 DW) primordia 1.70-fold vs. etiolated shoots 22.12-fold vs. callus

Total hypericins BAP (2.0 mg L 1) MS (30 g L 1 sucrose) (0.003–0.005% DW)

Hypericin (0.119 μg g 1)

Adhyperforin 3.7-fold vs. control (0.67 mg g 1 DW)

2-fold vs. control Adhyperforin (0.68 mg g 1 DW)

IBA/Kin (3.0/ 1.0 mg L 1) MS 30 g L 1 sucrose

L-isoleucine (2 mM) BAP/ascorbic acid (0.1/0.1 mg L 1) MS 30 g L 1 sucrose

L-threonine (3 mM) BAP/ascorbic acid (0.1/0.1 mg L 1) MS 30 g L 1 sucrose

Green multiple IAA/BAP (10 5/ 10 5 M) shoots Gelled media LS

Agitated shoot cultures 130 rpm Liquid medium Agitated shoot cultures 130 rpm Liquid medium Shoot cultures Gelled media Light 25  C Shoot cultures Gelled media Light 25  C Shoot cultures Gelled media Light 25  C Shoot cultures Gelled media

Secondary Metabolites in Shoot Cultures of Hypericum (continued)

[68]

[67]

[66]

[66]

[66]

[65]

[65]

10 283

Shoot culture lines (HP1/ HP3) Gelled media Shoot cultures Gelled media

Shoot cultures Gelled media

Shoot cultures Gelled media Leaves

H. perforatum

H. perforatum var. angustifolium

H. polyanthemum

H. perforatum var. angustifolium

Shoot cultures Gelled media

TDZ/IBA (3/2 μM) 2,4D/Kin/NAA (1.3/0.2/0.25 mg L 1) MS 30 g L 1 sucrose TDZ/IBA (3/2 μM) 2,4D/Kin/NAA (1.3/0.2/0.25 mg L 1) MS 30 g L 1 sucrose BAP (1.78 μM) MS 30 g L 1 sucrose

–

BAP/NAA (4.4/ 0.05 mM) MS 30 g L 1 sucrose

Culture medium composition Culture system [PGRs/elicitors-conc] Shoot cultures BAP (4.4 mM) Gelled media MS 30 g L 1 sucrose

H. perforatum

Species/ Cultivars H. perforatum

Table 1 (continued)

14.6-fold vs. green callus

1.79-fold vs. green callus

4 weeks

[72]

4 subcultures [71] every 20–25 days

26  1  C 16 h light/8 h dark photoperiod

[70]

4 subcultures [71] every 20–25 days

20-day intervals/ 10 years

Subculturing interval References 30 days short [69] experiment No subcultures 30 days short [69] experiment No subcultures

26  1  C 16 h light/8 h dark photoperiod

HP3 14-fold vs. HP1 – HP3 6-fold vs. HP1

Culture conditions photoperiod temperature 26  1  C 16 h light/8 h dark photoperiod 15 mE m 2 s 1 1.33-fold vs. control 26  1  C 1.56-fold vs. control 16 h light/8 h dark photoperiod 15 mE m 2 s 1)

Percent/fold enhancement 2.5-fold vs. control

Benzopyrans 12.71-fold vs. stems 25  1  C (0.89 mg g 1 DW) 16 h light/8 h dark photoperiod 45 μmol m 2 s 1

Cyanidin 3-Orhamnoside (4.38 μg g 1)

Phenolic compounds (35.3 mg g 1 FW) Flavonoids (4.52 mg g 1 FW) Pseudohypericin (6.35 mg g 1DW) Hypericin (0.495 mg g 1DW) Cyanidin 3-Oglucoside (7.11 μg g 1)

Products/active constituentscontent Hypericin

284 A. Coste et al.

H. perforatum

H. perforatum

H. perforatum

H. perforatum

H. perforatum

H. myorense

H. synaicum

TDZ/NAA (0.5/0.05 mg L 1)

Hypercin (84.70 μg g

1

DW)

2.56-fold vs. control 26  1  C 16 h light/8 h dark photoperiod Flavonoids Higher amount vs. 22  2  C Shoot cultures BAP (1 mg L 1) wild plants 16 h light/8 h dark Gelled media MS 30 g L 1 sucrose photoperiod 60 μmol m 2 s 1 1 Hypericin 150% vs. control 22  4  C Shoot cultures Sucrose (10 g L ) 16 h light/8 h dark Gelled media MJA (1 mg L 1) photoperiod 8000 lux Hyperforin 280% vs. control 22  4  C Shoot cultures MJA (1 mg L 1) 1 16 h light/8 h dark Gelled media MS 30 g L sucrose photoperiod 8000 lux 4-fold vs. control 24  C Agitated shoot Mannan (0.1 mg ml 1) Pseudohypericin (0.82 mg g 1 DW) 2-fold vs. control MS 30 g L 1 sucrose Continuous light (3000 cultures Hypericin lux) Liquid medium 100 rpm Agitated shoot Mannan (0.5 mg ml 1) Pseudohypericin 3-fold vs. control 24  C cultures MS 30 g L 1 sucrose Continuous light (3000 Liquid lux) medium 100 rpm Agitated shoot b-13-Glucan Pseudohypericin 2.5-fold vs. control 24  C cultures Continuous light (3000 (0.1 mg ml 1) Liquid lux) medium 100 rpm MS 30 g L 1 sucrose

Shoot culture Gelled media

[76]

15 days

15 days

Secondary Metabolites in Shoot Cultures of Hypericum (continued)

[76]

[76]

[75]

21 days

15 days

[75]

[74]

[73]

21 days

4 weeks

1.5 months

10 285

Shoot cultures Gelled media

Shoot cultures Gelled media

BB-bioreactor Liquid medium

H. hookerianum red shoots

H. perforatum

Pseudohypericin

Phenols Flavonoids Anthocyanins Hypericin

Hyperforin

K (2.325 μM) MS + Gelzan (1.5 g L 1)

Sucrose (45 g L 1) MS/B5 medium

Hypericin Pseudohypericin

Products/active constituentscontent Pseudohypericin

DEX [100 mg L 1] MS 30 g L 1 sucrose MS/B5 medium

Culture medium composition Culture system [PGRs/elicitors-conc] Agitated shoot PEC (0.1 mg ml 1) MS 30 g L 1 sucrose cultures Liquid medium 100 rpm Shoot cultures PEC [100 mg L 1] Gelled media MS 30 g L 1 sucrose MS/B5 medium

H. perforatum

H. perforatum

Species/ Cultivars H. perforatum

Table 1 (continued)

2-fold vs. green shoots 3-fold vs. green shoots 5-fold vs. green shoots 44-fold vs. green shoots 2-fold vs. control

1.7-fold vs. control

1.2-fold vs. control 1.5-fold vs. control

Percent/fold enhancement 1.8-fold vs. control

24  C 16 h light/8 h dark photoperiod 35 μmmol m 2 s-2

25 days

25  1  C 3 weeks 16 h light/8 h dark photoperiod 50 μmol
m2
s 1 25  1  C 3 weeks 16 h light/8 h dark photoperiod 50 μmol
m2
s 1 25  2  C to multiply 4 weeks etiolated shoots and in light (50–60 μmol m 2 S 1) 25  2  C under 12 h photoperiod red shoots

Culture conditions photoperiod Subculturing temperature interval 24  C 15 days Continuous light (3000 lux)

[79]

[78]

[77]

[77]

References [76]

286 A. Coste et al.

Transgenic shoot cultures A. rhizogenes

Transgenic plants A. rhizogenes

Transgenic plants T5 clone A. tumefaciens Agitated shoot cultures 140 rpm Liquid medium Agitated shoot cultures 140 rpm Liquid medium Agitated shoot cultures 140 rpm

H. perforatum

H. tomentosum

H. perforatum

H. perforatum cv. “Helos”

H. perforatum cv. “Elixir”

H. perforatum cv. “Helos”

Shoot cultures Gelled media

H. adenotrichum 1

sucrose

BAP/NAA (0.1/ 0.1 mg L 1) LS medium

BAP/NAA (0.1/ 0.1 mg L 1) MS medium

BAP/NAA (0.1/ 0.1 mg L 1) LS medium

MS

MS with B5 vitamins MS 30 g L 1 sucrose

MS 20 g L

PEG (10 g L 1) MS/Galzy

1

Hypericin (165.79 mg 100 g 1 DW)

Flavonoids (328.53 mg 100 g 1 DW)

Flavonoids (255.70 mg 100 g 1 DW)

Hypericin (276.8 μg g

Hypericin (5,2 μg g 1)

DW)

Naphtodianthrones Hyperforin

Hypericin Pseudohypericin

17.80-fold vs. “Elixir” on BAP/NAA (2.0/ 2.0 mg L 1) LS

4.39-fold vs. “Topas” on BAP/NAA (2.0/ 2.0 mg L 1) MS

3.38-fold vs. “Topas” on BAP/NAA (1.0/ 1.0 mg L 1) LS

5-fold increase

12-fold vs. nontransgenic plants 2-fold vs. nontransgenic plants 1.85-fold vs. nontransgenic plants

2.1-fold vs. control 2.3-fold vs. control

25  2  C Constant light 16 μmol m 2 s

25  2  C Constant light 16 μmol m 2 s

25  2  C Constant light 16 μmol m 2 s

1

1

1

22  1  C 16 h light/8 h dark photoperiod. 25  1  C under a of 16 h light/8 h dark photoperiod 50 μmol m2 s 1 23  1  C 16 h light/8 h dark photoperiod 85 μmol m 2 s 1 – [82]

[83]

–

–

6 weeks

6 weeks

(continued)

[84]

[84]

[84]

[81]

–

6 weeks

[80]

15 days

10 Secondary Metabolites in Shoot Cultures of Hypericum 287

H. tomentosum

H. perforatum

H. perforatum

H. perforatum

Species/ Cultivars H. perforatum

Culture medium composition Culture system [PGRs/elicitors-conc] Shoot cultures BAP (1 mg L 1) in non-aerated MS 30 g L 1 sucrose liquid medium (PB) Shoot cultures BAP (1 mg L 1) in non-aerated MS 30 g L 1 sucrose liquid medium (PI) Shoot cultures BAP (1 mg L 1) in non-aerated MS 30 g L 1 sucrose liquid medium (PB) Shoot cultures BAP (1 mg L 1) in non-aerated MS 30 g L 1 sucrose liquid medium (PI) Shoot cultures TDZ (5 μmol) MS/B5 MS 30 g L 1 sucrose Chemical mutagenesis

Table 1 (continued) Percent/fold enhancement 1.89-fold vs. both SS and TI

1.47-fold vs. both SS and TI

9-fold vs. SS 5-fold vs. TI

5-fold vs. SS 2.2-fold vs. TI

12-fold vs. wild plants

Products/active constituentscontent Hypericin (1.8 μg g 1 FW)

Hypericin (1.4 μg g 1 FW)

Phenolic compounds (9 mg g 1 FW) Phenolic compounds (4 mg g 1 FW) Melatonin (23.1 μg g 1)

24  C

16 h light/8 h dark photoperiod

Culture conditions photoperiod temperature 25  2  C 16 h light/8 h dark photoperiod 32.6 μmol m 2 s 1 25  2  C 16 h light/8 h dark photoperiod 32.6 μmol m 2 s 1 25  2  C

14 days

40 days

40 days

80 days

Subculturing interval 80 days

[86]

[85]

[85]

[85]

References [85]

288 A. Coste et al.

10

Secondary Metabolites in Shoot Cultures of Hypericum

289

H. perforatum cultured on medium with Z (0.23 μM or 0.46 μM), while opposite effects were observed in H. sampsonii cultured on the same medium [31]. Low BAP concentrations (0.1–1.0 mg L 1) increased hypericin and pseudohypericin production by approximately 1.25- to 1.1-fold, while higher BAP concentrations (1.0–2.0 mg L 1) led to inhibition of both shoot growth and secondary metabolite production [56]. It was reported that shoot cultures of H. perforatum are outstanding sources for hypericins and hyperforin, although concentration of hypericins in shoot cultures reached a sixfold (1.4% of DW) increase, while the concentrations of hyperforin were lower compared to wild or greenhouse plants [19]. On the other hand, in H. scabroides shoots cultured, only traces of total hypericins were found regardless of BAP concentration [67]. Various secondary metabolites in different Hypericum spp. were reported to be produced in shoot cultures, cell cultures, callus, or adventitious roots [91] although it is important to point that the accumulation of some molecules derived from secondary metabolism particularly hypericin requires differentiated tissues [46]. Research studies revealed a number of flavonoid aglycones and glycosides, especially in shoot culture systems or plant biomass with a high degree of organogenesis [92–94]. Čellárová et al. [95] reported that hypericin and its derivatives are accumulated in special morphological structures which are not found in callus and cell cultures. Similarly the content in polyphenols in differentiated tissues of H. erectum was higher than in callus [68]. In case of H. perforatum, the level of hypericin in callus (on 2 mg L 1, 2,4-D, and 1 mg L 1 Kin) [96] was lower compared to shoots or in vitro plantlets cultured on medium with 4.5 μmol L 1, BAP and 0.05 μmol L 1 NAA [88]. The production of hypericin in H. perforatum was increased in the presence of 4.4 mM BAP, while the production of phenolic compounds was promoted by addition of 4.4 mM BAP and 0.05 mM NAA [69]. Other reports registered a 3.3 to 10 times lower hypericin content under the influence of NAA and BAP (0.5 mg L 1) in shoot cultures of H. perforatum cultivars compared to Hyperici herba (12.2 mg 100 g 1 DW) [97]. Yet, a five- to sixfold higher contents of indole compounds were registered in the biomass from in vitro cultured H. perforatum cultivars (39.6–343.2 mg/100 g DW) compared to field-grown plants (Hyperici herba) (7.1–55.3 mg 100 g 1 DW). Varghese and Sooriamuthu [20] reported that hypocotyls and cotyledons from red H. hookerianum seedlings, cultured on medium with 2.32 μM Kin, produced hypericin (4.38 mg g 1 DW)-rich shoots and that red plant types on MS basal medium showed increased concentrations of total phenols antocyanins and flavonoids and a 19-fold higher concentration of hypericin. The content in secondary metabolites showed variation in two shoot cultures of H. perforatum derived from wild populations [70]. Although hypericin and pseudohypericin were detected in both shoot culture lines, the chemical analysis revealed that one of the lines produced 6 times more hypericin and 14 times more pseudohypericin than the other lines. The presence of hypericins and anthocyanins has been highlighted in shoots of H. perforatum var. angustifolium cultured on medium with TDZ in combination with IBA [71]. The hypericin content in shoots of H. perforatum varied according to explant type and culture medium composition. It was observed that in H. perforatum

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the accumulation of hypericins was superior in organized structures compared to cell suspensions and control [5]. Likewise HPLC analysis revealed that in H. polyanthemum benzopyrans are accumulated especially in the aerial part in various concentrations according to the age of the plant [72]. The total hypericins determined in shoot cultures of 12 Hypericum spp. ranged between 0.49 mg g 1 DW (H. tomentosum) and 7.98 mg g 1 DW (H. rumeliacum) [24], whereas for H. sinaicum shoots regenerated through indirect organogenesis, the highest hypericin content (84 μg g 1 DW) accounting for a 2.56-fold increase compared to control (32.99 μg g 1 DW) was obtained on MS medium supplemented with TDZ (0.5 mg L 1) and NAA (0.05 mg L 1) [73]. It was reported that the presence of BAP and indole-3-acetic acid (IAA) led to accumulation of large amounts of flavonoids in H. mysorense shoots, while the production of hyperforin could not be enhanced by biotransformation [74]. The content in total phenolics and flavonoids was significantly higher in H. calycinum compared to H. rumeliacum and H. tetrapterum [64] on both MS and Gamborg B-5 media [98]. Other studies reported that supplementation of culture medium with low concentrations of BAP (0.2 mg L 1) and IBA (0.1 mg L 1) only slightly affected the level of polyphenolics in hypericin-nonproducing H. calycinum, whereas IBA in higher concentrations inhibited polyphenolic levels in vitro [99]. An increasing interest toward essential oil (EO) production from several Hypericum spp. has arisen in the last few years. However the yield and complexity of these compounds in Hypericum shoot cultures was lower than in wild plants. This might be explained by either different growth conditions or the immaturity of the in vitro shoots compared to those of in vivo plants. Thus, lower EO yield (0.74 mg g 1 DW) was reported in in vitro cultured H. androsaemum shoots than the minimal values obtained from cultivated plants [100]. On the other hand, exposure to MS basal medium and Mg basal medium [101] induced a 2.14- and 2.31-fold increase, respectively, in EO levels in in vitro cultured H. undulatum compared to wildgrowing plants [102]. Plant defense signaling compounds such as jasmonic acid (JA), salicylic acid (SA), and methyl jasmonate (MJA) have been shown to positively influence secondary metabolism in Hypericum spp. Thus, shoot cultures treated with MJA (1 mg L 1) showed increased hypericin (1.5-fold) and hyperforin (2.8-fold) accumulation [75]. Higher MJA concentrations (50 μM/11.2 mg g L 1) resulted in increased pseudohypericin (1.76-/1.35-fold), hypericin (2.0-/1.40-fold), and hyperforin (1.65-/1.69-fold) in H. perforatum and H. sampsonii, respectively. Additionally, higher levels of hypericins were induced by 2,3-dihydroxypropyl jasmonate (DHPJA) (100 μM) supplementation in H. sampsonii than in H. perforatum [31]. Other studies, reported a 1.55- to 3-fold increase of hypericin content in H. maculatum and H. hirsutum, respectively, under the influence of high JA concentration (250 μM) [53]. SA in lower concentrations (50 μM) enhanced the production of hypericin (7.98-fold) and pseudohypericin (13.58-fold) in H. hirsutum while at 200 μM enhanced the production of hypericin (2.2-fold) and pseudohypericin (3.94-fold) in H. maculatum [53].

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Elicitation

Plants counteract with biotic and abiotic stresses via mobilizing their secondary metabolism that is the basis of elicitation [89]. Plants perceive the stress factors by receptors and activate the production of low molecular weight compounds that have an important role in plant defense response and possess important pharmacological properties [103]. Thus an elicitor can be an environmental factor or a signal molecule that activates a signal transduction cascade which mediates the expression of genes related to the biosynthesis of secondary metabolites. Nevertheless the type of culture rather than the type of elicitor defines the compounds induced in Hypericum spp. upon elicitation. Thus, most elicitation methods induce different organogenic processes, some of them being associated with the production of several compounds as well, but not in all situations. Hypericins are the most frequently induced compounds in seedlings and shoot cultures, probably due to the presence of hypericin nodules. Cell suspensions, calli, and root cultures mostly produce flavonoids and xanthones. Several other factors as the type of elicitor, concentration, incubation conditions, and duration of elicitor treatment can also influence the elicitation [103]. Based on their origin, elicitors are mainly classified into two categories: biotic and abiotic. The abiotic elicitors are further divided in chemical and physical triggers [104]. Among biotic elicitors, mainly fungi and bacteria have been used along with yeast extracts and components of microbial cell walls (chitin, chitosan, glucans and pectic poly- and oligosaccharides) in elicitation experiments [76, 77, 105]. Various biotic and abiotic elicitors tested for the manipulation of H. perforatum secondary metabolism are shown in Table 1.

3.2.1 Biotic Elicitors Poly- or oligosaccharides are signaling molecules within elicitation pathways that can induce plant defense response to pathogen invasion [89]. In this context, several studies have been carried out to investigate the effects of polysaccharide elicitors such as mannan, β-1,3-glucan, pectin (PEC), chitin (CHI), and dextran (DEX) on naphtodianthrone production in Hypericum spp. shoot cultures [76, 77, 105]. Thus, in H. perforatum shoot cultures, mannan (0.01–0.1 mg ml 1) stimulated pseudohypericin and hypericin production up to fourfold and twofold, respectively, while β-13-glucan and PEC slightly stimulated pseudohypericin production (about twofold) and had no effect on hypericin production [76]. Yet, other studies [105] revealed lower stimulatory potential of mannan compared to PEC, regarding the biosynthesis of hypericins in in vitro cultured H. adenotrichum seedlings. Thus, mannan (50 mg L 1) stimulated pseudohypericin and hypericin yield up to 2.8-fold and 1.7-fold, respectively, while 4.8- and 2.7-fold increase was registered for pseudohypericin and hypericin production under the influence of PEC (50 mg L 1) [105]. Moreover, Gadzovska Simic et al. [77] evaluated the effects of polysaccharides such as CHI, PEC, and DEX on the production of phenylpropanoids (phenolics and flavonoids) and naphtodianthrones (hypericin and pseudohypericin)

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in H. perforatum shoot cultures. Thus, treatment with CHI and DEX resulted in heterogeneous pattern of total phenolic production in shoot cultures, with levels of about 1.6- to 1.9-fold less than respective controls, while PEC showed the strongest inhibitory effect on total phenolic content in elicited shoots (about 3-fold) compared to control ones at day 21.

3.2.2 Bacteria and Yeast Extracts Meirelles et al. [106] proved that long-term addition of Nomuraea rileyi powder to H. polyanthemum acclimatized plants, following in vitro culture, can result in twofold increase in dry matter yield and total phenolic compound contents, also increasing benzopyrans and uliginosin B concentrations. Usually the biosynthesis of phenolic compounds was stimulated in most of the Hypericum species under elicitation, whereas other compounds were dependent on the species and elicitation treatment. The production of naphthodianthrones and emodin was predominantly stimulated by elicitors derived from Fusarium oxysporum and Trichoderma crassum, while Piriformospora indica promoted the phloroglucinol production. Among flavonoids the aglycone amentoflavone was increased by several elicitors up to 15.7-fold in H. humifusum treated by Potato Dextrose Broth [107]. Elicitation of H. perforatum shoot cultures with inactivated bacteria such as Agrobacterium tumefaciens promoted the hypericin and hyperforin production [75] most probably due to cold-shock protein, flagellin, peptidoglycan, and elongation factor-Tu [108, 109]. Furthermore, hypericin was found at elevated levels in adventitious shoots of H. perforatum after co-cultivation with A. rhizogenes, despite the co-cultivation not resulting in hairy root formation [110, 111]. Treatment of H. perforatum seedlings with Stenotrophomonas maltophilia increased hypericin and pseudohypericin contents [112]. Authors explored the ability of six rhizobacterial strains (N19.27, N11.37, N17.35, N6.8, N21.4, and N5.18), isolated from the rhizosphere of wild populations of Nicotiana glauca, to trigger secondary metabolism in H. perforatum seedlings and shoot cultures. Strain N5.18 significantly increased hypericin up to 1.2 ppm and pseudohypericin up to 3.4 ppm, over controls (0.3 and 2.5 ppm, respectively) when delivered to seedlings. In shoot cultures, only pseudohypericin was detected (168.9 ppm), and significant increases were observed under the effect of different elicitors, reaching even higher values [3164.8 ppm] under moderate elicitor treatment [112]. In H. perforatum shoot cultures, yeast extract did not show any stimulatory effect on either hypericin or pseudohypericin production [76] but increased the accumulation of phenolic compounds and flavonoids in plant cell suspension cultures [113]. 3.2.3 Abiotic Elicitors Physical factors such as exposure to high light intensity, ozone, UV, gamma irradiation, drought, and low temperatures (cryogenic treatment) have been tested for the induction of secondary metabolism in H. perforatum [114]. It is well known that the biosynthesis of hypericin is highly correlated with the cell differentiation

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and the oxidation of its precursor protohypericin is directly induced by light [115]. Yet, Kirakosyan et al. [18] found that neither light/dark treatments nor phytochromemediated (light/dark, red or light/dark, far-red treatments) seemed to play a significant role in regulation of hypericin production in H. perforatum shoot cultures. However, Sood et al. [66] reported higher hypericin content in H. perforatum shoots grown under light conditions (16 h day/8 h night photoperiod) at 25  2  C as compared to 15  2  C. Moreover, significant levels of target compounds (hypericin, hyperforin, flavonoids) were registered in photoactivated dark-grown shoot culture of H. hookerianum after 25 days of light exposure [78]. Thus, the red shoots in particular contained nearly two-, three-, five-, and fourfold higher concentrations of total phenols, flavonoids, anthocyanins, and hypericins than the control green shoot culture [78].

3.2.4 Carbon Source The excess of carbon source, either as sucrose or carbon dioxide, did not increase hypericin content in H. perforatum seedlings grown in bubble bioreactor system. However hyperforin content registered approximately a 2.5-fold increase in plantlets grown at 45 g L 1sucrose under carbon dioxide-non-enriched conditions. These findings provide evidence that the synthesis of the hypericin/pseudohypericin and hyperforin in H. perforatum is dependent on available nutrition and atmospheric conditions [79]. A 14-fold increase in hypericin content was quantified in the shoots grown in the controlled environment system (CCES) with CO2 enrichment [116]. 3.2.5 Cryogenic Treatment Modulation of anthraquinone and phloroglucinol biosynthesis in Hypericum spp. was also demonstrated by cryogenic treatment. The content of hypericins in both pre-cultured H. tetrapterum donor plants and H. perforatum shoots regenerated from cryopreserved meristems increased more than three times. The 38-fold enhancement of phloroglucinols was observed in H. rumeliacum shoots recovered after cryostorage [117]. 3.2.6 Chemical Factors Polyethylene glycol [PEG] at 10 g L 1, after a 15-day treatment, increased hypericin (2.1-fold) and pseudohypericin (2.3-fold) production in in vitro cultured H. adenotrichum seedlings [80]. On the other hand, PEG in similar concentrations (10 and 15 g L 1) had no influence on hypericin and hyperforin synthesis in H. perforatum shoot cultures, while PEG in lower concentrations (2.5 g L 1) in combination with saccharose (20 g L 1) synergistically increased the production of both abovementioned compounds [75]. Moreover, sucrose alone, in higher concentrations (30 g L 1), increased hypericin content from 3 to 4 μg g 1 (1.33-fold) and hyperforin level from 2200 to 3500 μg g 1 (1.59-fold) [75] (Table 1). Chromium treatment induced the production of protopseudohypericin, hypericin, and pseudohypericin in H. perforatum seedlings, whereas nickel treatment did not show any stimulatory effect [118] or demonstrated a 15–20-fold decrease in the concentration of pseudohypericin and hypericin [119]. In H. adenotrichum the

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amounts of hyperoside and isoquercitrin increased by 1.7- and 1.8-fold following 15 days treatment with 0.01 mM chrome, while no significant changes were registered after 30 days of exposure to the same treatment. In both 15- and 30-day 0.1 mM chrome applications, flavonoid amounts decreased significantly. By the application of 0.01 mM of chrome for 15 days, the amount of pseudohypericin increased up to 2.2-fold, and the amount of hypericin also increased up to 1.7-fold [120].

3.2.7 Nanoparticles Submicron-sized particles also known as nanoparticles are currently emerging as new class of elicitors and have been applied in different plant species. Thus, secondary metabolites increases were registered in response to silver nanoparticles in Artemisia annua hairy root cultures [121] and Calendula officinalis plants grown in a hydroponic system [122]. In addition multiwalled carbon nanotubes have been also reported to stimulate the biosynthesis of secondary metabolites in Satureja khuzestanica grown under in vitro conditions [123]. Unfortunately in H. perforatum, only one study has reported that zinc and iron nano-oxides stimulate hypericin and hyperforin production in cell suspension cultures [124]. The adsorption capacities of SiO2 nanoparticles toward quercetin may be further improved by TiO2 functionalization. The adsorption capacity increased linearly with surface coverage of TiO2 [125]. This phenomenon could be used as a new nanotrapping strategy for a wide variety of secondary metabolites.

3.3

Precursor Feeding

A variety of precursors and organic compounds have been used to stimulate the biosynthesis of hypericins especially in H. perforatum shoot cultures [8, 65]. Thus, precursor feeding with L-threonine (3 mM) and L-isoleucine (2 mM) enhanced the production of adhyperforin (2.0-fold and 3.7-fold) in liquid shoot cultures of H. perforatum, while the production of hyperforin could not be enhanced [65]. Improvement in the yield of pseudohypericin [2.4-fold] and hypericin [2.2-fold] was obtained with sodium acetate [50 mg L 1], while emodin [25 mg L 1] enhanced the production of pseudohypericin by 3.2-fold and hypericin by 2.5-fold compared to control. Prominent yields were achieved for pseudohypericin [4.5-fold] and hypericin [3.7-fold], when sodium acetate and emodin were fed simultaneously. Other organic compounds such as succinic acid (5 mM) and malic acid (10 mM) were also proven to stimulate the production of hypericins, although the effect was moderate [126].

3.4

Genetic Transformation

Plant genetic transformation and overexpression of target genes are already in use to promote secondary metabolite production in Hypericum genus [62, 127, 128]. Thus, an enhanced level of naphtodianthrones (12-fold) and hyperforin (2-fold) were reported compared to control in H. perforatum transgenic shoots regenerated from hairy roots after co-cultivation with Agrobacterium rhizogenes strain A4 [81].

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A. rhizogenes-mediated genetic transformation was also reported in other species besides H. perforatum [82, 129]. Thus, Nigutová et al. [82] reported in hairy rootderived transgenic shoots of H. tomentosum a 1.85-fold total hypericins increase versus non-transgenic plants. Among Agrobacterium spp., Agrobacterium tumefaciens-mediated transformation is the most preferred method of gene transfer due to frequent single-copy transgene integration into the plant genome and low incidence of transgene silencing [62, 128, 130]. A. tumefaciens (harboring pCAMBIA1301)-mediated genetic transformation studies resulted in the production of four transgenic H. perforatum clones, namely, T2, T3, T5, and T8, with different growth parameters [83]. The highest of fivefold increase in hypericin content was recorded by T5 (276.8  9.2 μg g 1 DW) in comparison to control non-transformed plants (35.6  2.7 μg g 1 DW) [83]. However, A. tumefaciens-mediated genetic transformation studies in recalcitrant Hypericum spp. have not been very fruitful as the secondary metabolites present in the plant itself were reported to inhibit the growth of the bacteria. Thus plant defense suppressing strategies in recalcitrant species were already investigated. Thus, antioxidant mixtures [131–133]; biolistic bombardment [128] and protoplast transformation have been used to avoid the presence of the cell wall, with crucial role in host defense [134, 135]; virus-mediated transformation [136, 137]; and nanoparticlemediated DNA delivery into plant cells [138] are only several strategies that were already tested and could successfully contribute to the genetic transformation of Hypericum plants in the future. Additionally, Hou et al. [62] proposed the use of different metabolic engineering technologies to control target compound-related genes and enzymes activity [RNAi technology; short palindromic repeat (CRISPR)/CRISPR-associated protein (Cas9) system] for secondary metabolite enhancement in Hypericum spp. Moreover, several Hypericum spp. were screened with next-generation sequencing technology with the aim to generate and enhance gene annotations, especially for genes coding the enzymes supposedly included in biosynthesis of valuable bioactive compound [139]. However, Hypericum-specific pathway engineering is still at its infancy, being currently limited by the lack of cloned genes and coding enzymes. Moreover, controlling the functions of hypericin synthesis-related genes might be possible only in a complex system, which contain the structures in which target compounds [with possible toxic effect on plant tissues] accumulate. In this context, in order to promote secondary metabolite production through genetic transformation, increase in knowledge of functional genomics and specific metabolic pathways and an efficient A. tumefaciens-mediated transformation protocol are mandatory.

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Other Factors Affecting Secondary Metabolite Yield in Shoot Cultures of Hypericum Species

Several research studies underlined the influence of media culture composition in combination with culture system type [liquid, semisolid or solidified], genotype, and plant’s developmental stage for the efficient production of target compounds [84].

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Culture System

Most Hypericum shoot cultures were carried out on solidifying media, although several studies have reported the successful use of liquid media for secondary metabolite production [53, 84, 85]. Though liquid systems proved several advantages over gelled media, several studies reported hyperhydricity and abnormal shoot morphology [53]. To counteract these problems, other procedures have been developed, including culture supports such as paper bridges (PB), liquid medium overlaying, and temporary immersion system or agitated liquid cultures [53, 85, 140]. Thus, enhanced adhyperforin production (3.7-fold) was reported in shoot cultures of H. perforatum grown in shaken liquid culture system (supplemented with 0.1 mg L 1 BA and L-isoleucine) [8]. Other authors reported high concentrations of hypericin in H. perforatum shoots grown in non-aerated, static liquid medium in partial immersion system (PI) [1.4 μg g 1 FW], and on paper bridge support [1.8 μg g 1 FW] as well as increased production of polyphenolic compounds [85]. Moreover, Coste et al. [53] applied a combination of several factors (agitated liquid medium, modified MS medium, PGRs, and JA/SA elicitation) for enhanced production of hypericin (7.98-/2.2-fold) and pseudohypericin (13.58-fold/3.94-fold) in agitated shoot cultures of H. hirsutum and H. maculatum, respectively. Yet, Sood et al. [66] revealed that H. perforatum shoots grown on solid MS media [supplemented with IBA/Kin] produced higher amounts of hypericin [15-fold increase] as compared to shoots grown on liquid media. Nonetheless, high amounts of flavonoids were also reported in H. perforatum agitated liquid shoot cultures [supplemented with 0.1 mg L 1 BAP/NAA] on both Linsmaier and Skoog (18-fold increase) [141] and MS (11-fold increase) [84]. Liquid culture systems were also implemented for essential oil production. Thus, Morshedloo et al. [140] compared the effect of three basal media including MS, B5, and half-strength B5, on volatile constituent production in H. perforatum agitated liquid shoot cultures. They have identified 44 volatile components with varying profile depending on the type of basal media used. Large-scale in vitro production of H. perforatum in bioreactors has also been reported [142]. Thus, in order to eliminate the quality impairment in St. John’s wort products, Zobayed et al. [142] reported six different culture systems to compare the biomass and secondary metabolite production with greenhouse plants. While liquid medium systems in bubble bioreactor effectively produced large quantities of biomass, the chemical profiles of these tissues were less desirable. Of the six culture systems, they have found that growth in large vessel with gelled medium under forced ventilation was optimal for the production of biomass and secondary metabolite production. However, the use of large-scale shoot cultures had limited success because the yields of secondary compounds in the cultures have been too low for commercialization.

4.2

Genotype and Ploidy Level

To improve yield of target compounds in Hypericum spp., effective breeding strategies based on the fundamental knowledge of reproduction biology and genetics

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should also be considered. H. perforatum is naturally distributed as facultative tetraploid apomict (2n = 4x = 32). Many studies have emphasized a high genotype variation with different ploidies/genome sizes for H. perforatum wild or cultivated plants. Thus, Matzk et al. [143] reported diploid, tetraploid, pentaploid, and hexaploid accessions of H. perforatum. As a consequence, many comprehensible investigations exist about its modes of reproduction and ploidy level investigation in order to identify superior germplasm, whether for traditional cultivation or for development of superior plant cell culture lines [44]. High-yielding, easy to harvest, and cost-effective genotypes are of interest for breeders. Thus, for traditional breeding, a few selected varieties, e.g., “Hyperimed,” “Elixir,” and “Topas,” are already commercially available [144]. Most plant tissue culture studies have looked especially at the regeneration of in vitro plants and secondary metabolite production in only one genotype [145, 146]. Several studies pointed out that genetic factors strongly affect the content of secondary metabolites in H. perforatum [147]. The use of different genotypes in combination with PGRs was reported to enhance hypericin production in shoot cultures of H. perforatum. Thus, a maximum level of hypericin was reported in shoot cultures of “Helos,” “Topas,” and “New Stem” genotypes under 1.0 mg L 1 BAP treatment, whereas 1.5 mg L 1 BAP was the suitable concentration for hypericin production in “Ardabil” accession [148]. Kwiecień et al. [84] performed the first comparison of flavonoid production in three cultivars of H. perforatum (“Elixir,” “Helos,” and “Topas”), reporting the highest amounts of flavonoids in shoots of cultivar “Elixir” grown on MS agitated liquid medium. Earlier studies of in vitro cultures of cultivars “Elixir,” “Helos,” and “Topas” focused on the analysis of nonhallucinogenic indole compounds and free phenolic acids in agar and agitated in vitro cultures, respectively [97, 149]. Extracts from shoots of the cultivar “Helos” were reported to contain high levels of serotonin (319.9 and 197.4 mg 100 g 1 DW) [97]. Moreover, analysis of the content of secondary metabolites in individual diploid and tetraploid H. perforatum seedlings revealed no significant differences between plants with different ploidy levels [147]. Interest in the development of H. perforatum lines with distinct bioactive compounds led to the establishment of different H. perforatum germplasm collections [86, 150]. Thus, a germplasm line of St. John’s wort with high levels of melatonin was selected in vitro using mutagenized tissues. The germplasm line has remained stable over a 5-year period and contained >12-fold melatonin content compared with wild plants [86]. Another in vitro collection was developed by Alan et al. [150], consisting of 29 clonal H. perforatum cv. New Stem lines, each derived from a single plantlet and selected on the basis of growth features, antioxidant potential, and bioactive compound composition. Flow cytometric (FCM) screening of in vitro maintained germplasm lines showed similar nuclear DNA amounts and ploidy level (2n = 4x) as donor plants. Thus, FCM analysis proved to be useful to evaluate the genetic uniformity of medicinal plants, an important aspect for long-term preservation of elite clones. Clonal propagation and preservation of elite genotypes, selected by their superior characteristics, require high degree of genetic uniformity among the regenerated plants [151]. The potential for improving plant productivity

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and their proper use in agriculture relies largely on newly developed DNA biotechnology and molecular markers. These techniques enable the selection of successful genotypes, better isolation and cloning of favorable traits, and the creation of transgenic organisms of importance to agriculture and industry [44].

4.3

Developmental Stage

Plant growth and developmental stage was also shown to affect the yield of target compounds in both wild populations and in vitro cultured plants [152]. Biosynthesis of hypericins is connected with the morphogenesis and formation of dark redcolored oil glands on the leaves of both intact plants and shoot organ cultures [6]. Thus, Kwiecień et al. [84] recommended the use of microshoot cultures, motivating that cells at this differentiating stage can possess high growth potential and can produce high levels of biologically active metabolites. Earlier studies reported that in the newly formed leaves of H. perforatum, the content of hypericins is higher than in intact plants [5]. In H. perforatum, large amount of flavonoid accumulation was related to leaf morphogenesis during advanced plant development stage [92]. Moreover, Košuth et al. [146] underlined that intraspecific chemical variation (related to dianthrones and phloroglucinols production) was present under controlled conditions even in the early stages of plant development.

5

Conclusions

Among all novel technologies, elicitation emerges as the most attractive strategy for the enhancement of secondary metabolite production in Hypericum spp. It is obvious that shoot cultures represent the best approach for the production of specific compounds, especially of hypericins, which accumulate mainly in dark glands in the aerial parts of the plants. However, recent studies identified the presence of these compounds in other tissues lacking secretory glands. The contradictory results obtained by different research groups in terms of secondary metabolites synthesis and accumulation by particular elicitation approaches in Hypericum spp. are closely linked with the complex nature of culture conditions, culture type and the particular elicitation potential and response of each species. Though, the elicitation potential is variable and subjected to the influence of diverse parameters, some strategies proved to be efficient in producing similar and even higher amounts of target compounds compared to field-grown plants. Therefore, further research is needed to optimize the best reproducible protocol for each species. Furthermore, it is important to understand first the biosynthetic pathways by an integrated OMICS approach in H. perforatum before extending it to other elite Hypericum spp. for novel value added compound production by metabolic engineering. Acknowledgments This work was partially supported by a grant of the Ministry of Research and Innovation through Program 1 – Development of the National R&D System Subprogram 1.2 – Institutional Performance – Projects for Excellence Financing in RDI ctr. no. 22PFE/2018 and

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partially by the core program PN2019-2022 – BIODIVERS 3 through the project BIOSERV project code 19270201 contract nr. 25 N/2019.

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Different Types of In Vitro Cultures of Schisandra chinensis and Its Cultivar (S. chinensis cv. Sadova): A Rich Potential Source of Specific Lignans and Phenolic Compounds

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Agnieszka Szopa, Adam Kokotkiewicz, Marta Klimek-Szczykutowicz, Maria Luczkiewicz, and Halina Maria Ekiert Contents 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.1 S. chinensis: Species Characteristics and Significance in Modern Phytotherapy and Cosmetology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.2 Schisandra lignans (Dibenzocyclooctadiene Lignans) and Phenolic Compounds . . . . 1.3 An Overview of Former Biotechnological Research on S. chinensis In Vitro Cultures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Production of Dibenzocyclooctadiene Lignans in S. chinensis Microshoot Cultures . . . . . 2.1 Optimization of Culture Type . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2 Optimization of Culture Lighting Conditions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3 Optimization of Elicitation Processes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.4 Optimization of Lignan Production in Bioreactors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.5 Optimization of Lignan Production Based on Culture Type in Microshoot Cultures of S. chinensis cv. Sadova . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Production of Phenolic Compounds in S. chinensis Microshoot Cultures . . . . . . . . . . . . . . . . . 3.1 Optimization of Culture Type . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2 Optimization of Culture Lighting Conditions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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A. Szopa (*) · H. M. Ekiert Department of Pharmaceutical Botany, Faculty of Pharmacy, Jagiellonian University, Medical College, Kraków, Poland e-mail: [email protected]; [email protected]; [email protected] A. Kokotkiewicz · M. Luczkiewicz Department of Pharmacognosy, Faculty of Pharmacy, Medical University of Gdańsk, Gdańsk, Poland e-mail: [email protected]; [email protected] M. Klimek-Szczykutowicz Department of Pharmaceutical Botany, Jagiellonian University, Collegium Medicum, Kraków, Poland e-mail: [email protected] © Springer Nature Switzerland AG 2021 K. G. Ramawat et al. (eds.), Plant Cell and Tissue Differentiation and Secondary Metabolites, Reference Series in Phytochemistry, https://doi.org/10.1007/978-3-030-30185-9_10

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3.3 Optimization of the Production of Phenolic Compounds in Bioreactors . . . . . . . . . . . . 3.4 Optimization of the Production of Phenolic Compounds in Microshoot Cultures of S. chinensis cv. Sadova . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Abstract

Chinese magnolia vine (Schisandra chinensis) is a well-known traditional Chinese medicinal plant species, which is very important in modern phytotherapy. The key role is assigned to specific compounds – dibenzocyclooctadiene lignans. This chapter describes the use of S. chinensis in vitro cultures as a tool to increase their production under controlled conditions as a promising biotechnological alternative to their extraction from ex vitro plant material or by chemical synthesis. Moreover, the chosen phenolic compounds (phenolic acids and flavonoids) were taken into consideration, too. The whole process of biotechnological research was applied to studying S. chinensis and its cultivar – S. chinensis cv. Sadova No. 1. The studies involved initiation of in vitro cultures, optimization of the basal composition of the culture media, duration of the growth cycles, type of cultures, culture lighting conditions, elicitors, and bioreactor design. In all the steps of biotechnological process, efficient production of specific lignans and phenolic compounds was achieved. The research proved that plant in vitro cultures of both Schisandras can be an efficient tool used to increase the production of the desired secondary metabolites. Keywords

Chinese magnolia vine · Microshoot cultures · Schisandra · Schisandra lignans · Cultivar · Phenolic acids · Flavonoids · Elicitation · Plant bioreactors Abbreviations

2,4-D BA DW GA3 LC-DAD LC-DAD-ESI-MS LC-UV LS MS NAA PGRs WV5

2,4-Dichlorophenoxyacetic acid 6-Benzyladenine Dry weight Gibberellic acid Liquid chromatography with diode-array detection Liquid chromatography with diode-array detection and electrospray ionization mass spectrometry Liquid chromatography with ultraviolet-visible detector Linsmaier and Skoog Murashige and Skoog 1-Naphthaleneacetic acid Plant growth regulators Westvaco

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1

Introduction

1.1

S. chinensis: Species Characteristics and Significance in Modern Phytotherapy and Cosmetology

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Schisandra chinensis (Schizandra chinensis, Chinese magnolia vine) (Turczaninow) Baillion – Chinese magnolia vine – is a dioecious climber plant belonging to the family Schisandraceae (Fig. 1a). The raw material is a berry-like red fruit, aggregated in raceme-like multiple fruits. The natural sites of S. chinensis occurrence are located in Northeastern China, in Korea, and in Japan, as well as in the eastern part of Russia, in Primorsko, on the Kuril Islands, and in the southern part of the Sakhalin Island. This species usually occurs on the periphery of mixed forests, often by streams. It prefers sites with moderate humidity and light, sheltered from the wind, and moist soils rich in humus [1–4].

Fig. 1 The morphological appearance of (a) Schisandra chinensis and (b) S. chinensis cv. Sadova No. 1 fruiting plants – specimens of the plants cultured in CLEMATIS “Source of Good Vines” ltd. with headquarters in Pruszków, Poland

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In the regions of natural occurrence, the fruits of Chinese magnolia vine are a valuable medicinal resource. The first description of this species can be found in the most valuable work of ancient Chinese medicine by Li Shih-Chen – “Pên T’shao Kang Mu” from 1596. The fruit of Chinese magnolia vine (Chin. Bei-Wuweizi; 五味 子) is called “the fruit of five flavors” and has been used for thousands of years in traditional Chinese medicine (TCM) [5]. Its individual parts are characterized by the following flavors: sweet, salty, sour, bitter, and “tart” (“astringent”). It was believed that the salty and sour tastes were responsible for improving the functioning of the liver and male gonads; the bitter and tart tastes affected the heart and lungs while the sweet taste the stomach. The Chinese magnolia vine fruit is also known in traditional Russian medicine, where it is described as a tonic, reducing hunger, thirst, and fatigue remedy [6]. S. chinensis is also a crop plant in China, Korea, and Russia due to its valuable properties. Import of raw materials into European countries for the needs of the pharmaceutical and cosmetic industries is carried out from those areas. Cultivation of Chinese magnolia vine in Europe as well as in North America is quite difficult. This species is most often planted as an ornamental plant in parks, as well as in home and monastery gardens [4, 6, 7]. S. chinensis has been introduced into official European medicine, including Poland, 11 years ago. The description of the raw material, which is the fruit of the Chinese magnolia vine – Schisandrae chinensis fructus – appeared for the first time in 2008 in Supplement 6.3 to the European Pharmacopoeia 6th [8]. This monograph was published in 2009 in the equally valuable supplement to the eighth edition of the Polish Pharmacopoeia [9]. The monograph remains unchanged in the latest ninth edition of the European Pharmacopoeia [10] and in the 11th Polish Pharmacopoeia [11]. The raw material has been used for many years in the official healthcare systems of Asian countries, mainly Japan [12], Korea [13], and especially China [14]. It is also known in North America, where it has been listed in the American Herbal Pharmacopoeia since 1999 [15]. Since 2007, it has also had its monograph in the International Pharmacopoeia published by World Health Organization (WHO) [16]. A highly fruit productive cultivar of Chinese magnolia vine, called S. chinensis cv. Sadova No. 1 (Fig. 1b), was selected in Ukraine by Ivan Szajtan (1914–2002) – an employee of the M. M. Gryshko National Botanical Garden in Kiev (Russian: Національний ботанічний сад ім. М.М. Гришка НАН України). This cultivar differs from the native species mainly in fruit traits; the fruits of S. chinensis cv. Sadova No. 1 are larger and matt. In addition, this species is characterized by higher fruit yields. Importantly, unlike S. chinensis, its cultivar is a monoicous, selfpollinating species and more resistant to climate change [1, 17, 18]. To the best of our knowledge, in the scientific literature, there is no information on phytochemical analyses or biotechnological research of S. chinensis cv. Sadova. The interest in the therapeutic values of S. chinensis fruits, mainly in East Asian countries but also in American and European markets, is proven by numerous phytoformulations including dietary supplements and medicinal preparations advertised mainly as adaptogenic, energetic, and hepatoprotective. There are also

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increasingly numerous cosmetic preparations based on Chinese magnolia vine fruits [4, 19]. All these product groups are based on the raw material of natural origin – fruit extracts. The main source of this raw material in Poland is imported from Southeast Asia, primarily from China. Theoretically, the possible chemical synthesis of the main S. chinensis secondary metabolites, called “Schisandra lignans,” is a multistage and very complicated process. Thus far, only one synthetic drug is available, DDB (bifendate, dimethyl diphenyl bicarboxylate), a derivative of schisandrin C (produced in China) applied in medicine as a hepatoprotective agent [20]. The raw material of natural origin – the fruits of Chinese magnolia vine – remains irreplaceable. In vitro cultures may be an alternative source for obtaining “Schisandra lignans” (dibenzocyclooctadiene lignans) [4, 21] (Fig. 2). The healing properties of Schisandra fruit, known for their traditional use, have been confirmed by numerous scientific studies. In addition, modern research results document new directions of their activity. Strong hepatoregenerative, hepatoprotective, and antioxidant effects of fruit extracts as well as extracted compounds have been proven, among others, for gomisin A, deoxyschisandrin, and schisandrin C. The hepatoregenerating effect is associated with the already established position of S. chinensis in the treatment of liver diseases, including infectious and autoimmune diseases as well as inflammation, steatosis, and cirrhosis [22–24]. Recent research indicates antitumor activity in colorectal cancer and stimulation of cancer cell apoptosis in leukemia and cancers of the skin, large intestine, ovaries, and liver [25–28]. Immunostimulatory and immunomodulating effects of polysaccharides isolated from Schisandra fruits have also been shown [29]. Fruit extracts have a positive effect on the nervous system; they protect neuronal cell against death and also increase the concentration of neurotransmitters in the central nervous system; thus they can be used as excipients in Alzheimer’s or Parkinson’s disease. Moreover, they improve concentration, attention span, learning, and memorizing abilities; they also exhibit antidepressant activity without causing adverse drowsiness; they act adaptogenously, supporting the body’s resistance to stress and traumatic experiences; they are used in states of exhaustion and anxiety; their inhibitory effect on HIV multiplication has been proven [30]. Fruit extracts also have a positive effect on the cardiovascular system, preventing heart attacks and regulating elevated blood pressure; they help to maintain normal body weight by preventing obesity [7, 31]; they exhibit an anti-asthmatic effect by reducing airway hyperresponsiveness; they act anti-osteoporotically, stimulating osteoblast proliferation, and anti-ulceratively, healing ulcer wounds in the stomach and duodenum [7]; in addition, Chinese magnolia vine fruits have strong antioxidant and anti-inflammatory properties [4, 19, 21, 32–34]. Biological properties of Chinese magnolia vine fruits have gained the interest of cosmetic companies, which have introduced fruit extracts for the production of various cosmetics. Antioxidant, UV-protection, so-called antiaging, skin whitening, soothing, and antiallergic effects of fruit extracts are valuable properties from a cosmetic point of view [19, 35]. Recently, more potent antioxidant, anti-inflammatory, antibacterial, and antifungal properties of Chinese magnolia vine leaf and shoot extracts, compared to fruits, have been proven [34].

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Deoxyschisandrin

Gomisin A

Schisandrin

dimethylgomisin J schisandrin A wuweizisu A

schisandrol B wuweizi alcohol B wuweizichun B

schisandrol A wuweizichun A

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gomisin B, schizandrer B wuweizi ester B

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H3C H

Fig. 2 Chosen dibenzocyclooctadiene lignans – their chemical structures and synonymous names

Plant in vitro cultures, and the ability to control and increase the endogenous production of secondary metabolites offered by various techniques used in plant biotechnology, create the possibility of their application in the pharmaceutical,

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cosmetic, and food industries. The valued strategies used to increase the endogenous production of active compounds include such treatments as optimization of culture conditions (selection of the basic composition of the culture medium, composition of plant growth regulators (PGRs)), type of culture (agar, liquid stationary, agitated, bioreactor cultures), and light conditions (light wavelength, lack of light), which have been applied in studying of in vitro cultures of S. chinensis performed by our team [36–43].

1.2

Schisandra lignans (Dibenzocyclooctadiene Lignans) and Phenolic Compounds

The valuable biological properties and the therapeutic applications resulting from them are determined by the unique chemical composition of S. chinensis [4, 7, 31]. Dibenzocyclooctadiene lignans are the main group of secondary metabolites specific to this species [31, 44, 45]. Due to the characteristic, very complex chemical structure of these compounds, as well as their occurrence only in Schisandra species, even in professional studies, they are referred to as “Chinese magnolia vine lignans” or “Schisandra lignans.” The content of lignans in fruits is influenced by the degree of fruit maturity, harvest time, area of plants, occurrence, and their habitat. This content varies within a wide range, from 4% to 19% DW (dry weight). Data from 2007 in the WHO monograph, mentioned the presence of 30 compounds from this group in S. chinensis; however, the number of newly identified structures is constantly increasing [16, 46]. In addition, these compounds have numerous synonymous names, which hinder the free exchange of scientific knowledge. The first discovered lignans, and at the same time occurring in the largest amounts in the fruits, were: schisandrin, gomisin A, deoxyschisandrin, schisandrin B, schisandrin C, γ-schisandrin, schisanhenol, schisantherin A, and schisantherin B (Fig. 2) [4, 21, 31, 47]. Chinese magnolia vine fruits, according to the requirements of the European Pharmacopoeia, must be standardized for the content of schisandrin, whose concentration in the dried material cannot be less than 0.4% [10]. The main pharmacological activities of S. chinensis are determined mainly by the presence of schisandra lignans. The fact of the presence of dibenzocyclooctadiene lignans, whose biogenesis is not yet fully understood, but its initial stages are known – the shikimic acid pathway, which is also characteristic of the biogenesis of polyphenolic compounds, such as phenolic acids and flavonoids, decided about interest on these polyphenols accumulation by our team. The aforementioned polyphenolic groups of compounds also have numerous, valuable pharmacological and cosmetic properties. Phenolic acids, both numerous derivatives of cinnamic and benzoic acids and depsides, are a pharmacologically attractive group of plant metabolites due to many valuable therapeutic properties, including anticancer, choleretic, diastolic, hypolipemic, antithrombotic, immunostimulant, as well as strong anti-inflammatory and antioxidant properties

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[48–54]. Flavonoids (both aglycons and flavonoid glycosides) are also an extremely valuable group of plant secondary metabolites from the point of therapeutic effects. Noteworthy are their strong antioxidant, anti-inflammatory, anticancer, diuretic, and antiulcer activities and the ability to seal the walls of capillary vessels [52, 54–59]. So far, phytochemical studies performed on S. chinensis plant-derived extracts have documented the presence of some phenolic acids and flavonoids in fruit extracts [34, 60–62]. Former analyses of S. chinensis fruit extracts by our team have confirmed the presence of chlorogenic, p-coumaric, p-hydroxybenzoic, protocatechuic, salicylic, and syringic acids [60]. Other authors have additionally proven the presence of gentisic acid and flavonoids: hyperoside, isoquercitrin, rutoside, and quercetin [34].

1.3

An Overview of Former Biotechnological Research on S. chinensis In Vitro Cultures

The in vitro cultures of S. chinensis are a very attractive object of biotechnological research. Thus far, research in this field has been carried out only in single research centers of East Asian countries: China, Japan, and South Korea. In Europe, this subject was recently undertaken only by two Czech scientific institutes from Brno. Most of the studies carried out to date have focused on the development of S. chinensis micropropagation methods mainly through somatic embryogenesis [63–66]. In addition to this research, several works have been published in recent years focusing on endogenous accumulation of the most important group of biologically active S. chinensis secondary metabolites – dibenzocyclooctadiene lignans. The possibilities of biotechnological production of selected Chinese magnolia vine lignans have been documented, apart from our own works, by groups from the Czech Republic and Japan [67–69]. In the studies described above, the obtained efficiencies of dibenzocyclooctadiene lignans production were very low compared to the results acquired under the research program on optimizing the production of Schisandra lignans in S. chinensis cultures performed by our team. S. chinensis in vitro cultures were successfully established in our laboratory in 2011 as cultures with different degrees of differentiation – shoot-differentiating callus cultures and undifferentiated callus cultures, from leaf buds of female S. chinensis plants from Rogów Arboretum, Forest Experimental Station, Warsaw University of Life Sciences, Rogów, Poland. The culture conditions were optimized by testing different variants of Murashige and Skoog (MS) culture media [70] differing in the concentration of selected PGRs: BA (6-benzyladenine; cytokinin) and NAA (1-naphthaleneacetic acid; auxin) in the concentration range from 0.1 to 3 mg/l. The studies were performed on different types of in vitro cultures – agar cultures (shoot-differentiating callus and undifferentiated callus cultures) and agitated cultures (shoot and suspension cultures). In the studied cell extracts, qualitative

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and quantitative analyses (LC-UV) of 4 main schisandra lignans and 11 phenolic acids, were performed [60, 71–74]. The results of that preliminary research proved that in vitro cultures of S. chinensis could be a rich, potential, independent of the seasons, climatic conditions, etc. source of these medically valuable metabolites. The further research was focused on cultures with a higher degree of organogenesis – microshoot cultures. The “optimal” culture medium (MS-based) variant for the growth and production of metabolites, containing 3 mg/l BA and 1 mg/l NAA, was chosen as a result of composition optimization based on the concentrations of PGRs. As part of the research, studies were performed on optimizing the other culture conditions – selection of the culture system, duration of the culture cycle, light conditions, and different types of elicitors. In addition, the optimization of S. chinensis microshoot cultures in different types of bioreactors was achieved. The quantities of lignan compounds in the biomass cultured in vitro were significant, similar to the content in the plant material growing in vivo. The experiments on microshoot cultures of S. chinensis cultivar Sadova No. 1 were an important, innovative aspect of the studies. As part of this research, we carried out an extensive, full optimization of the culture process, with the use of elicitation and large-scale laboratory cultures in bioreactors. Detailed aims and results of the studies are presented below.

2

Production of Dibenzocyclooctadiene Lignans in S. chinensis Microshoot Cultures

2.1

Optimization of Culture Type

The aim of the work was to optimize the conditions for S. chinensis microshoot cultures related to the type of the culture: agar and liquid cultures, stationary and agitated, in order to obtain high production of dibenzocyclooctadiene lignans and satisfactory biomass increments [75] (Fig. 3). In vitro cultures were carried out on the “production” MS medium containing 3 mg/l BA and 1 mg/l NAA, selected based on previous studies [72–74]. The following continuous culture growth cycles were tested: 30- and 60-day cycles; in addition, the so-called semicontinuous cultures (biomass fed with a portion of fresh culture medium after 30 days, with 60-day culture cycles) were also tested. The amounts of dibenzocyclooctadiene lignans were determined in methanol extracts from lyophilized biomass of in vitro cultures, in culture media, and in plant material (fruits and leaves of parent plant) for comparative purposes, using modern LC-DAD [76] and LC-DAD-ESI-MS techniques. In the in vitro microshoot extracts, the qualitive and quantitive analyzes of nine lignans: gomisin A and G, deoxyschisandrin, schisandrin, schisandrin C, schisantherin A, schisantherin B, schisanhenol, and γ-schisandrin, were performed. Moreover, five additional dibenzocyclooctadiene lignans were confirmed using LCDAD-ESI-MS method: schisandrin B, benzoylgomisin P, angeloyl /tigloylgomisin

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Fig. 3 Microshoot cultures of S. chinensis (MS medium containing 3 mg/l BA and 1 mg/l NAA, day 30 of culture): (a) agar cultures, (b) stationary liquid cultures, (c) agitated cultures [75]

H, angeloyl /tigloylgomisin Q, and schisantherin D (or its stereoisomer: benzoylgomisin O or benzoylysogomisin O). The amounts of the tentatively identified compounds were calculated according to the schisandrin standard curve (the main lignan of this group, to which the raw material is standardized according to pharmacopoeial requirements) [10] (Table 1). In total, 14 lignans were found in all the extracts analyzed in this study. The most quantitatively dominant were schisandrin (max. 65.62 mg/100 g DW), angeloyl-/ tigloylgomisin Q (max. 49.73 mg/100 g DW), deoxyschisandrin (max. 43.65 mg/ 100 g DW), and gomisin A (max. 34.36 mg/100 g DW). The highest lignan contents were found in biomass extracts from stationary cultures (maintained in Magenta™ vessels, Sigma) after 30-day culture periods: for agar cultures, 237.86 mg/100 g DW, and for liquid stationary cultures, 274.65 mg/100 g DW. The highest total lignan content (244.80 mg/100 g DW) in extracts from agitated cultures was found in biomass cultured in a semicontinuous system (30/60 days) (Table 1). No lignans were found in culture media. The analysis of dibenzocyclooctadiene lignans was performed for the first time in this study using the LC-DAD-ESI-MS technique in extracts from S. chinensis cultures grown in various in vitro systems. The results were compared with the amounts of lignans in the parent plant extracts, analyzed for comparison. It was found that the in vitro cultures were a rich source of Schisandra lignans, in particular

Lignans Schisandrin Gomisin A Gomisin G Schisantherin A Schisantherin B Schisanhenol Deoxyschisandrin γ-Schisandrin Schisandrin C Total content

In vitro cultures Agar 52.96  1.52 25.90  1.86 1.44  0.06 0.42  0.03 0.80  0.06 0.51  0.05 30.27  0.89 8.04  0.44 2.35  0.12 237.86 6 11.58 Stationary liquid 65.62  2.68 34.36  1.24 1.64  0.46 0.85  0.06 0.93  0.16 0.52  2.47 43.65  1.17 10.06  0.57 4.25  0.57 274.65 6 12.46

Agitated 42.24  5.55 27.66  1.99 1.91  0.29 2.97  0.28 5.70  0.50 1.28  0.02 35.65  1.60 10.28  0.79 3.68  0.22 244.80 6 21.16

Parent plant material Leaves 29.69  1.42 34.50  2.16 49.10  2.89 25.86  3.11 3.43  0.32 2.66  0.50 41.01  3.82 22.27  1.99 10.91  1.28 322.83 6 27.06

Fruits 132.39  9.42 109.40  8.26 46.06  3.09 25.48  2.50 4.73  0.99 3.62  0.42 60.72  5.20 66.50  2.51 6.10  1.03 754.56 6 52.78

Table 1 The total content and the maximal amounts (mg/100 g DW  SD) of the main detected dibenzocyclooctadiene lignans depending on the culture type tested, in biomass extracts from S. chinensis in vitro cultures, and in extracts from fruits and leaves of parent plant

11 Different Types of In Vitro Cultures of Schisandra chinensis and Its. . . 319

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schisandrin, gomisin B, and deoxyschisandrin, whose amounts were comparable to their contents in extracts from leaves of the parent plant (Table 1) [75].

2.2

Optimization of Culture Lighting Conditions

The aim of the study was to investigate the influence of light conditions of in vitro cultures on the accumulation of dibenzocyclooctadiene lignans. S. chinensis agar microshoot cultures were grown on a basic MS medium containing 3 mg/l BA and 1 mg/l NAA. The cultures were grown under continuous light of various spectra: farred, red, blue lights, and UV-A irradiation, and in darkness during the course of the whole experiment. Additionally, the control cultures were maintained under continuous white light [77]. The culture growth cycles lasted 30 days. The amounts of dibenzocyclooctadiene lignans (14 compounds) were determined in methanolic biomass extracts [76] (Table 2). Blue light was the most favorable for the accumulation of lignans – their total content was equal to 376.41 mg/100 g DW and was 1.31-fold higher than in biomass extracts cultured under white light. The contents of individual compounds increased from 1.19- to 1.87-fold depending on the applied light conditions. Quantitatively dominant metabolites were schisandrin (67.70 mg/ 100 g DW), deoxyschisandrin (55.19 mg/100 g DW), and gomisin A (36.97 mg/ 100 g DW) (Table 2) [77].

2.3

Optimization of Elicitation Processes

The concept of the study was to achieve efficient methods of S. chinensis microshoot culture elicitation in order to obtain high production of dibenzocyclooctadiene lignans. Table 2 The maximal total content and the maximal amounts (mg/100 g DW  SD) of the main detected dibenzocyclooctadiene lignans in microshoot cultures of S. chinensis cultured under different lighting conditions

Lignans Schisandrin Gomisin A Gomisin G Schisantherin A Schisantherin B Schisanhenol Deoxyschisandrin γ-Schisandrin Schisandrin C Total content

Maximal content 67.70  4.02 36.97  2.19 3.05  0.45 5.26  0.89 28.13  2.09 1.57  0.48 55.19  3.71 16.77  1.96 6.01  0.89 376.41  28.00

Optimal monochromatic light quality Blue light Blue light UV-A irradiation Far-red light Blue light Blue light Blue light Blue light Blue light Blue light

Control (white light) 52.08  3.94 25.41  1.76 2.05  0.94 2.81  0.33 22.89  2.09 1.10  0.25 45.73  2.66 14.06  1.02 4.80  0.22 286.62  22.65

Content increase toward control 1.30 1.45 1.49 1.87 1.23 1.43 1.21 1.19 1.25 1.31

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The study tested different concentrations and time of MS medium (containing 3 mg/l BA and 1 mg/l NAA) supplementation with four elicitors: cadmium chloride (CdCl2), chitosan (Ch), methyl jasmonate (MeJa), and yeast extract (YeE), and a permeabilizing agent – dimethyl sulfoxide (DMSO) [78]. A single culture cycle lasted 30 days. Assays were carried out in methanolic extracts from lyophilized biomass and culture media using the LC-DAD technique [76]. A significant influence of the test factors on the accumulation of lignans in biomass was found (Table 3). Only traces of lignans ( 1 η =
1 if β <
1 η = β if
1  β  1 η = 1 if β > 1 η = 0 if β < 0 η = β if β  0

Satlin

Symmetric saturating linear

Positive linear

Satlins

Poslin

Sigmoid

1 η ¼ 1þ exp
β

Hyperbolic tangent sigmoid

η ¼ eeβ
e þe
β

Tansig

Competitive

η = 1 if β maximum η = 0 other

Compet

Radial basis transfer function

η = e
β2

Radbas

β


β

Symbol

Logsig

(Fig. 7b). Multilayer ANNs are the most used architecture for modeling and optimizing the production of secondary metabolites by plant biotechnology. The feedback network or recurrent network (Fig. 7c) may have any architecture including loops which return to the inputs the value of one or more outputs. In general, the architectural choices for the construction of the ANN are the activation function and the number of neurons in the hidden layer. The model is more robust if the ANN consists of a reduced number of neurons, the model is more efficient if the ANN consists of a large number of neurons. Statistical measures such as the coefficient of determination R2 and the RMSE (Eq. 17) performed during the training and validation phases will allow the selection of the best ANN architecture. sffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffi n 1X RMSE ¼ ðy
yest Þ2 n i¼1 i where yi and yest are the measured and the predicted response, respectively.

ð17Þ

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Fig. 7 ANNs Architecture: Feed forward ANN (a: monolayers. b: Multilayer) and feedback network or recurrent ANN (c)

The ANN Learning Using an iterative algorithm, the ANN will seek to adjust the synaptic weights associated with each explanatory variable to minimize an error function: this is the training phase. Several training algorithms have been tested in plant biotechnology such as Gradient decent, Gradient decent with adaptive learn rate [131], LevenbergMarquardt [56, 131], and standard feed-forward back propagation [132]. Continuing multiple iteration cycles will result in overfitting of the ANN. To avoid this, the experimental data is divided into two sets: the training data that will be used for the algorithm in adjusting synaptic weights and the validation data which is only used to stop training at the threshold of overfitting.

4.2.2 Applications ANNs have been successfully applied in plant biotechnology to develop production processes that maximize the biomass and secondary metabolites content (Table 2). In order to develop a high performance process for the production of secondary metabolites of pharmaceutical interest by the cultivation of Glycyrrhiza glabra HRs in bioreactors, Prakash et al. [133] compared the efficiency of two types of ANNs: regression neural network and back propagation neural network. The input layer variables selected by the authors are inoculum density, sucrose concentration, pH, and volume of culture medium. The output layer variable represents the biomass of G. glabra HRs. The results show that both ANNs are effective in predicting optimal conditions with better accuracy for neural network regression.

Layer recurrent

Regression neural network Cascade forward

Biomass Biomass L-Hyoscyamine content Biomass

3 3

Biomass

Output Biomass

3

Biomass

Biomass

Inoculum size; mist ON time; mist OFF time; initial packing density; media volume; sucrose; time of culture

Inoculum size; mist ON time; mist OFF time; initial packing density; media volume; sucrose; time of culture

Inoculum density; pH; sucrose; volume of growth 4 in layer 1 medium 1 in layer 2

pH; sucrose; volume of medium; inoculum density; nitrate concentration Mg; Cu; Zn; nitrate; sucrose Salicylic acid concentration; exposure time

Network Number of Architecture Input neurons Feed Inoculum density; pH; sucrose; volume of growth 2 forward medium Inoculum size; mist ON time; mist OFF time; initial packing density; media volume; sucrose; time of culture - Gradient decent - Gradient decent with adaptive learn rate - LevenbergMarquardt Gradient descent

Training algorithm Gradient decent

Purelin (output layer) Purelin (output layer)

Tansig (hidden layer) Purelin (output layer)

[133]

[135] [56]

[132]

[131]

References [133]

- Gradient decent; [131] - Gradient decent with adaptive learn rate - LevenbergMarquardt - Gradient decent [131] - Gradient decent with adaptive learn rate - LevenbergMarquardt

Sigmoid Gradient descent TanH hyperbolic tangent LevenbergMarquardt Radbas (layer 1) Gradient descent Purelin (layer 2)

Activation function Purelin (output layer) Logsig (input layer) Tansig (hidden layer) Purelin (output layer)

Table 2 ANNs used for optimal conditions prediction maximizing increments of biomass and secondary metabolites content

822 R. Amdoun et al.

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Osama et al. [131] modeled the influence of Nutrient Mist Reactor on the final biomass of Artimisia annua HRs in different growing conditions. The input layer consists of seven variables: inoculum size, mist ON time, mist OFF time, initial packing density, media volume, initial sucrose concentration in media, and time of culture. The output layer represents the weight of the final biomass of the HRs. Three network architectures were studied, namely, Feed Forward, Cascade Forward, and Layer Recurrent. Each network has been trained with three types of algorithms. These are decent gradient algorithm, gradient descent algorithm with adaptive learn rate, and the Levenberg-Marquardt algorithm. Each network uses the tansig and purelin activation functions for the hidden layer and the output layer, respectively. The results show that the Levenberg-Marquardt training algorithm is the fastest and most accurate. ANNs are effective in modeling the growth of HRs in a Nutrient Mist Reactor, effectively predicting biomass productivity for different growing conditions. All networks were effective, but the recurrent neural network is the most accurate with 0.9679 of correlation coefficient. Centellosides extract from Centella asiatica are secondary metabolites whose specific constituents such as Asiatic acid, Asiaticoside, Madecassic acid, and Madecassoside are potentially antioxidants, anti-inflammatories, anticancer, and antidiabetic [134]. The ANN approach is applied by Prasad et al. [135] to determine the synergistic effect of five components of the culture medium maximizing the shoot biomass of Centella asiatica grown in vitro. The input layer variables are: Mg, Cu, Zn, nitrate, and sucrose. The back propagation neural network (BPNN) was used to predict optimal biomass for 35 days of culture. The optimum conditions of the culture medium to reach a maximum growth index of 1654.46 with a high yield of centellosides (62.37 mg DW) are: 1.5 mM Mg, 30 μM Zn, 0.1 μM Cu, 40 mM NO3, and 6% sucrose. According to the same authors, these results can be valued by the pharmaceutical industry for the production of these metabolites.

4.3

Kriging

Kriging is a linear unbiased approximation probabilistic method used for spatial interpolation [136–141]. Georges Matheron formalized kriging during the 1970s from the work of Danie G. Krige on the Witwatersrand deposit (South Africa) [136, 137]. Kriging is applied in empirical modeling for approximating functional relationships between impact factors and system response [142]. This modeling approach is now widely used in the oil industry, engineering, meteorology, and ecology [143–145]. However, only a few works using kriging in plant biotechnology have been reported in the literature [56, 146]. As with ANNs, Kriging is able to make complex function approximations but cannot easily separate statistically the influence of each factor and their interactions.

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Theory

The Semi-variogram The semi-variogram is the fundamental tool in the geostatistical study of natural phenomena spatially correlated. The natural phenomenon is considered as a random variable Z(x) in a domain D, defined by its probability distribution function: FV(x) = Prob(X  x) 8x; x  D. The semi-variogram γ(h) represents the moment of order 2 of a new random variable which is none other than the difference Z(x)
Z(x + h) which depends only on h: h i 1 1 γ ðhÞ ¼ Var½ZðxÞ
Z ðx þ hÞ ¼ E ðZ ðxÞ
Z ðx þ hÞÞ2 2 2

ð18Þ

This theoretical expression is approximated in practice by the experimental semivariogram and written according to the sample observations Z(xi): γ ð hÞ ¼

N X 1 ðzðxi Þ
zðxi þ hÞÞ2 2  N ðhÞ i¼1

ð19Þ

This function, usually increasing as a function of h (the variance between observations increases as the distance that separating them increases), γ(h) synthesizes a lot of information concerning the joint behavior of the random variables and the “continuity” of the phenomenon. The following parameters are used to describe semi-variogram models (Fig. 8): • The range a: it is the distance beyond which two observations are no longer similar on average; they are no longer linearly linked (zero covariance). At this distance, the semi-variogram value corresponds to the variance of the random variable σ 2. • The sill is equal to the variance of the random variable (Var[Z(x)]): σ 2 = C0 + C1. • The nugget effect C0: it can represent the variation on a very short scale, errors of localization or analysis. Sometimes the semi-variogram does not show a plateau (in this case, covariance and variance do not exist). However, when the semi-variogram shows a plateau then we can easily establish the link between γ(h) and the covariance function C(h) that are actually equivalent in that they provide the same information about the process: γ ðhÞ ¼ σ 2
CðhÞ

ð20Þ

When the range is reached, there is no more covariance between the random variables: C(h) = 0 si h  a.

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825

Fig. 8 Theoretical semivariogram

Ordinary Kriging There are different types of Kriging techniques, such as ordinary kriging, universal kriging, co-kriging, and others. Ordinary kriging is the most frequently used. All linear interpolators have in common Eq. 21 which produces an estimate Z* (xp) at the unknown point xp using a linear combination of n observations (xi). The major difference between these interpolators is the way to determine the weights λi = {λ1, λ2, . . .λn}T assigned to each of the known values. n   X λi :Z ðxi Þ Z  xp ¼

ð21Þ

i¼1

If we assume that Z(xp) is the true value in xp, then kriging poses two important conditions: 1. Minimum estimate variance: that is, Var[Z(xp)- Z(xp)] is minimal.    

σ 2e ¼ Var Z xp
Z xp n X n n X   X  

 

¼ Var Z xp þ λi λj  Cov Z ðxi Þ, Z xj
2  λi  Cov Z xp , Z ðxi Þ i¼1 j¼1

i¼1

ð22Þ 2. Unbiased estimator, i.e.,: n X

λi ¼ 1

ð23Þ

i¼1

where: n n X   X E Z  xp ¼ λi  E½Z ðxi Þ ¼ λi  m ¼ m 1

ð24Þ

1

The weights λi are the unknowns. To minimize σ 2e (Eq. 22), we have to solve the system of partial derivatives:

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dσ 2e ¼ 0, i ¼ 1 . . . n dλi

ð25Þ

The unbiased partial differential Eq. system to be solved will become a system with n + 1 Eqs. and only n unknowns: 8 2 dσ e > > ¼0 < dλi ,i ¼ 1...n n P > > : λi ¼ 1

ð26Þ

i¼1

To balance this system of Eqs. (with n + 1 Eqs. and n unknowns), we introduce a new unknown which is the Lagrange multiplier μ. We form the Lagrangian that must be minimized to find the n unknowns whose resolution makes it possible to obtain the weights λi for Eq. 25: LðλÞ ¼ σ 2e

þ 2μ

 

¼ Var Z xp þ

n X

!

λi i¼1 n X n X


1

 

λi :λj Cov Zðxi Þ, Z xj

i¼1 j¼1


2

n X

n X  

λi :Cov Z xp , Z ðxi Þ þ 2μ λi
1

i¼1

ð27Þ

!

i¼1

The minimum estimate variance σ e2, also called σ k2 kriging variance, distinguishes kriging from other estimation methods. The latter allows an appreciation of the quality of the estimate obtained in each kriged point. It is written as: n   X  

σ 2k ¼ σ 2e ¼ Var Z xp
λi  Cov Z xp , Zðxi Þ
μ

ð28Þ

i¼1

where: σ 2k ¼

n X

    λi  γ xp , xi
γ xp , xp
μ

ð29Þ

i¼1

where γ(xp,xi) the value of γ(h) for the separating vector xp of xi and μ is the Lagrange multiplier.

4.3.2 Applications The production of primary metabolites such as enzymes used in the food industry is an important part of industrial biotechnology. Freier al [146]. used kriging to maximize the production of the GFP protein (green fluorescent protein, primary

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metabolite) secreted by the bacterium Corynebacterium glutamicum. The authors conclude that Kriging is a powerful and effective approach to biotechnology research. The most statistically influential factors were screened by the application of the FFD. Subsequently, the functional relationship between the selected components was studied in detail by an iterative procedure. At each iteration, kriging interpolations were used to design the following experiments. Optimizing the composition of C. glutamicum culture medium has doubled GFP production. A comparative study of the accuracy of three modeling approaches, namely, RSM, ANN, and kriging, to predict L-hyoscyamine content of elicited Datura stramonium HRs shows that the kriging approach is more accurate than the RSM. It is however less precise than the ANN. Indeed, the probability of having an interpolation error σ