Plastids: Methods and Protocols (Methods in Molecular Biology, 2776) [2 ed.] 1071637258, 9781071637258

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Methods in Molecular Biology 2776

Eric Maréchal  Editor

Plastids Methods and Protocols Second Edition

METHODS

IN

MOLECULAR BIOLOGY

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

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

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

Plastids Methods and Protocols Second Edition

Edited by

Eric Maréchal Laboratoire de Physiologie Cellulaire et Végétale, IRIG, CEA-Grenoble, CNRS, CEA, INRAE, Univ. Grenoble Alpes, Grenoble, France

Editor Eric Mare´chal Laboratoire de Physiologie Cellulaire et Ve´ge´tale, IRIG, CEA-Grenoble CNRS, CEA, INRAE, Univ. Grenoble Alpes Grenoble, France

ISSN 1064-3745 ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-0716-3725-8 ISBN 978-1-0716-3726-5 (eBook) https://doi.org/10.1007/978-1-0716-3726-5 © The Editor(s) (if applicable) and The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024 This work is subject to copyright. All rights are solely and exclusively licensed by the Publisher, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. The publisher, the authors, and the editors are safe to assume that the advice and information in this book are believed to be true and accurate at the date of publication. Neither the publisher nor the authors or the editors give a warranty, expressed or implied, with respect to the material contained herein or for any errors or omissions that may have been made. The publisher remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. Cover Caption: Plastids capture CO2 and are the source of atmospheric O2. © Design by Nolwenn Gue´guen, LPCV, Grenoble, France. This Humana imprint is published by the registered company Springer Science+Business Media, LLC, part of Springer Nature. The registered company address is: 1 New York Plaza, New York, NY 10004, U.S.A. Paper in this product is recyclable.

Preface Following the success of its first edition, the second edition of Plastids: Methods and Protocols attempts to provide an updated view of classical and novel protocols required, when studying this key organelle in eukaryotic cells. A long time ago, on our planet, life has emerged, most likely from an initial prebiotic environment containing self-replicating RNA. Our understanding of the transition to the DNA-RNA-protein replicative system, characterizing what is meant by living cells and viruses, is poor. Nevertheless, progresses are being made thanks to the exponential increase of genomic sequences stored in gigantic databases, covering most branches of the tree of life, and to the improvement of mathematical methods used to mine and compare these data. Sophisticated reconstructions of phylogenies have moved the frontiers to the unknown deeper in the past. The precise series of very ancient events that have occurred more than one billion years ago is probably unattainable. Nevertheless, some major events appear as irreversible milestones: emergence of the first cell(s), elaboration of the ribosomes, radiation of the three domains of life (Bacteria, Archaea, Eukarya), development of the endomembrane system and the nucleus, acquisition of the mitochondrion, and eventually acquisition of the chloroplast. This being stayed, not all evolutionary events have been “memorized” in the molecular phylogenies of DNA or protein sequences. Key events, such as the emergence of thylakoids in cyanobacteria, are now reconstructed using other approaches, based, for instance, on the evolution of membrane glycerolipid phase transitions. Morphological, structural, biochemical, functional, molecular, and genetic evidence are consistent with a unique event, known as a “primary endosymbiosis,” at the origin of the “chloroplast” and all kinds of “primary plastids.” An ancestral photosynthetic prokaryote, related to present-day cyanobacteria, has been engulfed by an ancestral mitochondriate eukaryote. During this process of primary endosymbiosis, other prokaryotes, possibly a third partner related to present-day Chlamydiae, have provided some genetic material and have been critical to the success of the conversion of the initial primary endosymbiont into a semi-autonomous organelle transmitted from cell to cell by mitosis. The presence of the chloroplast, delineated by an envelope made of two membranes, has led to a dramatic reorganization of the bioenergetic and metabolic solutions used by eukaryotes to live in an oxygenic environment. Photosynthesis has allowed the capture of energy from light, of carbon from the atmosphere, and water and few minerals were just required to grow, divide, and colonize the environment, making up the foundation of ecosystems. Primary endosymbionts are called the Archaeplastidae. Three main lineages of algae derive from primary endosymbiosis: the Glaucophyta, the Rhodophyta (or Red algae), and the Chlorophyta (or Green algae). Plants derive from the Green algae, forming altogether the Viridiplantae super-phylum. A puzzling question has remained unanswered for decades until late in the twentieth century. How could plastids be also present in eukaryotic cells, but bounded by more than two membranes, i.e., three or four limiting membranes, sometimes with membrane connections with the nuclear envelope or the endoplasmic reticulum? Some of these protists were sometimes considered as branches of protozoa, such as the human parasites responsible for malaria or toxoplasmosis, containing a non-photosynthetic plastid limited by four membranes. Indeed, phytoplankton biodiversity comprise mostly photosynthetic protists

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of this kind, harboring so-called “complex” or “secondary plastids.” Knowledge has advanced. It is now clear that multiple events of “secondary endosymbiosis” have occurred, when ancestral algae, harboring primary plastids, have been engulfed by secondary eukaryotic cells, leading to this extremely complex cellular organization, which now needs to be characterized. The study of the chloroplast and primary plastids has been essential to the understanding of algae and plants, including crops. The study of secondary plastids is in its infancy and is critical to push the border to the unknown in branches of the biodiversity, populating oceanic and terrestrial biosystems and ignored until now. Eventually, understanding the structure and function of all types of plastids is a key scientific question for basic science, agronomy, biotechnology, and all recent developments aiming at capturing CO2, producing biofuels, and developing bio-sourced green chemistry. This book contains therefore updated and new introductory chapters summarizing our current view on plastid evolution, structure, and function, illustrating all the challenges we are facing to comprehend this unique organelle with multiple faces. This book then compiles methods, from robust techniques developed to visualize, fractionate, purify, and study primary plastids from plant materials to most recent techniques developed to study secondary plastids. Methods to analyze plastids by integrated biology strategies based on genetics, genomics, proteomics, or lipidomics are presented. Cutting-edge techniques to engineer plastid-localized metabolic pathways and processes are detailed. Eventually, key bioinformatic tools and databases are described. This book is the result of the contribution of prominent scientists, who have pioneered in the field of primary and secondary plastid biology. It is not exhaustive, and important methods are missing. Nevertheless, thanks to this collaborative effort, Plastids: Methods and Protocols represents one of the most complete compilation of methods and strategies. We hope therefore that reading these chapters will be useful to students, engineers, and researchers who explore this fascinating organelle, which is far from being fully understood. Readers are also encouraged to consider the first edition of Plastids for methods not repeated here. Grenoble, France

Eric Mare´chal

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

PART I

v ix

PLASTID EVOLUTION, PLASTICITY, STRUCTURAL, AND FUNCTIONAL DIVERSITY

1 How Did Thylakoids Emerge in Cyanobacteria, and How Were the Primary Chloroplast and Chromatophore Acquired? . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Eric Mare´chal 2 Complex Endosymbioses I: From Primary to Complex Plastids, Serial Endosymbiotic Events . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 21 ¨ ssy and Miroslav Obornı´k Zolta´n Fu 3 Complex Endosymbiosis II: The Nonphotosynthetic Plastid of Apicomplexa Parasites (The Apicoplast) and Its Integrated Metabolism . . . . . . . . . . . . . . . . . . . . 43 Nyamekye Quansah, Charital Sarah, Yoshiki Yamaryo-Botte´, and Cyrille Y. Botte´ 4 Diversification of Plastid Structure and Function in Land Plants . . . . . . . . . . . . . . 63 Henrik Aronsson and Katalin Solymosi 5 The Main Functions of Plastids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 89 Marcel Kuntz, Laura Dimnet, Sara Pullara, Lucas Moyet, and Norbert Rolland 6 Plastid Transient and Stable Interactions with Other Cell Compartments . . . . . . 107 Stefanie J. Mueller-Schuessele, Se´bastien Leterme, and Morgane Michaud

PART II

LABORATORY PROTOCOLS

7 Purification of Chloroplast Envelope, Thylakoids, and Stroma from Angiosperm Leaves. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Maryse A. Block, Catherine Albrieux, and Eric Mare´chal 8 Isolation of the Inner and Outer Membranes of the Chloroplast Envelope from Angiosperms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Maryse A. Block and Eric Mare´chal 9 Analysis of a Super-Complex at Contact Sites Between Mitochondria and Plastids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Morgane Michaud 10 Isolation of High-Quality Plastids from the Diatom Phaeodactylum tricornutum . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Fan Hu, Wenxiu Yin, Teng Huang, and Hanhua Hu 11 Determining the Subcellular Localization of Proteins in the Different Membranes of Diatom Secondary Plastid . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Xiaojuan Liu and Yangmin Gong

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Monitoring of Lipid Fluxes Between Host and Plastid-Bearing Apicomplexan Parasites . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Sarah Charital, Amandine Lourdel, Nyamekye Quansah, Cyrille Y. Botte´, and Yoshiki Yamaryo-Botte´ 13 Extraction and Quantification of Lipids from Plant or Algae . . . . . . . . . . . . . . . . . Vale´rie Gros, Josselin Lupette, and Juliette Jouhet 14 Quantitative Assessment of the Chloroplast Lipidome . . . . . . . . . . . . . . . . . . . . . . . Vale´rie Gros and Juliette Jouhet 15 The Use of Nanopore Sequencing to Analyze the Chloroplast Transcriptome Part I: Library Preparation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Se´bastien Skiada, Alexandra Launay-Avon, Arnaud Liehrmann, Etienne Delannoy, and Benoıˆt Castandet 16 The Use of Nanopore Sequencing to Analyze the Chloroplast Transcriptome Part II: Bioinformatic Analyzes and Virtual RNA Blots . . . . . . . . Etienne Delannoy, Arnaud Liehrmann, and Benoıˆt Castandet 17 Targeted Gene Editing of Nuclear-Encoded Plastid Proteins in Phaeodactylum tricornutum via CRISPR/Cas9 . . . . . . . . . . . . . . . . . . . . . . . . . . . Ce´cile Giustini, Jhoanell Angulo, Florence Courtois, and Guillaume Allorent 18 Isolation of Cytosolic Ribosomes Associated with Plant Mitochondria and Chloroplasts. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Laura Dimnet, Thalia Salinas-Giege´, Sara Pullara, Lucas Moyet, Chloe´ Genevey, Marcel Kuntz, Anne-Marie Ducheˆne, and Norbert Rolland

PART III 19

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IN SILICO TOOLS

A Practical Guide to the Representation of Protein Regulation in the Web Application ChloroKB . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 305 Gilles Curien

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

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Contributors CATHERINE ALBRIEUX • Laboratoire de Physiologie Cellulaire et Ve´ge´tale, CNRS, CEA, INRAE, Univ. Grenoble Alpes, IRIG, CEA Grenoble, Grenoble, France GUILLAUME ALLORENT • Laboratoire de Physiologie Cellulaire et Ve´ge´tale, CNRS, CEA, INRAE, Univ. Grenoble Alpes, IRIG, CEA Grenoble, Grenoble, France JHOANELL ANGULO • Laboratoire de Physiologie Cellulaire et Ve´ge´tale, CNRS, CEA, INRAE, Univ. Grenoble Alpes, IRIG, CEA Grenoble, Grenoble, France HENRIK ARONSSON • Department of Biological and Environmental Sciences, University of Gothenburg, Gothenburg, Sweden MARYSE A. BLOCK • Laboratoire de Physiologie Cellulaire et Ve´ge´tale, CNRS, CEA, INRAE, Univ. Grenoble Alpes, IRIG, CEA Grenoble, Grenoble, France CYRILLE Y. BOTTE´ • ApicoLipid Team, Institute for Advanced Biosciences, UMR5309, Centre National de la Recherche Scientifique, Universite´ Grenoble Alpes, U1209, Institut National de la Sante´ et de la Recherche Me´dicale, Grenoble, France; Centre National de la Recherche Scientifique, Institute for Advanced Biosciences, UMR5309, Universite´ Grenoble Alpes, INSERM, U1209, Grenoble, France BENOIˆT CASTANDET • Institute of Plant Sciences Paris-Saclay (IPS2), Universite´ Paris-Saclay, CNRS, INRAE, Universite´ Evry, Gif sur Yvette, France SARAH CHARITAL • ApicoLipid Team, Institute for Advanced Biosciences, CNRS UMR5309, Universite´ Grenoble Alpes, INSERM U1209, Grenoble, France FLORENCE COURTOIS • Laboratoire de Physiologie Cellulaire et Ve´ge´tale, CNRS, CEA, INRAE, Univ. Grenoble Alpes, IRIG, CEA Grenoble, Grenoble, France GILLES CURIEN • Laboratoire de Physiologie Cellulaire et Ve´ge´tale, CNRS, CEA, INRAE, Univ. Grenoble Alpes, IRIG, CEA Grenoble, Grenoble Cedex 9, France ETIENNE DELANNOY • Institute of Plant Sciences Paris-Saclay (IPS2), Universite´ ParisSaclay, CNRS, INRAE, Universite´ Evry, Gif sur Yvette, France LAURA DIMNET • Laboratoire de Physiologie Cellulaire et Ve´ge´tale, CNRS, CEA, INRAE, Univ. Grenoble Alpes, IRIG, CEA Grenoble, Grenoble, France ANNE-MARIE DUCHEˆNE • Institut de biologie mole´culaire des plantes, CNRS, Universite´ de Strasbourg, Strasbourg Cedex, France ZOLTA´N FU¨SSY • Scripps Institution of Oceanography, University of California San Diego, La ˇ eske´ Budeˇjovice, Czech Jolla, CA, USA; Faculty of Science, University of South Bohemia, C Republic CHLOE´ GENEVEY • Laboratoire de Physiologie Cellulaire et Ve´ge´tale, CNRS, CEA, INRAE, Univ. Grenoble Alpes, IRIG, CEA Grenoble, Grenoble, France CE´CILE GIUSTINI • Laboratoire de Physiologie Cellulaire et Ve´ge´tale, CNRS, CEA, INRAE, Univ. Grenoble Alpes, IRIG, CEA Grenoble, Grenoble, France YANGMIN GONG • Oil Crops Research Institute of the Chinese Academy of Agricultural Sciences, Wuhan, China VALE´RIE GROS • Laboratoire de Physiologie Cellulaire et Ve´ge´tale, CNRS, CEA, INRAE, Univ. Grenoble Alpes, IRIG, CEA Grenoble, Grenoble, France FAN HU • School of Foreign Languages, China University of Geosciences, Wuhan, China HANHUA HU • Key Laboratory of Algal Biology, Institute of Hydrobiology, Chinese Academy of Sciences, Wuhan, China

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TENG HUANG • Key Laboratory of Algal Biology, Institute of Hydrobiology, Chinese Academy of Sciences, Wuhan, China JULIETTE JOUHET • Laboratoire de Physiologie Cellulaire et Ve´ge´tale, CNRS, CEA, INRAE, Univ. Grenoble Alpes, IRIG, CEA Grenoble, Grenoble, France MARCEL KUNTZ • Laboratoire de Physiologie Cellulaire et Ve´ge´tale, CNRS, CEA, INRAE, Univ. Grenoble Alpes, IRIG, CEA Grenoble, Grenoble, France ALEXANDRA LAUNAY-AVON • Institute of Plant Sciences Paris-Saclay (IPS2), Universite´ ParisSaclay, CNRS, INRAE, Universite´ Evry, Gif sur Yvette, France SE´BASTIEN LETERME • Laboratoire de Physiologie Cellulaire et Ve´ge´tale, CNRS, CEA, INRAE, Univ. Grenoble Alpes, IRIG, CEA Grenoble, Grenoble, France ARNAUD LIEHRMANN • Institute of Plant Sciences Paris-Saclay (IPS2), Universite´ ParisSaclay, CNRS, INRAE, Universite´ Evry, Gif sur Yvette, France; Laboratoire de Mathe´ matiques et de Mode´lisation d’Evry (LaMME), Universite´ d’Evry-Val-d’Essonne, UMR CNRS 8071, ENSIIE, USC INRAE, Evry, France XIAOJUAN LIU • Shantou University, Shantou, Guangdong, China AMANDINE LOURDEL • ApicoLipid Team, Institute for Advanced Biosciences, CNRS UMR5309, Universite´ Grenoble Alpes, INSERM U1209, Grenoble, France JOSSELIN LUPETTE • Laboratoire de Physiologie Cellulaire et Ve´ge´tale, CNRS, CEA, INRAE, Univ. Grenoble Alpes, IRIG, CEA Grenoble, Grenoble, France ERIC MARE´CHAL • Laboratoire de Physiologie Cellulaire et Ve´ge´tale, IRIG, CEA-Grenoble, CNRS, CEA, INRAE, Univ. Grenoble Alpes, Grenoble, France MORGANE MICHAUD • Laboratoire de Physiologie Cellulaire et Ve´ge´tale, CNRS, CEA, INRAE, Univ. Grenoble Alpes, IRIG, CEA Grenoble, Grenoble, France LUCAS MOYET • Laboratoire de Physiologie Cellulaire et Ve´ge´tale, CNRS, CEA, INRAE, Univ. Grenoble Alpes, IRIG, CEA Grenoble, Grenoble, France STEFANIE J. MUELLER-SCHUESSELE • Molecular Botany, Department of Biology, RPTU, Kaiserslautern, Germany ˇ eske´ Budeˇjovice, MIROSLAV OBORNI´K • Faculty of Science, University of South Bohemia, C ˇ eske´ Czech Republic; Institute of Parasitology, Biology Centre, Czech Academy of Sciences, C Budeˇjovice, Czech Republic SARA PULLARA • Laboratoire de Physiologie Cellulaire et Ve´ge´tale, CNRS, CEA, INRAE, Univ. Grenoble Alpes, IRIG, CEA Grenoble, Grenoble, France NYAMEKYE QUANSAH • ApicoLipid Team, Institute for Advanced Biosciences, UMR5309, Centre National de la Recherche Scientifique, Universite´ Grenoble Alpes, U1209, Institut National de la Sante´ et de la Recherche Me´dicale, Grenoble, France; ApicoLipid Team, Institute for Advanced Biosciences, CNRS UMR5309, Universite´ Grenoble Alpes, INSERM U1209, Grenoble, France NORBERT ROLLAND • Laboratoire de Physiologie Cellulaire et Ve´ge´tale, CNRS, CEA, INRAE, Univ. Grenoble Alpes, IRIG, CEA Grenoble, Grenoble, France THALIA SALINAS-GIEGE´ • Institut de biologie mole´culaire des plantes, CNRS, Universite´ de Strasbourg, Strasbourg Cedex, France CHARITAL SARAH • ApicoLipid Team, Institute for Advanced Biosciences, UMR5309, Centre National de la Recherche Scientifique, Universite´ Grenoble Alpes, U1209, Institut National de la Sante´ et de la Recherche Me´dicale, Grenoble, France SE´BASTIEN SKIADA • Institute of Plant Sciences Paris-Saclay (IPS2), Universite´ Paris-Saclay, CNRS, INRAE, Universite´ Evry, Gif sur Yvette, France KATALIN SOLYMOSI ¨tvo¨s • Department of Plant Anatomy, Institute of Biology, ELTE Eo Lora´nd University, Budapest, Hungary

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YOSHIKI YAMARYO-BOTTE´ • ApicoLipid Team, Institute for Advanced Biosciences, UMR5309, Centre National de la Recherche Scientifique, Universite´ Grenoble Alpes, U1209, Institut National de la Sante´ et de la Recherche Me´dicale, Grenoble, France; ApicoLipid Team, Institute for Advanced Biosciences, CNRS UMR5309, Universite´ Grenoble Alpes, INSERM U1209, Grenoble, France WENXIU YIN • Key Laboratory of Algal Biology, Institute of Hydrobiology, Chinese Academy of Sciences, Wuhan, China

Part I Plastid Evolution, Plasticity, Structural, and Functional Diversity

Chapter 1 How Did Thylakoids Emerge in Cyanobacteria, and How Were the Primary Chloroplast and Chromatophore Acquired? Eric Mare´chal Abstract The emergence of thylakoid membranes in cyanobacteria is a key event in the evolution of all oxygenic photosynthetic cells, from prokaryotes to eukaryotes. Recent analyses show that they could originate from a unique lipid phase transition rather than from a supposed vesicular budding mechanism. Emergence of thylakoids coincided with the great oxygenation event, more than two billion years ago. The acquisition of semi-autonomous organelles, such as the mitochondrion, the chloroplast, and, more recently, the chromatophore, is a critical step in the evolution of eukaryotes. They resulted from primary endosymbiotic events that seem to share general features, i.e., an acquisition of a bacterium/cyanobacteria likely via a phagocytic membrane, a genome reduction coinciding with an escape of genes from the organelle to the nucleus, and, finally, the appearance of an active system translocating nuclear-encoded proteins back to the organelles. An intense mobilization of foreign genes of bacterial origin, via horizontal gene transfers, plays a critical role. Some third partners, like Chlamydia, might have facilitated the transition from cyanobacteria to the early chloroplast. This chapter further details our current understanding of primary endosymbiosis, focusing on primary chloroplasts, thought to have appeared over a billion years ago, and the chromatophore, which appeared around a hundred years ago. Key words Primary endosymbiosis, Chloroplast, Mitochondria, Chromatophore, Archaeplastida

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Introduction: Photosynthetic Membranes in Prokaryotic and Eukaryotic Cells Membrane structures specialized in oxygenic photosynthesis (called thylakoids) are present in both prokaryotic and eukaryotic cells. In the Bacteria domain of life, thylakoid lamellae are present in the cytoplasm of cyanobacteria cells, with the notable exception of a group considered as archaic, comprising Gloeobacter species, in which photosystems are anchored in the cytoplasmic membrane [1]. Thylakoids adopt a much broader variety of structures in cyanobacteria compared to chloroplasts in eukaryotic cells, from multiple parietal membrane layers to sacs piled in stacks [2]. The

Eric Mare´chal (ed.), Plastids: Methods and Protocols, Methods in Molecular Biology, vol. 2776, https://doi.org/10.1007/978-1-0716-3726-5_1, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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emergence and biogenesis of thylakoids in cyanobacteria have remained unresolved questions, until recent hypotheses based on nonvesicular elaboration of thylakoids [3]. Contrasting with prokaryotes, eukaryotic cells are characterized by a complex system of intracellular membranes. These compartments have a variety of shapes, from spherical vesicles to tubular or flattened cisternae, reticulated membrane networks, large-size multilayer subspherical systems, etc. An immense task has consisted in the description and definition of these organelles, mainly based on microscopy imaging techniques, but also cell fractionation and biochemistry, and this task is still incomplete. In eukaryotic cells, large-size organelles can be observed with a basic light microscope. This is the case for the nuclei, as well as vacuoles in plant cells. The chloroplasts are the easiest organelle to observe simply because of their pigmentation: they ‘look’ like cyanobacteria inside the cytoplasm. Smaller organelles or intricate membrane systems need higher-resolution imaging methods, such as electron microscopy, and more recently three-dimensional electron microscopy or tomography. The organization of eukaryotic cells and its dynamics have been initially described in simple cellular models. The organizational principles were then extrapolated to more elaborate models, such as multicellular animals and plants and all kinds of eukaryotic cells that have arisen from simple endosymbiotic events, described in this chapter. Higher levels of cellular complexity have emerged from secondary endosymbiosis events treated in the next chapter. When focusing on photosynthetic membranes, shared by cyanobacteria and chloroplasts, thylakoids form membrane lamellae, with nearly identical lipid compositions [4] and conserved function. Although some proteins are conserved, a large number of chloroplast proteins have diverged or are very different from their cyanobacteria counterparts. This is particularly evident in lipid biosynthesis pathways [4]. Two questions about emergence are therefore puzzling in relation to this remarkable structural conservation: How did thylakoids emerge in cyanobacteria and was a cyanobacteria at the origin of primary chloroplasts?

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How Did Thylakoids Emerge in Cyanobacteria? The origin of a membrane compartment may look simple, when it results from the budding or protrusion of a parent membrane, fusing with other vesicles or growing by membrane lateral expansion. It becomes a difficult question when we lack evidence of any vesicle budding. The biophysical rearrangement of membrane components, most notably lipids, during a fusion process, has

At the Origin of Plastids

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usually a positive enthalpy and requires an external energy source, usually provided by ATP or GTP. Such reorganization is therefore not spontaneous and involves also specific machineries. Thylakoid formation in cyanobacteria is unlikely vesicle-based, first because of the lack of ultrastructural observation of vesicular structures or protrusions generated from the plasma membrane toward the cell interior, but also because primitive thylakoids are concentric, uninterrupted, and it is not clear where their biogenesis could be initiated. A simple hypothesis was recently proposed to resolve the mechanism of this nonvesicular process [3]. It is based on the gradual production of lipids forming a nonbilayer phase, called Hexagonal II (or HexII), at the inner periphery of the cytosolic membrane of a primitive cyanobacterium, transformed in a few nanoseconds and without any external energy source into multilayer pro-thylakoids (Fig. 1). This model is derived from the opposite mechanism described by Marrink and Mark [5], in which lipids arranged in a lamellar phase (Lm) can be converted into a HexII phase when provided with energy (Fig. 1a). As a prerequisite for this model, cyanobacteria cells contain both types: HexII-forming lipids, most importantly monogalactosyldiacylglycerol (MGDG) and Lm-forming lipids, one being neutral, digalactosyldiacylglycerol (DGDG) and two being anionic, phosphatidylglycerol (PG) and sulfoquinovosyldiacylglycerol (SQDG). Most ancient cyanobacteria of Gloeobacter clade, which were unable to generate any thylakoid membrane, had a poor content in anionic Lm lipids, represented only by PG, whose proportion is controlled by the level of phosphorus in the environment. They contain their photosystems in foci at their cell periphery (Fig. 2a) [6]. The acquisition of the SQDG biosynthetic pathway contributed to a more stable supply of anionic lipids, allowing the HexII → Lm phase transition (Fig. 1a) to occur, enabling the formation of parietal thylakoids (Fig. 2b). The emergence of thylakoid coincides with the oxygenation of the atmosphere, during the great oxygenation event (GOE), which have occurred ~2.4 billion years ago. Thus the oxygenation of our atmosphere was not driven directly by the appearance of oxygenic photosynthesis, but on a latter metabolic innovation in thylakoid lipid biosynthesis [3]. This model for thylakoid emergence was elaborated before the description of the spatiotemporal biogenesis process of thylakoid membranes in the rod-shaped cyanobacterium Synechococcus elongatus PCC 7942 [7]. In this cyanobacterium, the plasma membrane and concentric layers of thylakoids have no physical connections. Newly synthesized thylakoid membranes emerge between the plasma membrane and preexisting thylakoids (Huokko et al., 2021). It should be kept in mind that this nonvesicular lipid phase transition does not exclude the involvement of proteins. Nevertheless, the role of companion proteins, such as Vipp1, whose characterization in a hypothetical vesicle-based biogenetic

Eric Mare´chal

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A

HexII phase can disrupt membrane multilayers and facilitate membrane fusion (Marrink & Mark model)

Stalks

Lm Triggers: - addition of HexII-forming lipids - hydration decrease; - temperature increase; - higher acyl desaturation - proteins, ATP

B

Lm

HexII

HexII

H > 0 (needs external energy) within nano- to microseconds

Vice versa, Lm lipids can generate membrane multilayers from an HexII phase via non-vesisular phase transition (origin of thylakoids)

Lm

Stalks

HexII

Triggers: - addition of Lm-forming lipids - hydration increase; - temperature decrease; - proteins; - lower acyl desaturation - no external energy required

HexII

Lm

H < 0 (spontaneous) within nano- to microseconds

Thylakoids

Fig. 1 Nonlamellar-to-lamellar lipid phase transition at the origin of thylakoids. (a) The Lm-to-HexII phase transition has been described as a driving process of membrane fusion, conditioned by a supply of an external energy source. The intermediate phase, forming “stalks” according to the model of Marrink and Mark is shown. (b) HexII to Lm phase transition as a driving process for thylakoid emergence. The major thylakoid lipid, monogalactosyldliacylglycerol, is a HexII lipid forming such structures. Based on the principle of reversibility, the phase transition is spontaneous, occurring within nanoseconds. It can be facilitated by molecular triggers, including Lm-forming lipids such as sulfoquinovosyldiacylglycerol

process remained unsatisfactory both in cyanobacteria and in chloroplasts, is probably not that of a biogenesis driver and needs to be reevaluated.

3

Was a Cyanobacteria at the Origin of Primary Chloroplasts?

3.1 Eukaryotic Cell Membrane Compartmentalization

In a simple eukaryotic cell, two major categories of membrane compartments can be considered: the endomembrane system and the semi-autonomous organelles, i.e., the mitochondrion and the chloroplast.

3.1.1 The Endomembrane System

The “endomembrane system” consists of the endoplasmic reticulum (ER), nuclear envelope, Golgi apparatus, trans-Golgi network, endosomes, vacuolar network, the plasma membrane, etc. All the

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photosynthetic domains (~55% MGDG)

A

HexII

Gloeobacter-type thylakoid-less Cyanobacteria

Lm

HexII-rich matrix (~70% MGDG)

No thylakoids Photosynthetic surface: 1 μm2 range per cell

Acquisition of sulfoquinovosyldiacylglycerol (SQDG), a bilyaer-forming sulfolipid

B

HexII-rich Matrix (MGDG) Cyanobacteria with parietal thylakoids (Cyanobium-, Synechocystis-, Leptolyngbya-type)

HexII → Lm Triggers: PG, SQDG, DGDG + Protein-helpers: Vipp1, etc.

Nanosecond scale

Lm

HexII

Iterations

Lm

HexII

HexII Lm

Hexagonal II-forming lipid (= MGDG) Bilayer-forming lipid PG, SQDG, DGDG)

Parietal thylakoids as concentric multilayers (uninterrupted or in fascicles) Photosynthetic surface: >20 μm2 per cell

Thylakoid expansion is one of the major drivers of the Great Oxygenation Event (~2.4 b.y.a.)

Fig. 2 Origin of thylakoids in early cyanobacteria. (a) In the ancient type of cyanobaceria, like Gloeobacter, cells have domains in their cytosolic limiting membrane, in which photosystems are concentrated, with a balanced composition of HexII and Lm-forming lipids. An MGDG-rich matrix can therefore create local HexIIrich domains. This matrix can possibly form a HexII phase protruding inward. In Gloeobacter, the level of Lm lipids is not sufficient to trigger a transition from HexII to Lm phase. (b) Later in the evolution, in the first SQDGcontaining cyanobacteria, Lm-forming lipids are not limiting and can trigger serial HexII-to-Lm phase transitions, giving rise to concentric parietal thylakoids. Parietal thylakoids can be uninterrupted in Cyanobium cells (as illustrated) or forming fascicles in Synechocystis or Leptolyngbya. Given the increase of photosynthetic membrane surface thus introduced, a multiplier effect is compatible with a decisive role in the accumulation of O2 during the great oxygenation event (GOE)

membranes of the endomembrane system are dynamically connected either by vesicular shuttles or tubules. It is well known that the “eukaryotic” cell is named after the presence of the nucleus; however, the nuclear envelope “disappears” in some eukaryotes during mitosis (in so-called open mitosis). The ER is the only membrane sack that is always present and it can be considered as the core of the endomembrane system. Expansions of the ER make the nuclear envelope; vesicles budding from the ER are at the origin of other compartments, such as the Golgi network, vacuoles, etc., and eventually the plasma membrane. Membrane biogenesis, combining lipid biosynthesis with protein cotranslational membrane insertion, is therefore very intense at the level of the ER, and then membrane lipids and proteins are sorted, remodeled, modified, etc., fitting with the desired composition in the final (or transitory) functional compartments, in which they sit.

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3.1.2 The Mitochondrion, the Primary Chloroplast, and the Chromatophore

Some membrane structures are not connected to this endomembrane system and are called “semi-autonomous organelles.” They contain DNA and need to divide to be transmitted to daughter cells. If they are not transmitted following mitosis or if they are lost or degraded, they cannot be reconstituted from another membrane system. Best-known semi-autonomous organelles are the mitochondrion and the primary chloroplast, both limited by an envelope made of two membranes. A less common semi-autonomous organelle, containing DNA and bounded by two membranes, is the chromatophore, currently restricted to a group of amiboids called the Paulinellidae. Secondary plastids are limited by 3 to 4 membranes and have a hybrid membrane organization, having their outermost membrane connected to the endomembrane system. The architecture of some secondary plastids can even include direct connections with the nuclear envelope and also contain DNA inherited from a nuclear material, called the nucleomorph (see Subheading 2). In the case of secondary plastids, the architecture of the photosynthetic “organelle” is hybrid, and a cooperation between the endomembrane system and the organelle is required for maintenance and transmission; for such subcellular architecture, the definition as ‘semiautonomous organelles’ makes little sense. Therefore, the term ‘semi-autonomous organelle’ needs to be restricted to the mitochondrion, the primary chloroplast, and the chromatophore.

3.1.3 The Origin of Eukaryotes and the Question of the Acquisition of the Semi-autonomous Organelles

Understanding the evolution of eukaryotes and the acquisition of semi-autonomous organelles is a difficult task, probably unattainable [8] and so is the reconstruction of a putative universal tree of life [9]. Reasonable hypotheses are based on molecular features found in genetic sequences, protein structures, conserved metabolites, function, cell architecture, etc., and a few experimental data. Since the 1980s, it is considered that ribosome encoding organisms, excluding viruses, comprise three major groups or domains, called “Archaea,” “Bacteria,” and “Eukarya” [9]. Life is supposed to have emerged from self-replicating systems in an RNA-rich environment. Maybe as early as 4.5–3.8 billion years ago, this initial RNA-based system has transitioned to the modern DNA/RNA/protein system [10, 11]. Whereas limiting membranes predated the origin of life is still a matter of debate; nevertheless, the last unicellular common ancestor (LUCA) is proposed to have emerged more than 3.5 billion years ago, followed by the divergence between Bacteria and Archaea [10–12]. Based on the most recent phylogenetic studies, Eukarya are now considered as a sister group of a superphylum of Archaea, called the ‘Asgard Archaea’ [13]. The transition from Asgard Archea to Eukarya is supposed to have occurred via an intense remodeling of genetic material, including, on the hand, transmission of Archaea genes, tending to be involved in information processes, and on the other

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hand, horizontal gene transfers (HGT) from Bacteria, linked to metabolic processes [13]. This results in the current questioning of the definition of the Eukarya as an actual “domain of life” or more likely a “hybrid” domain originating from Bacteria and Archaea [14]. The acquisition of the mitochondrion and the primary chloroplast is probably one of the most important events in the evolution of eukaryotes. Mitochondria provided a unique gain in bioenergetics efficiency influencing the formation of the eukaryotic cell, as we know it today [15]. Both the mitochondrion and the primary chloroplast are limited by two membranes, both organelles are central in the bioenergetic metabolism, both contain circular DNA, with (nearly) all the necessary machineries for DNA replication and gene translation. They are like guests ensuring the energetic supply of their host cells, and this function is therefore likely central in their successful integration. The reconstruction of the acquisition of these organelles has fascinated biologists for more than a century. In 1905, the Russian biologist C. Mereschkowsky [16] developed a theory postulating that plastids were the evolutionary descendants of endosymbioticlike organisms, calling them the “little green slaves.” From this first visionary insight, the endosymbiotic theory was not considered until the 1970s [17]. Cytological and molecular evidence accumulated and confirmed that plastids have probably originated from the engulfment of an unknown cyanobacterial ancestor inside a eukaryotic cell, predated by the enslavement of a mitochondrion inherited from an α-proteobacterium-like ancestor. Evidence for an endosymbiotic origin includes (1) the presence of “naked” (without histones) organelle-specific DNA, (2) high degrees of sequence homology between DNA of mitochondria and plastids and of bacteria and cyanobacteria, respectively, (3) organelle ribosomes similar to those of prokaryotes and sensitive to chloramphenicol, (4) initiation of messenger RNA translation by means similar to those in prokaryotes, (5) lack of actin/tubulin system, (6) fatty acid biosynthesis occurring via a prokaryote-like fatty acid synthase and coupled to an acyl carrier protein (ACP), and (7) occurrence of β-barrel-membrane proteins in the outer envelope membrane like in gram-negative bacteria and cyanobacteria, etc. This being said, how and when did these endosymbiotic events occur? How many times? How did this association become irreversible, with such a high level of dependence on the host cell? 3.2 The Mechanism of Entry and the Question of the Outermost Membranes of the Mitochondrion and the Chloroplast

The First eukaryotic common ancestor (FECA) is inferred to be identical to the last common ancestor of Archaea and Eukarya [13]. Since we do not know how the endomembrane system was acquired, FECA is shown either containing an ER and a nucleus in Fig. 3 or without any membrane compartments in Fig. 2. About 1 to 1.9 billion years ago, features of modern eukaryotes are evidenced [13], and this is therefore during this period that the Last

10

Eric Mare´chal Unknown α-proteobacterium

1 Asgard-Like Archaeon

2 phagosome? First eukaryotic common ancestor (FECA) Acquisition of the mitochondrion

Emergence of the endomenbrane system

3

The outer membrane of the mitochondria envelope IS NOT the relic of a phagosome

Last eukaryotic common ancestor (LECA) Unknown cyanobacteria

4

NO

YES

phagosome 5

The outer membrane of the chloroplast envelope IS NOT the relic of a phagosome

Acquisition of the chloroplast

Algae and plants = ARCHAEPLASTIDA

YES

NO

Fig. 3 From syntrophy and phagocytosis to endosymbiosis. In this scheme, the First eukaryotic common ancestor (1) is shown containing an endomembrane system (in blue). The Last eukaryotic common ancestor (2) appears when an unknown α-proteobacterium is engulfed within the cell, giving rise to the mitochondrion. Should it be via the formation of a network of cellular “tentacles” or of actual phagosome, the phagosome-like structure is not conserved (3). In this simple scheme, the primary chloroplast derives from the engulfment of an unknown cyanobacterium (4). Again, the phagosome is not conserved (5). The two membanes limiting the mitochondrion and the chloroplast are therefore supposed to derive, mainly, from the outermost membranes of the α-proteobacterium and the cynaobacterium, respectively

eukaryotic common ancestor (LECA) is positioned. The mitochondrion and the primary chloroplast are bounded by two membranes. Alternative models are possible regarding the origin of these membranes (Fig. 3). The ancestral prokaryotes are supposed to have entered the cell via either syntrophy (when two organisms interact without endosymbiosis), specific bacteriotrophy, or unspecific phagocytosis (these two latter mechanisms requiring phagocytosis) [15, 18]. No contemporary example of a bacterial endosymbiont resident in an archaeal host has been described to date, which would support such hypothesis for the acquisition of the mitochondrion [19]. The mechanism of acquisition of the mitochondrion is

At the Origin of Plastids Genes of bacterial origin acquired by horizontal gene transfer (HGT)

Sister group to Asgard Archaea

11

Bacteria-to-Archaea HGT

1

Gene of archaeal origin Gene of a-proteobacterial origin Gene of cyanobacterial origin

Bacteria-toa -proteobacterium HGT

First eukaryotic common ancestor (FECA)

Bacteria-to-pre-LECA HGT 2

Unknown α-proteobacterium

Organelle-to-nucleus LGT

Last eukaryotic common ancestor (LECA) Bacteria-to-cyanobacteria HGT Bacteria-to-eukaryote HGT

Unknown cyanobacteria

3

Organelle-to-organelle LGT

Organelle-to-nucleus LGT 4

First photosynthetic eukaryote (primary endosymbionts)

Fig. 4 Importance of horizontal gene transfers in Eukarya evolution and in primary endosymbioses. In this scheme, the First eukaryotic common ancestor (1) contains genes originating from Archaea and unique Eukarya origin (blue circles). Some bacterial genes (blue square) could be incoporated via Bacteria-to-Archaea horizontal gene transfer (HGT). The acquisition of the mitochondrion could involve Bacteria-to-α-proteobacterium HGT (2), explaining the presence of genes that do not carry an α-proteobacterial signature in mitochondria (brown square). This endosymbiotic event was followed by the escape of some of the mitochondrial genes to the nucleus by a specific HGT, called here lateral gene tranfer (LGT). Likewise, the acquisition of the chloroplast could involve Bacteria-to-cyanobacteria HGT (3), explaining the presence of genes that do not carry a cyanobacterial signature in chloroplasts (green square). Primary endosymbiosis of the chloroplast was followed by the escape of some genes to the nucleus by LGT. Some LGT between both organelles could then have occurred (4)

difficult to reconstruct: It was possibly captured in a net of membrane protrusions acting as a phagocytic membrane (or phagosome) ending with an invagination of the plasma membrane of the host cell (Figs. 1, 2, and 4). The acquisition of the chloroplast seems more simply due to a more classical phagocytic process. Some textbooks propose that the outermost membrane of

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the mitochondria and the primary chloroplast envelope derive from this phagosome. However, the protein and lipid compositions of mitochondria and chloroplast outer envelope membranes do not support this hypothesis and are rather reminiscent of the outermost membrane of an ancestral α-proteobacterium and cyanobacteria, respectively. One can therefore consider that a reasonable scenario is either an entry within the cell, without any phagocytic membrane, or a loss of the phagosome over evolution. An alternative option is that a hybrid membrane system might have appeared, combining features from the ancestral prokaryotic endosymbionts, the eukaryotic host, and possibly other prokaryotic partners (see below). Phylogenetic and molecular analyses support that the mitochondria derive from a unique endosymbiotic event, with a debate whether the α-proteobacterium might be related to the free-living Pelagibacter group or the parasitic Rickettsiales [20]. Different models for eukaryotic origins were proposed, which were compatible with both mitochondrial-early and mitochondria-late scenarios [19], positioning therefore this event in a very broad period from 1.9 to 1 billion years ago. The primary chloroplast derives from a single endosymbiotic event occurring later, from 1.5 to 1 billion years ago [21]. It is believed to originate from, at least, a close relative of the deep-branching Gloeomargarita cyanobacterium, in combination with gene transfers from a variety of other sources [22–24]. 3.3 The Genetic and Molecular Integration of the Mitochondrion and the Chloroplast

Mitochondrial and chloroplast chromosomes exist in both circular and linear forms [25, 26]. They contain conserved genes coding for ribosome components, tRNA, and organellar proteins. They vary in size from a few kilobase pairs to thousands of kilobase pairs. A common feature is that none of these chromosomes is sufficient to ensure the complete biogenesis and maintenance of these organelles. A considerable reduction of the ancestral eubacterial genomes has occurred, following the massive loss and/or transfer of genes to the nucleus of the post-endosymbiotic cell [27, 28]. This loss of autonomy is probably the most achieved in mitochondrial evolution, since DNA-free organelles, bounded by two membranes, are evidenced to derive from mitochondria, i.e., the hydrogenosome, which continues to generate energy for the host cell, and the mitosome, which does not [11]. Presence of cardiolipin, the mitochondrial-specific phospholipid, has been demonstrated in hydrogenosomes [29], whereas at least some of the mitosomes are cardiolipin-free [30]. No such DNA-free organelle deriving from chloroplasts has been observed to date. The most reduced form of a plastid is a secondary plastid, found in Apicomplexa parasites, missing galactolipids that are specific to all plastids studied to that date [4, 31–33]. The function of genes transferred to the nucleus was preserved in many cases, schematically by introducing a short

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additional sequence at the N-terminus of nuclear-encoded proteins, allowing protein precursors to be actively translocated back from the cytosol to the endosymbiotic organelles [34]. Mechanisms allowing the transfer of genetic material from the mitochondrion or the chloroplast to the nucleus are still active. This is actually one of the most robust aspects in our understanding of endosymbiosis, since this transition from free-living organisms to full integration, based on a gene escape from the organelles to the nucleus, is still ongoing and can be evidenced by experiments. The principle is simply to incorporate a gene in the mitochondria of yeast [35] or in the chloroplast of tobacco [36], carrying a nuclear selectable marker gene that allows the efficient selection of yeasts or plants, with a nuclear genome that carries pieces transferred from the mitochondria or the chloroplast genome, respectively. The rate of a gene escape from the mitochondrion to the nucleus is in the range of 2.10-5 per cell per generation in yeast [35]. The rate at which DNA migrates from the nucleus to mitochondria seems at least 100,000 times less [35]. The frequency of a gene escape from the chloroplast to the nucleus is at least 1 per 5 million cells in tobacco [36]. A more puzzling question is how organellar sequences could be remodeled following their integration in the nuclear chromosomes, so as to contain domains allowing their addressing back to the mitochondrion, the chloroplast, and sometimes both. Specific N-terminal sequences, called the mitochondrial and chloroplast transit peptides (Mtp and Ctp) thus allow entry into the mitochondrion via a large translocon complex, at the outer and inner envelope membrane (called TOM/TIM) and in the chloroplast via another translocon complex (called TOC/TIC). Following entry inside the organelles, the Mtp and Ctp are cleaved by specific peptidases in the mitochondrial matrix and chloroplast stroma, respectively. A convincing model has been introduced recently, proposing that these addressing N-terminal sequences derived from a specific property of the ancestors of the organelles, i.e., a resistance strategy to antimicrobial peptides that consisted in their rapid internalization and proteolytic disposal by microbial peptidases [27]. 3.4 Inheritance of Non-α-Proteobacterial and Noncyanobacterial Genes by Horizontal Gene Transfers

The description we have given on the current reconstruction of the evolution of early eukaryotes seems simple. This picture might be satisfying if only genes tracing back to FECA (initial pool of Archaea, Bacteria, and unique Eukarya genes), α-proteobacterial and cyanobacterial genes were detected. The reduction of mitochondrial and chloroplast genomes should therefore coincide with an increase of nuclear-encoded genes with α-proteobacterial and cyanobacterial signatures. However, this simple scheme is far from being satisfactory. The major issue is that a large number of genes have other signatures, i.e., non-α-proteobacterial genes and proteins in mitochondria [20] and noncyanobacterial genes and

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proteins in chloroplasts [4, 24, 37]. This presence of mostly bacterial genes, and sometimes genes with no prokaryotic signatures, highlight the importance of horizontal gene transfers (HGT) from other Eukarya and Bacteria [13]. In Fig. 4, HGT is considered at all stages of Eukarya evolution, and is still ongoing. Some bacterial genes could be incorporated via Bacteria-to-Archaea HGT before and after FECA emergence. The acquisition of the mitochondrion could involve Bacteria-to-α-proteobacterium HGT, followed by the escape of some of the mitochondrial genes to the nucleus. Likewise, the acquisition of the chloroplast could involve Bacteriato-cyanobacteria HGT, followed by the escape of some genes to the nucleus. Some lateral gene transfers between both organelles could then have occurred. 3.5 Could an Ancestral Chlamydia Have Helped Primary Chloroplast Emergence? The Me´ nage-a`-trois Hypothesis (MATH)

A novel hypothesis has been recently proposed based on the intriguing detection of genes having Chlamydial signatures in photosynthetic eukaryotes [21, 38], often harboring a plastid-targeting signal [39]. Up to 55 genes are proposed to be transferred from Chlamydiae to primary photosynthetic eukaryotes [40–43]. Chlamydia are obligate intracellular bacteria, including pathogens of animals, and found in unicellular organisms such as amoebae [21]. The primary endosymbiosis at the origin of chloroplasts might have therefore involved an ancestral cyanobacterium and an ancestral Chlamydia and be conserved, thanks to Chlamydia-toeukaryote LGT [40, 41, 44]. The presence of the bacterial pathogen would have been critical in transmitting genes allowing the long-term residence of the cyanobacterial symbiont. Two models are currently considered (Fig. 5). In the first model, cyanobacteria and Chlamydia entered the host cell simultaneously but in distinct phagosomal vacuoles (Fig. 5, left). In the second model, both cyanobacteria and Chlamydia occupied the same phagocytic vacuole (Fig. 5, right). The cooccurrence of a cyanobacteria and Chlamydia is then supposed to have led to the development of a protochloroplast, combining genes obtained via intense LGT, making the basis for modern primary plastids.

3.6 Three Lineages Deriving from Chloroplast Primary Endosymbiosis: Glaucocystophytes, Chlorophytes, and Rhodophytes

Following the primary endosymbiosis, which has led to the emergence of the chloroplast, three major lineages have been defined, based on pigments in photosynthetic machineries [45] (Fig. 6a). • The green lineage of primary endosymbionts (Viridiplantae), in which chlorophyll a is associated with chlorophyll b, contains the “green algae” (Chlorophyta), such as Chlamydomonas reinhardtii, and the so-called plants (Streptophyta), such as Arabidopsis thaliana; • The red lineage of primary endosymbionts, in which chlorophyll a is associated with phycobiline, contains the “red algae” (Rhodophyta), such as Cyanidioschyzon merolae.

At the Origin of Plastids

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Acquisition of the chloroplast following the MATH model

Ancestral cyanobacteria

Ancestral cyanobacteria 2

Ancestral Chlamydia

1 Ancestral Chlamydia

Loss of Chlamydia cells and LGT facilitating chloroplast stable residence

Fig. 5 Acquisition of the primary chloroplast following the Me´nage-a`-Trois Hypothesis (MATH). In this scheme, the acquisition of the ancestral cynaobacteria coincides with the presence of parasitic Chlamydia, either in distinct (1) or identical (2) phagocytic vacuoles. The presence of Chlamydia cells provides a genetic environment adapted to the residence of a bacterium within a eukaryote. Following HGT, Chlamydia genes are proposed to have facilitated the cyanobacteria-to-chloroplast transition

• A nongreen, nonred (sometimes called “blue”) lineage of primary endosymbionts, in which chlorophyll a is associated with phycocyanin and allophycocyanin, is a small group of unicellular organisms (Glaucocystophytes), such as Cyanophora paradoxa, in which the chloroplast still contains a peptidoglycan cell wall.

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Fig. 6 Chloroplast-containing and chromatophore-containing eukaryotes. (a) The organisms containing a primary chloroplasts belong the Archaeplastida kigdom. The primary endosymbiosis has occurred more than 1 billion years ago. Three lineages have been defined: the green lineage comprising Green Algae and Plants (also called Viridiplantae), the red lineage comprising Red algae, and the nongreen, nonred lineage comprising Glaucophyta, such as Cyanophora species. Glaucophyta contain a chloroplast with peptidoglycans, which origin might be distinct from those of the ancestral cyanobacteria. (b) An independent primary endosymbiosis has occurred 100–60 million years ago leading to the emergence of another type of photosynthetic organelle, the chromatophore, in Paullinellidae. The organelle contains peptidoglycans, like in cyanobacteria. Pept, peptidoglycans

4 A Second and Independent Cyanobacterial Endosymbiosis in Rhizaria at the Origin of the Chromatophore The study of Paulinella chromatophora, a unicellular photosynthetic eukaryote, has led to the discovery of a second independent primary endosymbiosis leading to the integration of a cyanobacterium into a Rhizarian amoeba, 100–60 million years ago [46–48] (Fig. 4b). The organelle is not called a chloroplast, but a chromatophore. Phylogenetic analyses have shown that a quarter of proteinencoding genes could be found in modern free-living Synechocystis species [36]. Just like in chloroplasts, a genome reduction of the chromatophore is ongoing and genes have escaped to the nucleus and encode proteins imported back into the organelle (almost 450 nuclear-encoded proteins targeted to the chromatophore) [48–52]. Like mitochondria and chloroplasts, the chromatophore

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function relies on a large number of bacterial genes originating from HGT [53]. This discovery is therefore fascinating. First, this endosymbiotic event shows that although it is rare, the emergence of a semi-autonomous organelle can still occur. Second, features observed in the α-proteobacterium-to-mitochondrion and cyanobacteria-to-chloroplast transition are also observed. Third, we are witnessing the emergence of a novel eukaryotic organization and the evolution of chromatophore-containing species might lead to a very rich and complex biodiversity in terms of molecular, structural, and functional diversity that we cannot extrapolate today.

5

Conclusion In this chapter, the current understanding of thylakoid emergence in cyanobacteria and the acquisition of chloroplasts by primary endosymbiosis has been summarized, leaving some frustrating unresolved questions, which might actually stay unresolved unless novel data are made available. The presented model for the emergence of thylakoid membranes points to the actual function of companion proteins, such as Vipp1, and to the overlooked role of HexII $ Lm phase transitions promoted by such lipids as SQDG, in the biogenesis of membrane compartments. The emergence of semi-autonomous organelles seems to share general features, i.e., an acquisition of a bacterium/cyanobacteria likely via a phagocytic membrane, a genome reduction coinciding with an escape of genes from the organelle to the nucleus, and, finally, the appearance of an active system translocating nuclear-encoded proteins back to the organelles. An intense mobilization of foreign genes of bacterial origin, via HGT, plays a critical role. Some third partners, like Chlamydia, might even have facilitated the transition from cyanobacteria to chloroplasts. Interestingly, although the evolution of mitochondria can lead to the complete loss of DNA, occurring independently in multiple lineages, forming mitochondria-derived hydrogenosomes and mitosomes, no such phenomenon is currently demonstrated for chloroplasts. Altogether, this chapter illustrates also the diversity of organelle forms and functions that can derive from primary endosymbiosis. An increase in complexity has arisen from multiple events of secondary endosymbioses between nonphotosynthetic and photosynthetic eukaryotes and has raised superbly complicated cell structures. The understanding of the evolution, structure, and function of these secondary or complex plastids is far less advanced compared to primary plastids, representing therefore a major goal and challenge for future research.

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Acknowledgments This work was supported by the French National Research Agency (GRAL Labex ANR-10-LABEX-04, EUR CBS ANR-17-EURE0003, Alpalga ANR-20-CE02-0020, DIM ANR-21-CE02-0021). References 1. Raven JA, Sanchez-Baracaldo P (2021) Gloeobacter and the implications of a freshwater origin of Cyanobacteria. Phycologia 60(5): 4 0 2 – 4 1 8 . h t t p s : // d o i . o r g / 1 0 . 1 0 8 0 / 00318884.2021.1881729 2. Mares J, Strunecky O, Bucinska L et al (2019) Evolutionary patterns of thylakoid architecture in cyanobacteria. Front Microbiol 10:277. https://doi.org/10.3389/fmicb.2019.00277 3. Gueguen N, Mare´chal E (2022) Origin of cyanobacterial thylakoids via a non-vesicular glycolipid phase transition and impact on the great oxygenation event. J Exp Bot 73(9): 2721–2734. https://doi.org/10.1093/jxb/ erab429 4. Petroutsos D, Amiar S, Abida H et al (2014) Evolution of galactoglycerolipid biosynthetic pathways--from cyanobacteria to primary plastids and from primary to secondary plastids. Prog Lipid Res 54:68–85. https://doi.org/ 10.1016/j.plipres.2014.02.001 5. Marrink SJ, Mark AE (2004) Molecular view of hexagonal phase formation in phospholipid membranes. Biophys J 87(6):3894–3900. https://doi.org/10.1529/biophysj.104. 048710 6. Rexroth S, Mullineaux CW, Ellinger D et al (2011) The plasma membrane of the cyanobacterium Gloeobacter violaceus contains segregated bioenergetic domains. Plant Cell 23(6):2379–2390. https://doi.org/10.1105/ tpc.111.085779 7. Huokko T, Ni T, Dykes GF et al (2021) Probing the biogenesis pathway and dynamics of thylakoid membranes. Nat Commun 12(1): 3475. https://doi.org/10.1038/s41467021-23680-1 8. Koskela M, Annila A (2012) Looking for the last universal common ancestor (LUCA). Genes (Basel) 3(1):81–87. https://doi.org/ 10.3390/genes3010081 9. Forterre P (2015) The universal tree of life: an update. Front Microbiol 6:717. https://doi. org/10.3389/fmicb.2015.00717 10. Cantine MD, Fournier GP (2017) Environmental adaptation from the origin of life to the last universal common ancestor. Orig Life

Evol Biosph 48:35. https://doi.org/10.1007/ s11084-017-9542-5 11. van der Giezen M, Tovar J, Clark CG (2005) Mitochondrion-derived organelles in protists and fungi. Int Rev Cytol 244:175–225. https://doi.org/10.1016/S0074-7696(05) 44005-X 12. Gribaldo S, Poole AM, Daubin V et al (2010) The origin of eukaryotes and their relationship with the Archaea: are we at a phylogenomic impasse? Nat Rev Microbiol 8(10):743–752. https://doi.org/10.1038/nrmicro2426 13. Eme L, Spang A, Lombard J, Stairs CW, Ettema TJG (2017) Archaea and the origin of eukaryotes. Nat Rev Microbiol 15:711–723. https://doi.org/10.1038/nrmicro.2017.154 14. Nobs SJ, MacLeod FI, Wong HL et al (2022) Eukarya the chimera: eukaryotes, a secondary innovation of the two domains of life? Trends Microbiol 30(5):421–431. https://doi.org/ 10.1016/j.tim.2021.11.003 15. Raval PK, Garg SG, Gould SB (2022) Endosymbiotic selective pressure at the origin of eukaryotic cell biology. elife 11:e81033. https://doi.org/10.7554/eLife.81033 16. Mereschkowsky C (1905) Ober Natur and Ursprung der Chromatophoren im Pflanzenreiche. Biol Zentralbl 25:593–604 17. Margulis L (1970) Origin of eukaryotic cells. Yale University Press, New Haven 18. Imachi H, Nobu MK, Nakahara N et al (2022) Isolation of an archaeon at the prokaryoteeukaryote interface. Nature 577(7791): 519–525. https://doi.org/10.1038/s41586019-1916-6 19. Poole AM, Gribaldo S (2014) Eukaryotic origins: how and when was the mitochondrion acquired? Cold Spring Harb Perspect Biol 6(12):a015990. https://doi.org/10.1101/ cshperspect.a015990 20. Lopez Alonso D, Garcia-Maroto F, RodriguezRuiz J et al (2003) Evolutiuon of membranebound fatty acid desaturases. Biochem Syst Ecol 31:1111–1124 21. Jensen PE, Leister D (2014) Chloroplast evolution, structure and functions. F1000Prime Rep 6:40. https://doi.org/10.12703/P6-40

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34. McFadden GI (1999) Endosymbiosis and evolution of the plant cell. Curr Opin Plant Biol 2(6):513–519 35. Thorsness PE, Fox TD (1990) Escape of DNA from mitochondria to the nucleus in Saccharomyces cerevisiae. Nature 346(6282):376–379. https://doi.org/10.1038/346376a0 36. Stegemann S, Hartmann S, Ruf S et al (2003) High-frequency gene transfer from the chloroplast genome to the nucleus. Proc Natl Acad Sci USA 100(15):8828–8833. https://doi. org/10.1073/pnas.1430924100 37. Reyes-Prieto A, Moustafa A (2012) Plastidlocalized amino acid biosynthetic pathways of Plantae are predominantly composed of non-cyanobacterial enzymes. Sci Rep 2:955. https://doi.org/10.1038/srep00955 38. Horn M, Collingro A, Schmitz-Esser S et al (2004) Illuminating the evolutionary history of chlamydiae. Science 304(5671):728–730. https://doi.org/10.1126/science.1096330 39. Brinkman FS, Blanchard JL, Cherkasov A et al (2002) Evidence that plant-like genes in Chlamydia species reflect an ancestral relationship between Chlamydiaceae, cyanobacteria, and the chloroplast. Genome Res 12(8): 1159–1167. https://doi.org/10.1101/gr. 341802 40. Ball SG, Subtil A, Bhattacharya D et al (2013) Metabolic effectors secreted by bacterial pathogens: essential facilitators of plastid endosymbiosis? Plant Cell 25(1):7–21. https://doi.org/ 10.1105/tpc.112.101329 41. Huang J, Gogarten JP (2007) Did an ancient chlamydial endosymbiosis facilitate the establishment of primary plastids? Genome Biol 8(6):R99. https://doi.org/10.1186/gb2007-8-6-r99 42. Huang J, Gogarten JP (2008) Concerted gene recruitment in early plant evolution. Genome Biol 9(7):R109. https://doi.org/10.1186/ gb-2008-9-7-r109 43. Moustafa A, Reyes-Prieto A, Bhattacharya D (2008) Chlamydiae has contributed at least 55 genes to Plantae with predominantly plastid functions. PLoS One 3(5):e2205. https://doi. org/10.1371/journal.pone.0002205 44. Cenci U, Bhattacharya D, Weber AP et al (2017) Biotic host-pathogen interactions as major drivers of plastid endosymbiosis. Trends Plant Sci 22(4):316–328. https://doi.org/10. 1016/j.tplants.2016.12.007 45. Archibald JM, Keeling PJ (2002) Recycled plastids: a ‘green movement’ in eukaryotic evolution. Trends Genet 18(11):577–584

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50. Mackiewicz P, Bodyl A, Gagat P (2012) Protein import into the photosynthetic organelles of Paulinella chromatophora and its implications for primary plastid endosymbiosis. Symbiosis 58(1–3):99–107. https://doi.org/10. 1007/s13199-012-0202-2 51. Gagat P, Bodyl A, Mackiewicz P (2013) How protein targeting to primary plastids via the endomembrane system could have evolved? A new hypothesis based on phylogenetic studies. Biol Direct 8:18. https://doi.org/10.1186/ 1745-6150-8-18 52. Bodyl A, Mackiewicz P, Gagat P (2012) Organelle evolution: Paulinella breaks a paradigm. Curr Biol 22(9):R304–R306. https:// doi.org/10.1016/j.cub.2012.03.020 53. Nowack EC, Price DC, Bhattacharya D et al (2016) Gene transfers from diverse bacteria compensate for reductive genome evolution in the chromatophore of Paulinella chromatophora. Proc Natl Acad Sci U S A 113(43): 12214–12219. https://doi.org/10.1073/ pnas.1608016113

Chapter 2 Complex Endosymbioses I: From Primary to Complex Plastids, Serial Endosymbiotic Events Zolta´n Fu¨ssy and Miroslav Obornı´k Abstract A considerable part of the diversity of eukaryotic phototrophs consists of algae with plastids that evolved from endosymbioses between two eukaryotes. These complex plastids are characterized by a high number of envelope membranes (more than two) and some of them contain a residual nucleus of the endosymbiotic alga called a nucleomorph. Complex plastid-bearing algae are thus chimeric cell assemblies, eukaryotic symbionts living in a eukaryotic host. In contrast, the primary plastids of the Archaeplastida (plants, green algae, red algae, and glaucophytes) possibly evolved from a single endosymbiosis with a cyanobacterium and are surrounded by two membranes. Complex plastids have been acquired several times by unrelated groups of eukaryotic heterotrophic hosts, suggesting that complex plastids are somewhat easier to obtain than primary plastids. Evidence suggests that complex plastids arose twice independently in the green lineage (euglenophytes and chlorarachniophytes) through secondary endosymbiosis, and four times in the red lineage, first through secondary endosymbiosis in cryptophytes, then by higher-order events in stramenopiles, alveolates, and haptophytes. Engulfment of primary and complex plastid-containing algae by eukaryotic hosts (secondary, tertiary, and higher-order endosymbioses) is also responsible for numerous plastid replacements in dinoflagellates. Plastid endosymbiosis is accompanied by massive gene transfer from the endosymbiont to the host nucleus and cell adaptation of both endosymbiotic partners, which is related to the trophic switch to phototrophy and loss of autonomy of the endosymbiont. Such a process is essential for the metabolic integration and division control of the endosymbiont in the host. Although photosynthesis is the main advantage of acquiring plastids, loss of photosynthesis often occurs in algae with complex plastids. This chapter summarizes the essential knowledge of the acquisition, evolution, and function of complex plastids. Key words Complex endosymbiosis, Plastid replacement, Reductive evolution

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Introduction Photosynthesis plays a vital role in sustaining life on Earth by harnessing the energy of sunlight to create chemical bonds, thereby serving as a significant driver of primary productivity for all life. Light-harvesting complexes couple the energy transfer of photons to produce NADPH and a proton gradient across membranes, yielding energy needed to incorporate CO2 into organic

Eric Mare´chal (ed.), Plastids: Methods and Protocols, Methods in Molecular Biology, vol. 2776, https://doi.org/10.1007/978-1-0716-3726-5_2, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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compounds. The inventors of this complex process and molecular machinery, cyanobacteria, have not only colonized aquatic environments but also have formed crucial bonds with eukaryotes through endosymbiosis and thus expanded into every conceivable niche on Earth with sufficient light [1]. As endosymbiotic organelles, plastids are found within their eukaryotic hosts in environments ranging from oceans to land and are of the most numerous and diverse biological entities. The history of plastids dates to the first association between a unicellular heterotrophic eukaryote, an early ancestor of Archaeplastida, and a phototrophic cyanobacterium, which we refer to as prokaryote-to-eukaryote or primary endosymbiosis (Fig. 1) [2]. Streptophytes (land plants), chlorophytes (green algae), rhodophytes (red algae), and glaucophytes, together constituting Archaeplastida, harbor primary plastids that are thought to have evolved from a domesticated cyanobacterium in a single endosymbiotic event. Another exciting case of primary endosymbiosis was found in rhizarian amoebae of the genus Paulinella [3], which acquired their endosymbiont cyanobacterium more recently, allowing the examination of the early phases of this process. However, multiple other lineages of phototrophs are scattered throughout the eukaryote tree of life and represent a significant portion of their diversity (Fig. 2). These algal lineages are essential for global food chains because diatoms, dinoflagellates, or haptophytes are responsible for most of the primary production in the ocean. In fact, the primary production of diatoms is comparable to that of all terrestrial rainforests combined [4]. In contrast to Archaeplastida and Paulinella, plastids in these lineages did not arise by primary endosymbiosis with a cyanobacterium. Instead, they were acquired “horizontally,” by engulfment of a photosynthetic eukaryote (an alga), which is called a eukaryote-to-eukaryote or complex endosymbiotic event (Fig. 2). Some complex algae possess a residual nucleus of the engulfed endosymbiont in their plastids; these so-called nucleomorphs are the most convincing evidence for the eukaryotic origin of complex plastids. Classification of plastid endosymbiosis events reflects their historical order, and so when we talk about secondary endosymbiosis, we mean that the endosymbiont is a primary alga. Higher-order endosymbioses (tertiary, quaternary, etc.) seem to be just as common though and result from more entangled interactions involving eukaryotic hosts and complex algal endosymbionts, much like Matryoshka dolls (Fig. 1). The origins of complex plastids in some lineages are still rather hotly debated. These evolutionary events occurred hundreds of millions of years ago, so the molecular phylogenetic signals have largely eroded, and sometimes we can only tell for sure that the endosymbionts are rhodophyte-derived. We, therefore, prefer to use the term “complex plastids” not only

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Fig. 1 Evolution of plastid envelopes. (a) Phases of the endosymbiont reduction over the course of time. 1, Feeding on algal prey; 2, gradual reduction of the endosymbiont structures and gene transfer to host nucleus resulting in a reduced endosymbiont nucleus, the nucleomorph; 3, progressing dependence on host factors, nucleomorph lost; 4, loss of the endosymbiont-derived membrane. (b) Envelope structures under various endosymbiosis scenarios. As apparent from the scheme, different-order complex endosymbioses may result in plastids with the same envelope arrangement. (c) More complex envelopes require additional proteintranslocating machinery, reflected by altered protein targeting presequences. Color code of protein domains: green, plastid transit peptide; orange, signal peptide; red, transmembrane anchor domain; gray, mature protein. TOC/TIC, translocons of the outer/inner chloroplast membrane; SEC, signal peptide translocon at the endoplasmic reticulum membrane; SELMA, symbiont-derived translocon (see text)

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when discussing general principles of endosymbiosis but also when considering the unresolved sequence of plastid acquisitions in different rhodophyte-derived lineages (Fig. 2) [5, 6]. To understand how plastids establish as organelles, it is also important to learn what happens to plastids later after an endosymbiotic event. Plastids play a role in cell biochemistry that extends beyond photosynthesis. They play key roles in the regulation of carbon and energy balance, nitrogen and sulfur assimilation, and the biosynthesis of essential compounds such as vitamins, tetrapyrroles, fatty acids, and isoprenoids. It is, therefore, not surprising that many lineages retain plastids even when they lose photosynthesis. In this chapter, we summarize the essentials about the evolution of complex plastids and their role in organisms that possess them.

2

Distribution of Plastids Among Eukaryotes Currently, six or seven species-rich “supergroups” of eukaryotes are recognized (and a handful of incertae sedis, uncertainly placed lineages; Fig. 2, [7]). Obazoa (Apusomonada, Breviates, and Opisthokonta) and Amoebozoa likely never had permanent plastids, although endosymbionts of other origins occasionally occur in different lineages (e.g., [8, 9]). In Archaeplastida, heterotrophic Picozoa form a sister group to rhodophytes and thus appear nested among phototrophic lineages [10]. This suggests either an early loss of plastids in Picozoa or multiple primary endosymbioses in the Archaeplastida, most likely in the ancestor of rhodophytes and rhodelphids and in the ancestor glaucophytes, chlorophytes, and streptophytes (Fig. 2). Four and two lineages contain complex rhodophyte- and chlorophyte-derived plastids, respectively. In early works, the “red lineage” and the “green lineage” were considered monophyletic [11] to fit the most parsimonious evolutionary scenario of two endosymbioses, sometimes referred to as early plastid acquisition. However, accumulating evidence has consistently rejected these notions, and it has become clear that these six lineages most likely arose independently, through “late acquisition” [12–15]. First of all, all complex plastid-bearing lineages represent crown lineages in their respective clades, unambiguously surrounded by heterotrophic, plastid-less relatives: ciliates basal to myzozoans (dinoflagellates and apicomplexans), heterotrophic stramenopiles (i.e., Oomycota, Labyrinthulea, Opalozoa) basal to ochrophytes, centrohelids to haptophytes, katablepharids to cryptophytes, discobids to euglenophytes, and rhizarians to chlorarachniophytes. Moreover, host phylogenomic analyses suggest that these ancestral heterotrophic lineages branch deeply in the evolution of eukaryotes and predate the diversification of red and green algae

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Fig. 2 Eukaryotic tree of life with an accent on the diversity of algae. Major eukaryotic groups are shown around the outermost circle, in gray. Inner circles mark lower-rank groups of organisms, nonphotosynthetic clades boxed in shades of gray, the “green lineage” boxed in shades of green, and the “red lineage” boxed in shades of red and brown. Note that Cryptista and Haptista do not robustly associate with currently recognized major eukaryotic groups (*). The cladogram shows a schematic relationship between taxa, with the course of evolution from the center to the margin, as marked by the black arrow. Red- and green-colored nodes and branches denote the red- and green-algal plastid descendants, respectively, light-green indicates the primary endosymbiont of Paulinella and Archaeplastida before the divergence of red algae, glaucophytes, and green algae. Letters mark levels of particular symbiotic events: P – primary, S – secondary, C – complex (unclear higher-order), K plus a dashed branch line – kleptoplasty. Narrow rectangles show losses of photosynthesis (white) or entire plastids (black). Taxa abbreviations: chl. – chlorarachniophytes, For. – Foraminifera, Gl. – glaucophytes, H. – Hematodinium, Hapt – Haptista, Paul. – Paulinella, * – incertae sedis, uncertain evolutionary position

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[16, 17]. Lastly, the plastid genomes of euglenophytes and chlorarachniophytes show affinity with different lineages of chlorophytes, corroborating their independent origins [12]. Altogether, available phylogenetic data suggest six complex endosymbiotic events (Fig. 2, representative algae shown in Fig. 3) [12]. In agreement with their phylogeny, complex plastids substantially differ lineage-to-lineage by their morphology and function. Within each lineage, the envelope ultrastructure usually remains conserved, supporting the notion that endosymbiotic events define monophyletic clades. While primary plastids of Archaeplastida and Paulinella have two surrounding membranes, dinoflagellates (except those that have undergone serial endosymbiosis) and euglenophytes have plastids with three membranes, while other complex algae have plastids with four, exceptionally five (dinotoms) membranes (Fig. 1). Cryptophytes and chlorarachniophytes both possess a distinctive residual nucleomorph in their plastids, with three tiny chromosomes [18]. This structure localizes in the so-called periplastidial space, which is squeezed between the two outermost and the two inner plastid membranes and, as expected, corresponds to the cytosol of the endosymbiont (Fig. 1). The ultrastructure of plastids is thus thought to directly reflect the physical mechanisms of primary or complex endosymbiosis. The nucleomorphs remained in plastids where the host genome could not provide all the necessary factors for their function, and the endosymbiont had to step in. The additional membranes evolved from the host endomembrane system, which engulfed the organelle, and/or the cytoplasmic membrane of the symbiont [2]. Interestingly, none of the higher-order plastids are enclosed by more than five membranes (Fig. 1) [19] suggesting that some of the membranes must disappear during the engulfment [20]. These nuances all document that there have been multiple successful approaches to endosymbiosis. While plastids in various lineages clearly differ in important characteristics, there are undeniable and conspicuous similarities between them, too. For instance, all rhodophyte-derived complex algae utilize chlorophylls a and c, the latter apparently being their evolutionary innovation, because rhodophytes only use chlorophyll a, and the green lineage (including chlorarachniophytes and euglenophytes) use chlorophylls a and b. How do we resolve this discrepancy between host and plastid phylogenies? The apparent similarity of rhodophyte-derived complex plastids most likely results from horizontal “grafting” of established organelles from one lineage to another via higher-order endosymbioses (e.g., [6, 14, 21, 22]). Under this scenario, chlorophyll c evolved in a secondary plastid-containing alga, most likely cryptophytes, which later formed direct or indirect higher-order relationships with the ancestors of haptophytes, ochrophytes, and myzozoans, although the order remains a conundrum. One hypothesis, based on the number of genes that were transferred to the host nucleus with

Fig. 3 Examples of algae with complex plastids. (a) Diatom Phaeodactylum tricornutum, (b) peridinin dinoflagellate Amphidinium carterae, (c) dinoflagellate with a diatom endosymbiont, also called a dinotom, Kryptoperidinium foliaceum, (d) autofluorescence of the diatom plastids in the dinoflagellate K. foliaceum (plastid – magenta; nucleus – blue), (e) excavate alga Euglena gracilis, (f) excavate colourless (osmotrophic) alga Euglena longa, (g) alveolate alga Chromera velia, (h) alveolate alga Vitrella brassicaformis. Image courtesy of K. Jiroutova´ (A), Z. Fu¨ssy (B, E, F), J. Cihla´rˇ (C), J. Richtova´ (D), and D. Modry´ (G, H)

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the plastid (see below), posits that ochrophytes possess tertiary plastids and were taken up as quaternary plastids by myzozoans and haptophytes, although haptophytes could have also obtained their plastid via tertiary endosymbiosis with cryptophytes [23–25]. Another controversy arose when the plastid genome and proteome of the apicomonad Vitrella brassicaformis (Fig. 3h) revealed phylogenetic relationships with those of eustigmatophytes, a subgroup of ochrophytes [26–28]. According to some data, apicomonad algae [20, 26, 29, 30] and related sporozoan parasites (Plasmodium, Toxoplasma) may have obtained their plastids via higher-order endosymbiosis with an ochrophyte [27, 28, 30, 31], independently of dinoflagellates. Another indication of this is the absence of chlorophyll c in both chromerids (phototrophic apicomonads) and eustigmatophytes [30]. Some researchers suggested that higher-order endosymbiosis with an ochrophyte alga led to the origin of the entire Myzozoa (including dinoflagellates) [14]. However, the differences between the plastids of dinoflagellates and chromerids/sporozoans, particularly the ultrastructure of the plastid envelopes, pigmentation, and organization of the genome, suggest a separate origin of the endosymbionts in these sister lineages [13]. This would not seem surprising, as dinoflagellates showcase the possibilities of endosymbiotic relationships [2, 19, 32]. Dinoflagellates, generally exhibiting great ecological plasticity, repeatedly underwent a spectrum of events ranging from retention of rhodophyte-derived plastid in peridinin-pigmented species, through kleptoplasty (theft of plastid from prey) and serial endosymbiosis (plastid replacement), to losses of photosynthesis or the plastid altogether in parasitic and predatory species (Fig. 2). The best-known examples of serial endosymbiosis are Lepidodinium, “MGD,” and “TGD” (undescribed dinoflagellates), with green algal endosymbionts, dinotoms maintaining diatom endosymbionts (Fig. 3b), and Kareniaceae with haptophyte endosymbionts (Fig. 2) [2, 32–34]. Importantly, these evolutionary events in dinoflagellates occur on smaller time scales and allow for “realtime” examination of the processes involved. Dinophysis is the best-known kleptoplastic lineage, i.e., employing a strategy to repeatedly steal plastids from algae [35], but frequent plastid promiscuity in dinotoms, some of which have been found frozen in the state of kleptoplasty, seems promising for studying early plastid evolution [36].

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Integration of Plastids Plastid evolution is multifaceted but involves two main processes. The first component, critical for the transformation into fully integrated organelles, is the establishment of host control over the endosymbiont’s biochemistry (via channeling) and its division (via anterograde signaling) [37–39]. To support the cell biological

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and metabolic processes of the newborn organelle, some level of evolutionary innovation is necessary, represented by new or repurposed proteins and complexes. These allow proper targeting (translocation) of proteins encoded in the host nucleus across the plastid envelope [40, 41], compound exchange [38, 42], and control of organelle division [43]. Along with the establishment of communication mechanisms, genetic material is massively transferred from the endosymbiont to the host nucleus or lost. Reductive evolution is therefore the second important component of symbiogenesis [44, 45]. As a result, plastids lose most of their genome complexity (and autonomy) compared to free-living cyanobacteria. Analogously during complex endosymbioses, the nuclear genome of the endosymbiont, which ranges from 16 to 105 Mb in free-living red algae [46], is significantly reduced to the form of a nucleomorph or, more frequently, lost altogether. At the end of the process, plastid proteomes of primary algae (and plants) consist of about 1000–1500 proteins (perhaps up to 3000 according to [47]), with only 87 proteins encoded by the plastid genome (in A. thaliana) [48]. Proteome richness, as well as actual coding capacity, is quite similar in complex plastids, although their genomes generally contain fewer genes [18, 24, 34, 49]. Therefore, the mechanisms of instating host control and organellar reduction work hand in hand to allow this change of command. Almost all the hundreds of nuclear-encoded proteins targeted to plastids have a targeting presequence that is recognized by the translocation machinery that guides the protein across the plastid envelopes (Fig. 1) [50, 51]. Genes transferred from the plastid genome to the host nucleus, or any other new proteins directed to the compartment, must acquire this targeting presequence to (re)gain their plastid function. The number of membranes in plastid envelopes greatly affects the complexity of the machinery by which nuclear-encoded proteins are imported into the organelle. Proteins with a so-called chloroplast transit peptide presequence can cross the two-membrane envelopes of primary plastids. Proteins destined for complex plastids must cross additional membranes and are therefore equipped with an additional domain, the signal peptide, just upstream of the transit peptide [50]. In terms of intracellular connectivity (topologically), therefore, primary plastids are localized in the cytosol of Archaeplastida because they do not have additional membrane barriers that they did not inherit from the two-membrane cyanobacteria (Fig. 1). In comparison, complex plastids are accessible only through the host endomembrane system from which the outer membranes of the envelope originate. Indeed, the pathways of protein import in primary and complex algae reflect this fundamental ultrastructural difference (Fig. 1) [50]. The presequences (transit peptides) of proteins targeted to primary plastid are recognized directly by the translocon complexes at the outer and inner membranes of the

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envelope (TOC and TIC). To cross the additional membrane(s) of the complex plastids, the proteins must enter the endomembrane system via the endoplasmic reticulum (ER) and therefore require the signal peptide (SP) in their presequence. After entry into ER and cleavage of SP, the fate of proteins differs in three- and fourmembrane plastids (Fig. 1). In three-membrane plastids, the cleavage of SP exposes the transit peptide, which then directs the entry of the protein across the two inner membranes via the TOC/TIC. Remarkably, proteins imported into three-membrane plastids typically have a hydrophobic domain, a membrane anchor, downstream of the transit peptide. This possibly allows vesicular transport between the ER and the plastid outermost membrane, which are not physically connected [52, 53]. Complex plastids with a fourmembrane envelope require an additional protein translocation system to pass through the second outermost membrane. This translocon originates from the endosymbiont ERAD (ER -Associated Protein Degradation), which originally exported misfolded proteins from the ER. After redirection to the second membrane (thought to be homologous to the endomembrane system or cytoplasmic membrane of the endosymbiont), this complex began translocating proteins further across. Passing this transtermed SELMA (symbiont-specific ERAD-like locator, machinery), plastid proteins are now facing TOC/TIC complexes in the innermost two membranes (Fig. 1) [54]. Five-membrane plastids are thought to be inaccessible to host-encoded proteins, which is why they have a nucleomorph under the outermost membrane that encodes all necessary plastid proteins that are translocated as in four-membrane plastids [55]. The inability to import dinoflagellate host proteins into these endosymbionts is likely the reason for their unprecedented conservation—they still contain mitochondria and an almost unreduced nucleus. The need for hundreds of proteins to obtain a correct targeting signal is considered one of the main reasons why primary and secondary endosymbioses are so rare [56]. Moreover, molecular machines such as photosystems are unlikely to evolve de novo in eukaryotes. Not only are they highly sophisticated in terms of subunit composition, but also bind strictly to specific (“genetic” or inherited) membranes that do not appear de novo. Genetic membranes, such as plastid envelope membranes, often have specific lipid and protein compositions and derive only semi-conservatively from preexisting membranes, much like a new DNA strand derives from the template strand [56]. Thus, the evolution and spread of photosynthesis in eukaryotes was mediated exclusively by endosymbiosis. For metabolic integration, plastids use a series of transporters to connect to the cytosolic pool of compounds [38]. Triose phosphate/phosphate translocators (TPTs) were supposedly the pioneers of the connection between the host and the endosymbiont,

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as they allow the exchange of three-carbon sugar intermediates synthesized during CO2 fixation (Calvin-Benson cycle). Other compounds transported include glucose-6-phosphate, xylulose-5phosphate, phosphoenolpyruvate, and glutathione, although the distribution of their transporters is not necessarily universal among complex plastids [57]. Members of the mitochondrial carrier family facilitate the transport of substrates such as folates, S-adenosylmethionine, NAD, ADP-glucose, or adenosine nucleotides; ATP:ADP antiporters ensure the exchange of ATP and thus the maintenance of the physiological ATP/ADP ratio in the plastid [57]. Dicarboxylate transporters play a role in nitrogen assimilation by allowing the circulation of 2-oxoglutarate and glutamate into and out of the plastid, respectively. 2-oxoglutarate is the acceptor for ammonia in the glutamate synthase reaction, whereas glutamate then serves as a nitrogen donor for the biosynthesis of nitrogen-containing compounds. The presence and function of other transporters, such as the pyruvate carrier, are unknown in most complex algal species [57]. Once established, plastids play a dominant role in algal biochemistry. With four major multi-subunit protein complexes and about 80 proteins involved [58], photosynthesis is the most prominent process of plastids. These proteins are required for the assembly, function, and regulation of light-harvesting antennae, photosystems, and electron transfer factors, and enable the production of ATP and the reducing agent NADPH as co-substrates for carbon fixation. In addition to photosynthesis, plastids function as biochemical factories, synthesizing vitamins, polysaccharides, amino acids, fatty acids, isoprenoids, tetrapyrroles, and Fe-S clusters. Most of these compounds are directly required for photosynthesis, but in many organisms, plastid synthesis supplies the entire cell. As discussed below, plastids tend to take over metabolic functions of other cellular compartments to achieve a streamlined and light-regulated biochemistry. Plastids also cooperate with mitochondria and the cytosol to balance metabolic and energy flows during the day-night cycle and under nutrition limitation [59, 60]. However, plastids from distinct lineages differ in their metabolic capabilities. For example, unlike most others, the plastids of Apicomplexa (Sporozoa and Apicomonada) are not involved in amino acid synthesis [31, 61]. Due to the extensive attempts of the host to patch the biological functions of the endosymbiont, plastid proteomes are highly mosaic with respect to their evolutionary origin. Individual plastid proteins in primary algae originate from eukaryotes (host nuclear genes), cyanobacteria (introduced with the plastid), alphaproteobacteria (introduced with the mitochondrion), and genes from other sources, such as those derived from non-endosymbiotic, horizontal gene transfer (HGT)

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[62, 63]. While metabolic pathways that do not have a eukaryotic counterpart originate almost entirely from the cyanobacterium (e.g., photosystems, type II fatty acid synthesis, and nonmevalonate isoprenoid synthesis), other metabolic pathways consist of a mosaic of cyanobacterial proteins and proteins obtained by HGT or retargeted from other cellular compartments. Typical examples of chimeric metabolic pathways include tetrapyrrole biosynthesis and the Calvin-Benson cycle [64–67]. Complex endosymbioses greatly increase the genetic complexity of organisms by incorporating genes from additional eukaryotic symbiotic partners into the pool. Secondary endosymbiosis brings together two mosaic genomes, that of the primary alga and that of the host (i.e., additional eukaryotic, proteobacterial, and horizontally acquired genes with different evolutionary histories) [44]. Higher-order endosymbioses therefore further increase this genetic complexity. These boosts of gene richness, comparable to genome-wide duplications, could enhance metabolic adaptations to changing environmental conditions and drive the rapid radiation of complex algae. Unfortunately for researchers, though, this genetic chimerism complicates the interpretation of the history of complex endosymbiosis, beyond the conflicting phylogenies of nuclear and plastid genes [44].

4

Reductive Evolution of Plastids The increased genetic complexity of the union of the host and its endosymbiont is a transient stage in endosymbiosis; reductive evolution necessitates a gradual loss of unnecessary genes from this composite. This sudden genetic expansion is faster than exponential, followed by exponential contraction, and this biphasic pattern appears to recur in evolution [68]. Genomic simplification is a process general for both organisms with small effective populations (parasites and endosymbionts) and evolutionarily successful freeliving organisms with larger effective populations [68]. Different genes, however, have different propensities to being lost after a genetic expansion. Orphaned or redundant proteins are lost almost immediately, whereas proteins that function in multiprotein complexes and pathways are more frequently retained [69]. Occasionally, the loss of one component can lead to a domino effect in which the entire module is lost [70]. Continued reductive evolution resulted in an extremely reduced genome of dinoflagellate plastids; the genomes of peridinin-pigmented plastids consist of only a few minicircles (up to 16), i.e., small molecules of 2–3 kbp in size, usually encoding one gene each [69, 71]. Reductive evolution also affects serial complex plastids, although presumably at different rates for individual acquisition events [19]. Acquired plastid endosymbionts were redundant with their host in many metabolic pathways; carbohydrates, fatty acids,

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isoprenoids, and tetrapyrroles are synthesized in both the host and the endosymbiont. These compounds are essential for eukaryotic cells, and thus we hypothesize that there was an intermediate stage during organellogenesis in which both host and endosymbiont pathways were used in parallel [67, 72]. For example, chlorarachniophytes (Bigelowiella natans) and euglenophytes (Euglena gracilis, Fig. 3e) have two redundant pathways for tetrapyrrole biosynthesis, one originating from the host (C4-type) and the other localized in the plastid (C5-type) [65, 67]. Similarly, dinotoms represent an intermediate (or evolutionarily frozen) lineage, as both the host and the diatom endosymbiont operate independent tetrapyrrole pathways (both C5). This arrangement may result from the inability of host-encoded proteins to translocate to the endosymbiont compartment [32, 55, 67]. However, in the long term, streamlining of cellular biochemistry results in the retention of only one of the complementary metabolic pathways, while the other is doomed to disappear [65, 73, 74]. In most other eukaryotic algae studied, tetrapyrroles are synthesized exclusively in the plastid. In some exceptional cases, such as apicomplexans [75], and an early rhodophyte relative Rhodelphis [76], heme synthesis is initiated in the mitochondrion by the C4 pathway and continues in the plastid. While in sporozoans the pathway terminates in the mitochondrion, in apicomonads and Rhodelphis the rest of the pathway putatively localizes in the plastid [75–77]. Of these, however, only chromerids are photosynthetic and thus are the only known phototrophs to synthesize chlorophyll from glycine rather than glutamate [75]. Reductive evolution also affects photosystems to a certain degree, as cyanobacterial photosystems appear to be more complex than those of their plastid descendants. Photosystems and the ATP synthase complex are reduced in plants, primary algae, and complex algae, with chromerids known to have the most reduced photosystems. On the other hand, chromerid photosystem I contains additional subunits and superoxide dismutases (SODs) permanently bound to the photosystem. Thus, reductive and constructive evolution can act simultaneously on the same molecular machinery [28].

5

Loss of Photosynthesis in Complex Plastids Algae do not always keep their photosynthetic capabilities on evolutionary timescales. Loss of photosynthesis can be seen as a continuation of reductive evolution, and the many examples of independent loss of this hallmark plastid feature suggest there might be patterns to this phenomenon. Plastids, surprisingly, often outlast photosynthesis, as nonphotosynthetic (cryptic) plastids have been discovered in most nonphotosynthetic algae upon

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closer inspection. Remarkably, about 50% of dinoflagellate species are heterotrophs or parasites secondarily, implying numerous losses of photosynthesis [32, 78]. Secondarily nonphotosynthetic taxa indeed appear in all other phototrophic lineages, except haptophytes and chlorarachniophytes (Fig. 2). Some lineages tend to produce heterotrophic nonphototrophs, some tend to be parasitic [13, 14, 45]. Colorless lineages of plants and rhodophytes are often parasitic, and while most nonphotosynthetic green algae are freeliving, Helicosporidium is also a parasite [79–81]. Euglenophytes have lost photosynthesis several times, yet the reason is not well understood. The plastid of the phototrophic Euglena gracilis (Fig. 3e) can be bleached by antibiotics or physical stress, and natural hetero-osmotrophic mutants such as Euglena longa are quite common (Fig. 3f) [82, 83]. This tendency to secondary heterotrophy is probably due to an intermediate and highly redundant state of cellular biochemistry in euglenophytes [65]. Cases of complete plastid loss appear to be strikingly rare, and the only widely accepted cases are sporozoans Cryptosporidium spp. and Gregarina niphandrodes [84, 85] and the parasitic dinoflagellate Hematodinium (Fig. 2) [86]. Plastids are thought to be maintained well after loss of photosynthesis because they harbor other essential biochemical pathways [87–90]. The apicoplast of sporozoan parasites (e.g., Plasmodium falciparum), the best-studied relict plastid, illustrates this. It is four-membrane bound, holds a reduced circular genome of about 35 kbp, and does not contain any genes involved in photosynthesis (Fig. 4) (e.g., [87, 91, 92]). The organelle appears to be essential for the survival of the parasite, and its disruption causes the so-called “delayed death effect,” whereby the parasite’s progeny ceases to develop in erythrocytes. The bloodstream form of P. falciparum requires the apicoplast for isoprenoid synthesis, and the insect form relies on its fatty acid and heme synthesis capacity [93]. These compounds are essential for survival in most eukaryotes (but see [94]) and must either be synthesized autonomously or obtained from external sources such as prey or host. The role of the apicoplast in the cell’s biochemistry begins with phosphoenolpyruvate (PEP) and dihydroxyacetone phosphate (DHAP) import from the cytosol. These carbohydrate phosphates are the initial substrates for the abovementioned fatty acid, phospholipid, and isoprenoid biosynthesis (heme biosynthesis requires aminolevulinic acid, an intermediate synthesized by the mitochondrion). The conversion of PEP to pyruvate and acetyl-CoA generates ATP and NADH, both of which are required for phosphorylation and reduction steps in the above metabolic pathways. Performing several intermediate steps of heme synthesis makes apicoplast a biochemical hub of the parasite cell (Fig. 4). Evidence of the actual metabolic functions of other cryptic plastids is growing slowly. Some of them harbor metabolic pathways similar to those of

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Fig. 4 Apicoplast. (a) Apicomplexan plastid as seen in DAPI stained trophozoites of Sarcocystis muris. (b) Electron micrograph of Goussia janae showing four membranes surrounding the apicoplast. Image courtesy of M. Obornı´k (A), and J. Lukesˇ (B). (c) Pathways of the apicoplast. Numbered circles denote enzymes, according to KEGG pathways. Turquoise, glycolysis: 1, hexokinase; 2, glucose-6-phosphate isomerase; 3, 6-phosphofructokinase; 4, aldolase; 5, triose-phosphate isomerase; 6, glyceraldehyde-phosphate dehydrogenase; 7, phosphoglycerate kinase; 8, bisphosphoglycerate mutase; 9, enolase; 10, pyruvate kinase. White, fatty acid synthesis: 1, pyruvate kinase; 2, pyruvate dehydrogenase complex; 3, acetyl-CoA carboxylase; 4, FabD; 5, FabH; 6, FabG; 7, FabZ; 8, FabI; 9, FabB/F; 10, acyl-carrier protein synthase. Red, isoprenoid precursor biosynthesis: 1, DOXP synthase; 2, DOXP reductase; 3, CDP-ME synthase; 4, CDP-ME kinase; 5, MEcPP synthase; 6, HMB-PP synthase; 7, HMB-PP reductase. Purple, phospholipid synthesis: 1, glycerol-3phosphate dehydrogenase; 2, glycerol-3-phosphate acyltransferase; 3, acyl-glycerol-3-phosphate acyltransferase; 4, phosphatidic acid cytidyltransferase; 5, phosphatidylglycerol phosphate synthase; 6, phosphatidylglycerol phosphatase. Brown, heme synthesis: 1, ALA synthase; 2, ALA dehydratase; 3, PBG deaminase; 4, URO synthase; 5, URO decarboxylase; 6, CP oxidase; 7, PP oxidase; 8, heme ferrochelatase. Yellow, ironsulfur cluster assembly machinery SUF, three-step synthesis, seven proteins required. Only important metabolites are shown, ALA – δ-aminolevulinic acid, DHAP – dihydroxyacetone-phosphate, DOXP – 1-deoxy-D-xylulose 5-phosphate, PEP – phosphoenolpyruvate; in bold, production of co-substrates ATP and NADH from PEP

apicoplasts, with amino acid biosynthesis [80, 95–99], while those of Euglena retain only carbohydrate metabolism, hypothesized to redox-balance cytosolic carbon metabolism by uniquely using CO2 as electron sink [82, 83]. To combat parasitic organisms such as apicomplexan parasites, we can exploit their dependence on plastid biosynthetic pathways, if we study them thoroughly. Sporozoans are unicellular, obligate parasites with typical morphological features that animals do not possess, such as the apicoplast and the apical complex, a series of tubular and secretory organelles used to penetrate the host cell. Diseases caused by sporozoans have a high impact on humans, e.g., hundreds of thousands of deaths per year (malaria, toxoplasmosis) and severe economic losses (eimeriosis, toxoplasmosis, cryptosporidiosis). Although various drugs are used to treat malaria, drug-

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resistant strains of Plasmodium are spreading rapidly in affected countries in Asia and Africa. Recently, combined treatment with antibiotics such as azithromycin has been frequently used [100]. Antibiotics target organellar replication, transcription, and translation and affect mitochondria and the apicoplast. Some antimalarials act on proteins unique to the Plasmodium apicoplast, i.e., therapeutic targets that do not exist in the mammalian host, and are presumably less harmful to patients [93, 101]. These new targets include transporters and the biosynthesis of amino acids (pyrimethamine, cycloguanil), fatty acids (thiolactomycin, cerulenin, triclosan), heme (succinylacetone), and isoprenoids (fosmidomycin). Unfortunately, some of these compounds may be toxic to humans at high doses [93, 100], and clinical trials are needed to determine their efficiency. In addition, further research is expected to be conducted to develop even more effective therapeutics.

6

Conclusion The photosynthetic organelles of eukaryotes, the plastids, exhibit extraordinary diversity and have been the driving force for significant changes in the cell biology of many lineages. The evolution of the endosymbiont into an organelle was a gradual process. The host cell had to evolve mechanisms for metabolic exchange and protein import into the endosymbiont, leading to complete dependence of the symbiont on its host and vice versa, and to mutual control by anterograde and retrograde signaling. The relocation of the plastid into the endomembrane system (secondary endosymbiosis), likely occurring in cryptophytes, required the introduction of a more complex targeting and integration machinery. Once established, secondary plastids could have been spread horizontally across eukaryotic supergroups via higher-order endosymbioses, possibly using their unique protein-targeting machinery. In some lineages, most obviously in dinoflagellates, complex plastids were replaced by plastids from other lineages (serial or higher-order endosymbioses). Horizontal spread of plastids between eukaryotic lineages resulted in massive evolutionary radiations, and complex algae (especially diatoms, haptophytes, and dinoflagellates) became key players in Earth’s aquatic environments. Other, nonphotosynthetic complex algae are parasites of animals and humans. Indeed, complex plastids became nonphotosynthetic in many lineages, but most of these organelles persisted and are biochemically essential for their host cells. While complex plastids became an evolutionary advantage over aplastidic lineages, their uniqueness represents a chance to develop specific drug targets to combat some of the deadliest parasitoses in humans.

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Acknowledgments ZF is supported by the Simons Collaboration on Principles of Microbial Ecosystems (PriME) (Grant ID: 970820 to Andrew E. Allen). MO is supported by a grant by the Czech Science Foundation, grants number 21-03224S, and the European Regional Development Fund ERDF/ESF, grant number CZ.02.1.01./0.0/0.0/16_019/0000759. References 1. Obornı´k M (2019) In the beginning was the word: how terminology drives our understanding of endosymbiotic organelles. Microb Cell 6:134–141. https://doi.org/10.15698/ mic2019.02.669 2. Keeling PJ (2013) The number, speed, and impact of plastid endosymbioses in eukaryotic evolution. Annu Rev Plant Biol 64:583–607. https://doi.org/10.1146/annurev-arplant050312-120144 3. Nowack ECM, Melkonian M, Glo¨ckner G (2008) Chromatophore genome sequence of Paulinella sheds light on acquisition of photosynthesis by eukaryotes. Curr Biol 18:410– 418. https://doi.org/10.1016/j.cub.2008. 02.051 4. Armbrust EV, Berges JA, Bowler C et al (2004) The genome of the diatom Thalassiosira pseudonana: ecology, evolution, and metabolism. Science 306:79–86. https:// doi.org/10.1126/science.1101156 5. Burki F, Kaplan M, Tikhonenkov DV et al (2016) Untangling the early diversification of eukaryotes: a phylogenomic study of the evolutionary origins of Centrohelida, Haptophyta and Cryptista. Proc R Soc B 283: 20152802. https://doi.org/10.1098/rspb. 2015.2802 6. Petersen J, Ludewig AK, Michael V et al (2014) Chromera velia, endosymbioses and the rhodoplex hypothesis – plastid evolution in cryptophytes, alveolates, stramenopiles, and haptophytes (CASH lineages). Genom Biol Evol 6:666–684. https://doi.org/10. 1093/gbe/evu043 7. Derelle R, Torruella G, Klimesˇ V et al (2015) Bacterial proteins pinpoint a single eukaryotic root. Proc Natl Acad Sci U S A 112:E693– E699. https://doi.org/10.1073/pnas. 1420657112 8. Hurst GDD (2017) Extended genomes: symbiosis and evolution. Interface Focus 7: 20170001. https://doi.org/10.1098/rsfs. 2017.0001

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Chapter 3 Complex Endosymbiosis II: The Nonphotosynthetic Plastid of Apicomplexa Parasites (The Apicoplast) and Its Integrated Metabolism Nyamekye Quansah, Charital Sarah, Yoshiki Yamaryo-Botte´, and Cyrille Y. Botte´ Abstract Chloroplasts are essential organelles that are responsible for photosynthesis in a wide range of organisms that have colonized all biotopes on Earth such as plants and unicellular algae. Interestingly, a secondary endosymbiotic event of a red algal ancestor gave rise to a group of organisms that have adopted an obligate parasitic lifestyle named Apicomplexa parasites. Apicomplexa parasites are some of the most widespread and poorly controlled pathogens in the world. These infectious agents are responsible for major human diseases such as toxoplasmosis, caused by Toxoplasma gondii, and malaria, caused by Plasmodium spp. Most of these parasites harbor this relict plastid named the apicoplast, which is essential for parasite survival. The apicoplast has lost photosynthetic capacities but is metabolically similar to plant and algal chloroplasts. The apicoplast is considered a novel and important drug target against Apicomplexa parasites. This chapter focuses on the apicoplast of apicomplexa parasites, its maintenance, and its metabolic pathways. Key words Secondary endosymbiosis, Secondary plastid, Complex plastid, SAR

1

Introduction Apicoplast is a relict nonphotosynthetic plastid found in most Apicomplexa, a phylum of unicellular eukaryotes (or protists), which mainly comprises obligate intracellular parasite, including pathogens of medical importance, such as the malaria parasite, Plasmodium spp. The apicoplast (Apicomplexa plastid) has been acquired by the secondary endosymbiosis of red algae and is thus delimited by four surrounding membranes (see below). There is only a single apicoplast in each parasitic cell, which then elongates and branches before dividing and being attributed to each future daughter cell before cytokinesis of the parasite. The apicoplast is usually found in close vicinity to the mitochondrion and was even thought to be bound to it. The apicoplast is essential for the

Eric Mare´chal (ed.), Plastids: Methods and Protocols, Methods in Molecular Biology, vol. 2776, https://doi.org/10.1007/978-1-0716-3726-5_3, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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Fatty acids Apicoplast

Glycolysis

PA synthesis

FASII pathway

Pyruvate

AcetylCoA

GA3P

DOXP pathway

Replication DNA

PEP DHAP

Transcription RNA

Isoprenoid precursors

? Translation

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FeS cluster synthesis Heme synthesis

?

?

Proteins

?

Mitochondrion (Cytochrome)

?

Nuclear encoded proteins

Protein prenylation ? Mitonchondrion (quinones) ? Others (dolichol, Abscissic acid) ?

Fig. 1 The apicoplast of Plasmodium falciparum: a relict nonphotosynthetic plastid involved in essential metabolic functions. The apicoplast possesses a 35 kilobase genomic DNA which is replicated, transcribed into RNA, and translated into proteins via its own machineries of prokaryotic origin. The apicoplast hosts four remarkable metabolic pathways: a type II fatty acid synthesis pathway (FASII), a non-mevalonate pathway, or 1-deoxy-D-xylulose 5-phosphate (DOXP) pathway, which catalyzes the synthesis of isoprenoid precursors, a heme synthesis pathway and an iron/sulfur (FeS) cluster synthesis pathway. The FASII and DOXP pathways both rely on the import of triose phosphates, i.e., phosphoenolpyruvate (PEP) and dihydroxyacetonephosphate (DHAP), generated in the parasite’s cytosol and converted into pyruvate, acetylCoA, and glyceraldehyde-3phosphate (GA3P) in the apicoplast. (Adapted from [3])

parasite survival and, besides the loss of photosynthetic capacities, is metabolically similar to plant and algal chloroplast. Biochemical analyses combined with robust bioinformatic analysis by mining P. falciparum genome enabled the identification and assembly of apicoplast proteins involved in five major metabolic pathways of plant origins: fatty acid synthesis, lysophosphatidic acid synthesis, isoprenoid synthesis (Fig. 1), heme synthesis (Fig. 2), and Fe-S cluster synthesis (Fig. 3), which are discussed below [1, 2]. In contrast to pathogenic bacteria, Apicomplexan parasites are unicellular eukaryotes, and they share numerous metabolic pathways with their animal hosts, making therapeutic target development difficult. Two Apicomplexa genera, Plasmodium and Toxoplasma, which are responsible for serious diseases in humans, have been the focus of most studies on apicoplasts in this parasite group. Here, we will focus on the apicoplast of Plasmodium spp., the agent of malaria,

Roles and Maintenance of the Apicoplast

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ALA Glycine

PBGS

ALA

ALAS Succinyl-CoA

PBG PGDB/ UroS

Heme b Cytosol

FeChl

PPIX

Uro

Mitochondrion

Apicoplast

UroD PPo

PP

CPIII

?

CPIII

CPO PP

Fig. 2 The apicoplast heme pathway. The heme biosynthetic pathway starts in the mitochondrial matrix with the condensation of succinyl-CoA and glycine, generating δ-aminolevulinic acid (ALA) by an ALA synthase (ALAS). ALA is then exported to the cytosol and then imported into the apicoplast by an unknown transporter. ALA is then successively converted into porphobilinogen (PBG), hydroxymethylbilane, uroporphyrinogenIII (Uro), and coproporphyrinogenIII (CPIII) via the apicoplast resident proteins PBG synthase (PBGS), PBG deaminase (PBGD) and uroporphyrinogen deaminase (UroD), respectively. CPIII is exported to the parasite cytosol to be oxidized into protoporphyrinogenIX (PP) by a coproporphyrinogen oxydase (CPO). PP can then be imported into the mitochondrial intermembrane space where it is converted into protoporphyrinIX via a PP oxydase (PPo). A ferrochelatase (FeCHl) eventually adds a ferrous iron at the center of the tetrapyrole structure to form hemeb

the death toll due to which is estimated close to one million people per year, mainly children via infections with one of the most virulent strain, P. falciparum. Appearance and spreading of Plasmodium lines resistant to commonly used drugs, even to artemisinin, the front-line malaria drug, and the extreme difficulty encountered in developing efficient vaccines against some species, both argue for the pressing need for new targets and new drugs. The apicoplast therefore represents a unique and promising target to fight these increasingly resistant parasites against which there is no current vaccine [3].

2 Apicoplast History and Origin Although observed for decades, apicoplasts were first identified as the chloroplast counterpart of the Apicomplexa parasites in the late 1990s, which came as a major breakthrough in the biology and

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alanine

cysteine

ADP

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SufC

SufC SufC

SufA

S2-

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[Fe-S]

SufE

SufB SufD

NFU

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Holo-protein

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Fd-red

e-

FNR Fd-ox

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Fig. 3 The apicoplast [Fe-S] cluster synthesis pathway. The [Fe-S] pathway is initiated by the scavenging of a sulfide group from a cysteine residue by SufS. This sulfide group is carried by SufE, which works in tandem with SufS. An iron atom is then added to the sulfide group to form a pre-[Fe-S] cluster, held by the multimeric ABC carrier formed by SufC, SufB, and SufD. This BC2D carrier complex activates the pre [Fe-S] into a [Fe-S] cluster by transferring electron from the Ferredoxine (Fd) donor, which, thus, gets oxidized (Fd-ox). Fd-ox is then reduced (Fd-red) by a Ferredoxine NADP+ reductase (FNR). The [Fe-S] cluster is actively transferred by the BC2D complex to the next carrier NFU. SufA then catalyzes binding of the [Fe-S] cluster to a cysteine residue of the targeted protein (apo-protein), ready to perform its function (holo-protein)

evolutionary origin of this group of organisms then considered as typical animal cells [4–6]. The apicoplast has been acquired by the unique secondary endosymbiosis event of a unicellular algae, whose origin has long been debated, and that gave rise to the large Chromoalveolates group comprising Apicomplexa but also dinoflagellates and even ciliates. The 2008 discovery of Chromera velia—the closest relative of Apicomplexa retaining a photosynthetic apicoplast—confirmed the red algal origin of the apicoplast [7]. Chromera velia is an exciting new model to understand the apicoplast evolution from an autotrophic (photosynthetic) to the heterotrophic stage found in Apicomplexa parasites. The apicoplast is found in nearly all members of the Apicomplexa phylum except Cryptosporidium sp., which seems to have lost it in a later event [8], similarly to the ciliates. As a consequence of its endosymbiotic origins, the apicoplast is limited by four surrounding membranes, in Apicomplexa parasites, including Plasmodium spp.. The two

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innermost membranes together form the outer and inner envelope, as found in all primary plastids and correspond to the membranes of the initial cyanobacterial ancestor. The third membrane (from within) or periplastid membrane corresponds to the plasma membrane of the red algal ancestor, and the outermost membrane is likely the remnant of the phagotrophic membrane from the second endosymbiotic event. Because of its algal origin, the apicoplast harbors a broad range of prokaryotic/plant-like pathways (homologous to those in the plant or algal chloroplast) that are unique to the parasite. Importantly, the apicoplast is involved in unique and vital biological processes for the parasite [2, 9]. Considerable effort has thus been put into unraveling the role of that peculiar nonphotosynthetic plastid in apicomplexan parasites. However, it is unclear why parasites are dependent on their apicoplasts and how their metabolic pathways are required under specific cellular context, intracellular environment, and life stage. Deeper analysis of the content and the metabolic roles of the apicoplast have long been hampered by the lack of a purification method for the organelle. A novel protocol to isolate the apicoplast has provided the first lipid profile of the apicoplast and enabled new discoveries on this peculiar organelle [10].

3

Genome Maintenance and Protein Import The apicoplast is one of the genetically active organelles in Apicomplexa. All Apicomplexa parasites, except Cryptosporidium spp., possess three genetically active compartments: the nucleus, the mitochondrion and the apicoplast, two semi-autonomous organelles of prokaryotic origins, thus containing their own circular genome. The apicoplast genome is only 30–35Kb encoding for about 50 apicoplast proteins, which makes it the smallest plastid genome to date. Upon endosymbiosis events, the apicoplast had undergone a drastic reduction of the original prokaryotic genome, which was either lost (such as photosynthesis-related genes) or laterally transferred to the host genome in the nucleus. This transfer of genetic material from plastid to host is still an ongoing process that can be directly measured. As a direct consequence, most apicoplast proteins are encoded in the nucleus and later imported in the apicoplast (or so-called NEAT proteins: Nuclear Encoded Apicoplast Targeted proteins). Bioinformatic analysis allowed identifying this nuclear-encoded repertoire of more than 350 proteins forming the virtual proteome of the apicoplast, which greatly benefit our understanding of the apicoplast functions [11]. Even if encoding only a fistful of proteins, the apicoplast genome is essential for proper apicoplast functions and thus needs to be replicated, transcribed, and translated via the organelle’s own

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maintenance machinery. These prokaryotic machineries of apicoplast genome maintenance can be fully considered as part of the organelle’s integrated pathways [3]. 3.1

Replication

In Plasmodium blood stages, replication of both the apicoplast and the nucleus genomes is simultaneously initiated at the beginning of schizogony. Replication is initiated by the formation of replication bubbles between the genes of the small and large rRNAs. All proteins involved in apicoplast DNA replication are encoded by the nucleus and later imported in the organelle. Several of these are considered or used as targets against malaria, such as DNA gyrases, inhibited by ciprofloxacin. The main enzymatic functions of the replication process (DNA polymerase, DNA primase, and helicase) are believed to be held by a multifunctional protein named Pfprex, which is considered a potential drug target.

3.2

Transcription

The apicoplast genome transcription is a highly coordinated event that occurs at the schizont stage. Transcription relies on a chloroplast-like RNA polymerase, which comprises three subunits RpoB, RpoC1, and RpoC2, which are encoded by the apicoplast genome and a nuclear-encoded subunit named RpoA. To date, there is, however no experimental evidence of apicoplast transcriptional activity.

3.3

Translation

The apicoplast encodes most of the prokaryotic machinery required for the translation of its proteins such as all rRNAs, 17 or 18 ribosomal proteins, a full set of 35 tRNAs, and the elongation factor EF-Tu. On the other hand, its genome does not express any tRNA synthetase, which catalyzes the specific attachment of each amino acid to its corresponding tRNA. Most of these specific apicoplast tRNA synthetases are encoded in the nucleus and later imported in the organelle, alongside some ribosomal proteins, initiation, elongation, and release factors. Interestingly, some genes’ tRNA synthetases genes are only present as single copies in the nuclear genome although they are required in both the apicoplast and the cytosol. Recent studies showed that the subsequent proteins can be dually targeted to fulfill their function in each compartment. Apicoplast translation is an acknowledged target for antimalarials, mostly directed against rRNA and ribosomes. For instance, doxycycline is an antibiotic used as a prophylactic against malaria, which specifically blocks the apicoplast translation and leads to a delayed-death phenotype, typically observed when disrupting the apicoplast.

3.4

Protein Import

As mentioned earlier, apicoplast presence, maintenance, and metabolic functions rely on the trafficking of more than 300 nuclearencoded proteins from the cytosol into the apicoplast. Two import systems have been identified to date, a canonical import pathway and a vesicular transport pathway. The former is the most

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characterized and is dependent on an N-terminus peptide sequence referred to as a bipartite leader. This leader is necessary and sufficient for the import of most proteins into the organelle. This sequence is composed of a signal peptide (SP), which directs the neosynthesized polypeptide during its translation into the ER, and a transit peptide (TP) that mediates the entry into the apicoplast. The TP contains charged amino acids but lacks a consensus and any secondary structure. The mechanism by which the TP is recognized remains unclear. Trafficking from ER to apicoplast is direct, likely vesicular and does not involve the Golgi apparatus, unlike the secretory pathway. The latter is much less understood. The latter pathway mainly concerns outermost membrane proteins that lack a bipartite import signal, and its current consensus states a vesicular transport involving the recognition of an internal signal comprising putative tyrosine and glycine within the protein sequence. The canonical pathways require trafficking machineries, which allows the protein’s transport through each of the four surrounding apicoplast membranes. The import machinery through the two innermost membranes, originally the envelope from the chloroplast ancestor, is performed via the plastid specific machineries called TOC (translocon of the outer chloroplast membrane) and TIC (translocon of the inner chloroplast membrane). Each of these translocons is composed of several proteins, performing different roles during the protein translocation, such as chaperones, import channel, and peripheral proteins. Not all protein partners of the complexes have been experimentally characterized in Apicomplexa. However, the most important ones have either been studied or predicted such as Tic20, an essential part of the core pore complex, and Tic22, an essential chaperone. The import machinery for the periplastidial membrane (i.e., third apicoplast outermost membrane) is one of newly acquired or invented machineries after the complex apicoplast and parasite evolution. This enabled the parasite to “invent” new strategies by diverting preexisting systems for novel functions. Indeed, the Endoplasmic Reticulum Associated Degradation (ERAD) machinery originally reexports misfolded proteins out of the ER for further degradation by the proteasome (and preubiquitinylation of concerned proteins). This machinery, including its core membrane protein Der1 and the subsequent ubiquitinylation system, including plastid ubiquitin-like protein and E1/E2/E3 ubiquitin ligases, was found and then characterized in the parasites [12]. Surprisingly, this complex was localized at the apicoplast and found responsible for the essential import of proteins through the periplastidial membrane, i.e., an “outside-in” direction from the original transport flux and in a different compartment [13, 14]. Protein trafficking from the ER through the outermost apicoplast membrane remains questionable but may have partly found an answer via recent studies. The current hypothesis supports a vesicular transport sorted, towards the apicoplast,

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via the recognition of membrane phosphoinositides generated by PI3P kinases [14]. This mechanism, if confirmed as the canonical transport cargo of the outermost membrane, would be another recycled machinery typically used to target proteins to early endolysosomes in animal cells.

4

Apicoplast Metabolic Pathways

4.1 Heme Synthesis Pathway, the Shared Burden with Sister Mitochondrion

Heme is an essential prosthetic group of many proteins, which is formed of a tetrapyrrole molecule including a chelated ion in its structure, either Fe2+ or Mg2+. Heme allows its hosting protein to reversibly bind diverse small molecules or electrons. A well-known example of such protein is hemoglobin, which can bind oxygen and allows its transport in erythrocytes. Heme synthesis pathway is one of the most complex of the apicoplast metabolic functions. Indeed, the pathway is carried out within three different compartments: It is initiated in the mitochondrion, continues in the apicoplast, then in the cytosol to eventually end where it started, in the mitochondrion. This shared pathway could explain the tight association of the two organelles observed throughout the parasite life cycle. One could also wonder why heme synthesis is commonly accepted as one of the apicoplast pathways. The answer lies, again, in the evolution of the apicoplast and how genomes and enzymes of photosynthetic and nonphotosynthetic origins were reshuffled to obtain the pathway as can now be observed. The pathway likely evolved from the coexistence of a chloroplast and a mitochondrialcytosolic tetrapyrrole pathway, which can still be found today in Euglena (a protist with a secondary plastid of green origin). That evolutionary hypothesis was confirmed by the discovery and phylogenetic analysis of the heme pathway in Chromera velia, the closest homolog of Apicomplexa parasites still harboring a photosynthetic apicoplast. The heme synthesis starts with a conversion of glycine and succinyl-CoA into δ-aminolevulinic acid (ALA) by ALA synthase (ALAS) in mitochondrion. ALA is then exported towards the apicoplast stroma via a yet-to-determine transport system. ALA is then successively converted into porphobilinogen (PBG), hydroxymethylbilane, uroporphyrinogenIII (Uro), and coproporphyrinogenIII (CPIII) via the apicoplast resident proteins PBG synthase (PBGS), PBG deaminase (PBGD), and uroporphyrinogen deaminase (UroD), respectively. It is noteworthy to mention that in P. falciparum, PBGD is believed to generate both hydroxymethylbilane and Uro and thus to possess both PBGD and Uro synthase (UroS) activities. This statement is based on the initial absence of UroS in P. falciparum genome and further biochemical evidence proving the proper activity of UroS are still needed. However, a putative homolog of UroS has been identified in P. falciparum

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genome, which has yet to be characterized. Once generated, the CPIII is then exported to the parasite cytosol to be oxidized into protoporphyrinogen IX (PP) by a coproporphyrinogen oxydase (CPO). PP can then be imported back into the mitochondria where it is converted into protoporphyrinIX at the inner mitochondrial membrane via a PP oxydase (PPo). A ferrochelatase (FeCHl) eventually adds a ferrous iron at the center of the tetrapyrole structure to form heme b [15]. The utilization of heme is quite relevant in P. falciparum since Plasmodium blood stages live in and feed from the red blood cell hemoglobin. Once hemoglobin is engulfed by the parasite, it goes to the parasite’s food vacuole where it can be degraded into amino acids by a range of proteases and used for protein synthesis. This degradation releases heme, which is toxic for the parasite due to high oxidative stress of iron contained in the heme molecules. To confine heme, the parasite polymerizes it into an insoluble pigment called hemozoin, which is imprisoned in the food vacuole. So, what use for neo-synthesized heme when an unlimited supply is available from the host? Plasmodium genome mining suggests that the mitochondrial electron transport chain is a likely acceptor because it requires heme-containing cytochromes (cyt c, b, c1) and possesses enzymes required for heme utilization (cox1, cox10, cox15). The electron transport chain is essential for parasite survival and is already the target for therapeutic intervention by atovaquone. Recent studies have shown that heme requirement for the parasite are strongly affected by changes in parasite energy requirements. These changes can be due to changes to the host environment or the stage of the life cycle the parasite is at [16]. A study showed, by radioactive metabolic labeling, that both neosynthesized-heme and hemoglobin-generated heme are found incorporated into both hemozoin and mitochondrial cytochromes during blood stages [17]. This suggests that the two sources of heme provide a redundant supply for the parasite. It was also shown that this heme synthesis pathway was only essential during the mosquito and liver stage in P. berghei, its absence leading to a drastic reduction in oocyst formation, complete disruption of sporozoite formation, and invasion. The results could not determine the precise role of the pathway during these stages. 4.2 Iron-Sulfur Clusters, The Mystery Function

In the apicoplast, there is also a pathway to synthesize iron-sulfur, [Fe-S], clusters. [Fe-S] cluster is an active center, which is formed of iron and sulfur atoms attached to its protein via a cysteine residue and plays an indispensable role in some enzymatic functions as prosthetic groups. However, this [Fe-S] cluster pathway has received the least attention due to the lack of known inhibitors potentially disrupting this pathway or its potential targets. Progress has nonetheless been made in the study of [Fe-S] cluster pathway in Toxoplasma gondii, showing that this pathway had pleiotropic

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effects on parasite metabolism and survival. The loss of the cluster impacts the parasite’s motility, fatty acid generation, and parasite division [18]. The clusters’ role is to accept electrons from a donor and to transfer them onto an acceptor, thus having a strong redox potential. The [Fe-S] prosthetic group is usually added posttranslationally onto the peptide sequence of an apo-protein, which, in return, activates the protein (i.e., holo-protein). This reaction is usually catalyzed by the mitochondrial iron-sulfur cluster (ISC), the cytoplasmic iron-sulfur protein assembly (CIA) or the plastid-based sulfur utilization factor (SUF). Homologs of the mitochondrial ISC could be identified in P. falciparum, whereas the CIA machinery seems incomplete and thus inactive. P. falciparum also possesses a complete SUF pathway, which proteins are predicted to be targeted to the apicoplast. In the apicoplast, [Fe-S] cluster synthesis is initiated by the dual action of SufC and SufE. SufC is a cysteine desulfurase that catalyzes scavenging of a sulfide group from a cysteine, releasing alanine. The sulfide group is then transferred by SufE, which carries the group and activates SufC. Iron, likely from hemoglobin digestion, is then added to the sulfide group for a pre-[Fe-S] cluster. This pre-cluster is transferred to a scaffold-carrier complex formed by SufB, C, and D, assembled in a BC2D multimeric complex. They, thus, form a nonintegral ABC transporter capable of (i) carrying the precluster, (ii) transferring electron to activate the [Fe-S] cluster, and (iii) transferring this cluster to the next scaffold complex fueled by ATP hydrolysis. The [Fe-S] cluster is transferred to NFU, another scaffold partner, and the final transfer to the target apo-protein is believed to be catalyzed by SufA [19]. Out of the seven proteins putatively identified to perform apicoplast [Fe-S] cluster assembly, one is encoded by the apicoplast genome,(SufB), five have a predicted N-Ter bipartite leader (SufC, SufD, SufE, SufS, and NFU), and only two have been experimentally confirmed to be apicoplast resident proteins, SufC and NFU. The assembly of [Fe-S] clusters heavily relies on the Ferredoxin (Fd)-Ferredoxin NADP+ Reductase (FNR) duo as Fd serves as an electron donor to the precluster and FNR regenerates some reduced Fd. Fd is a holoprotein that participates in the pathway but also requires a [Fe-S] cluster to be functional. The role of the apicoplast [Fe-S] cluster pathway remains unclear as IspG (aka LytB) from the DOXP pathway is the only experimentally proven acceptor of apicoplast-made [Fe-S]. LipA, catalyzing the synthesis of lipoic acid, also seems a likely electron acceptor from Fd. A study analyzing an NFU KO parasite line showed that the pathway is not essential throughout the whole life cycle of the rodent malaria parasite P. berghei [20]. The only significant difference was a reduced formation of merosomes (i.e., infected liver cells) in vitro. These results are relatively surprising

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knowing that both the isoprenoid pathway and the fatty acid pathway require holo-proteins and are essential for the parasite. Several apicoplast proteins are likely candidates for the requirement of [Fe-S] cluster: IspG and IspH, involved in the apicoplast isoprenoid pathway, LipA required for apicoplast lipoic acid synthesis, MiaB, a tRNA methylthiotransferase and ferredoxin (Fd), part of the apicoplast [Fe-S] cluster synthesis pathway. All of these candidate acceptors are nuclear-encoded and thus need to be imported posttranslation. As a consequence of this import, they need to be unfolded to reach the apicoplast stroma. Due to this mechanism and protein import process, it is likely that the apicoplast is the sole de novo source for [Fe-S] clusters for these apicoplast proteins rather than from the mitochondria. Recent studies in Plasmodium also point at the role of SufS for providing sulfur to allow the [Fe-S] synthetic pathway but also sulfur for the use of MnmA to modify apicoplast tRNAs [21]. 4.3 DOXP Pathway, The New Promised Land for Anti-parasitic Therapeutic Developments?

In the apicoplast, there is a DOXP (1-deoxy-D-xylulose-5-phosphate) synthesis pathway, also known as the non-mevalonate pathway (Fig. 4). The DOXP pathway and its eukaryotic counterpart, the mevalonate pathway, generate precursors of isoprenoids, which are an important class of lipid compounds essential for the synthesis of sterols, chlorophylls, and many components in both animal and plant cells. The mevalonate pathway usually takes place in the cytosol of many eukaryotes whereas the DOXP pathway is found in many bacteria, plastids, and, thus, Apicomplexa. The pathway generates two end products: Isopentenyl pyrophosphate (IPP) and Dimethylallylpyrophosphate (DMAPP), which form the repetitive units in isoprenoids and can be incorporated in numerous parasitic metabolites from protein anchors to electron transport chains and even putative carotenoids. Experimental data on the incorporation of apicoplast IPP and DMAPP are lacking, making it difficult to grasp the exact role of this pathway for the parasite despite its obvious requirement. DOXP pathway has sparked renewed interest in the past few years since it might be the sole essential function of the apicoplast during malaria blood stages [22]. On one hand, the raison d’eˆtre of the apicoplast seems much more complex, directly linked to life stages and environmental conditions. And indeed, as further developed below, it seems that other metabolic functions, such as the FASII, become essential depending on the host’s nutritional content. This points to the fact that essentiality vs dispensability is not a binary answer but is rather more dependent on the physiological conditions found in the host that make the importance of pathways. So one should be careful when interpreting results of essentiality scores from recent whole genome loss-offunction screens in Toxoplasma gondii [23] and Plasmodium falciparum, which provide an idea of importance but only in high host nutrient content, hence potentially hiding the real importance/

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Glc MEP IspD

Glycolysis

PK PEP TPT PEP

CDP-ME

IspE

IspF Pyruvate

DXS

MEc-PP

DOXP IspC

IspG

GA3P

HMB-PP

TPI DMAPP IPP

DHAP TPT DHAP Glycolysis

CDP-MEP

IspH

Apicoplast

Glc

?

DMAPP

IPP

Geranyl-PP

Fig. 4 The apicoplast DOXP pathway. Cytosolic glycolysis provides the two triose phosphate precursors required for the apicoplast DOXP pathway: phosphoenol-pyruvate (PEP) and dihydroxyacetone phosphate (DHAP). Both are imported into the apicoplast by the triose phosphate transporters (TPT). PEP is converted into pyruvate (pyr) via the action of a pyruvate kinase (PK), and DHAP into glyceraldehyde-3-phosphate (GA3P) by a triose phosphate isomerase (TPI). Initiation of the DOXP pathway is catalyzed by a DOXP synthase (DXS), which allows the condensation of pyr and GA3P to form 1-deoxy-D-xylulose-5-phosphate (DOXP). DOXP is converted into methylerythritol-4-phosphate (MEP) by a DOXP reductase (IspC). MEP is then converted into 4-diphosphocytidyl-2-C-methyl-D-erythritol 2-phosphate (CDP-MEP), 2-C-methyl-D-erythritol 2,4-cyclopyrophosphate (MEcPP), and (E)-4-Hydroxy-3-methyl-but-2-enyl pyrophosphate (HMB-PP) by a 4-diphosphocytidyl-2-C-methyl-D-erythritol synthase (IspD), a 4-diphosphocytidyl-2-C-methyl-D-erythritol kinase (IspE), a 2-C-methyl-D-erythritol 2,4-cyclodiphosphate synthase (IspF), and a HMB-PP synthase (IspG). Eventually, an HMB-PP reductase (IspH) catalyzes the last step of the apicoplast DOXP pathway, which results in the formation of isopentenyl pyrophosphate (IPP) and dimethyallyl pyrophosphate (DMAPP). IPP and DMAPP can then be exported by an unknown mechanism to be condensed into geranyl pyrophosphate (geranyl-PP). Geranyl-PP serves as a central precursor for all downstream products and pathways. (Adapted from [26])

function of a proteins of interest. Furthermore, another study showed that even when disrupted and absent as a whole by IPP supplementation and disruption of the DOXP pathway, the apicoplast remains metabolically active in the form of vesicles, hinting at the possibility of more than the DOXP function being essential in the apicoplast [24]. This was further confirmed by the recent possibility of ablating the DOXP pathway in the apicoplast by complementation with mevalonate, hinting at its putative dispensability when host nutrients are compensating [25]. On the other hand, the DOXP pathway constitutes a very promising target to

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treat patients since the blood stages are responsible for the symptoms and deadly consequences of malaria. The DOXP pathway is initiated by the condensation of pyruvate and glyceraldehyde-3-phosphate (GA3P) and is catalyzed by a DOXP synthase (DXS). This reaction generates DOXP, giving its name to the pathway. Both the DXS substrates are intermediates of glycolysis, which takes place in the cytosol and represents the main source of ATP for the parasite. Together with the FASII pathway (and the related lipid synthesis pathway, see below), the DOXP pathway relies on the import of two important glycolytic metabolites into the apicoplast, phosphoenolpyruvate (PEP) and Dihydroxyacetone phosphate (DHAP). These metabolites are imported by two transporters, homologs to the chloroplast phosphate transporters and the sole apicoplast transporters characterized to date: the outer Triose phosphate transporter (oTPT) and the Inner Triose Phosphate Transporter (iTPT). Upon their import, PEP is transformed into pyruvate by a pyruvate kinase (PK) and DHAP is converted into GA3P by a triose phosphate isomerase (TPI). Once generated from these essential imported metabolites, DOXP is then converted to methylerythritol-4-phosphate (MEP) and eventually to the end products the isoprenoid precursors IPP and DMAPP via the consecutive action of a DOXP reductase (IspC), a CDP-MEP synthase (IspD), a CDP-ME kinase (IspE), a MEcyclodiphosphate synthase (IspF), a hydroxymethyl butenyl-PP synthase (IspG), and a HMP-PP reductase (IspH). All seven enzymes of the pathway are predicted to be nuclear encoded [3]. The DOXP pathway was initially shown to be essential since fosmydomycin, an inhibitor of IspC and known antibiotic, killed P. falciparum blood stages. A more recent study, also using fosmydomycin treatment combined with IPP complementation, showed that the DOXP pathway is likely to be the only essential function of the apicoplast during the acute phase of the disease, the blood stage [22]. These data suggest that fosmydomycin is a promising candidate for malaria treatment. It has therefore been tested for clinical trial on the field in combination with clindamycin but showed lesser efficacy than front-line artemisinin-based combination therapies. Nonetheless, other clinical trials have shown that fosmidomycin– piperaquine combination treatment was effective for uncomplicated malaria making it a promising treatment candidate [26]. Controversial reports have been published on the potential of fosmidomycin as a prophylactic and its putative inhibition of apicoplast development in the liver stage and the issue is yet to be solved. Recent studies have shown that the inhibition of isoprenoid biosynthesis prevented the branching and inheritance of the apicoplast by the daughter merozoites [27]. So, yes, the DOXP pathway is essential but what does it provide to the parasite? One obvious role is the synthesis of prenyl groups (repetitive units of IPP:

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Geranyl/farnesyl/geranylgeranylpyrophsophates) for protein anchoring. Inhibitors of prenyltransferases are active against P. falciparum. Furthermore, recent analysis using proteomic approaches have shown that many proteins (between 15 and 25) are actually prenylated in Plasmodium falciparum blood stages, especially trafficking protein Rabs and SNARES [28, 29]. Yet, no protein has been proven to be directly prenylated via the apicoplast DOXP products. Another predicted role for the DOXP is tRNA isopentenylation, by addition of DMAPP to a tRNA moiety. Such modification is essential for proper tRNA function, but again, no experimental evidence yet supports this hypothesis. Dolichol, an ER polyprenyl unit required for polysaccharide transport and thus GPI anchor synthesis, was drastically reduced under fosmidomycin treatment. These data indirectly support the putative role of the DOXP pathway for their synthesis. Ubiquinones and menaquinone (mitochondrial electron transporters) could require prenylation from the DOXP as suggested by the loss of mitochondrial potential in a TPT KO mutant in T. gondii. Finally, a recent report that both carotenoids and abscisic acid, typical plant and chloroplast metabolites usually generated from neosynthesized isoprenoid precursors in plants, could be synthesized from apicoplast IPP and DMAPP [30]. 4.4 Fatty Acid Synthesis and Lipid Precursor Synthesis: A Dispensable Yet Essential Pathway

Apicoplast possesses a set of enzymes for prokaryotic type II fatty acid synthesis pathway (FASII) and potentially for the synthesis of major glycerophospholipids (i.e., major membrane lipids) precursor (Fig. 5). Plasmodium had long been thought unable to make fatty acids (FA, i.e., long aliphatic chains used to synthesize membrane glycerophospholipids) for its membrane biogenesis, rather relying on its host cell to provide these essential molecules. The discovery of the apicoplast and its FASII challenged this dogma, suggesting that the parasite was able to synthesize fatty acids for further glycerophospholipids synthesis on top of massively scavenging from its host. Indeed, the parasite requirements for lipids are huge: Upon a single sporozoite infection into a liver cell, 20,000–70,000 new merozoites with their own membrane compartments are made and released into the blood flow. Then, a single merozoite infection into a red blood cell induces a dramatic lipid increase of an average of 400% in the infected red blood cell. Due to the cyanobacterial origin of this pathway, it was immediately considered a very promising drug target and has thus attracted much attention for the search of herbicides/antibiotics with anti-parasitic properties targeting the FA synthesis. A study even showed that triclosan, a known herbicide inhibiting FASII, was able to kill P. falciparum blood stages and to cure infected mice, thus raising high hopes for the development of new efficient treatments. In 2008, however, two independent studies succeeded in generating parasite lines with a disrupted FASII pathway without any effect on

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Glc Apicoplast

Glycolysis

PEP TPT PEP PK

Pyruvate FabH

PDH Acetyl-CoA

ACP

Malonyl- ACP

ACP

ACP FabD

Malonyl-CoA

FabG

O

ACCase

Lipoic acid

O O

FabF

O OH

ACP AC FabI

FabZ

O

Octyl- ACP ATS2 ?

?

ATS1

PA

C12:0-C16:0 LPA ?

?

Membrane lipid synthesis

ER

Fig. 5 The apicoplast FASII and PA synthesis pathways. The apicoplast harbors the FASII biosynthetic machinery. Cytosolic glycolysis results in the production of phosphoenolpyruvate (PEP), which is translocated across the four membranes of the apicoplast by triosephosphate transporters (TPT) and converted into pyruvate in an ATP-producing reaction catalyzed by a type II pyruvate kinase (PK). This is then converted to acetyl-CoA, the precursor of FASII, by the lipoylation-dependent pyruvate dehydrogenase (PDH). Initiation of FASII occurs by the conversion of acetyl-CoA to malonyl-CoA by ACCase1, transfer of the malonyl group to ACP by FabD, and condensation with an acyl-promed FabH, forming α-ketoacyl-ACP. This then enters FASII elongation with additional malonyl units being incorporated into the growing acyl chain by sequential action of FabB/F, -G, -Z, and –I, producing octyl-ACP and other fatty acids most likely up to C16 in length. These fatty acids, and especially C14:0 (myristic acid), are then used by a glycerol-3-phosphate acyltransferase (ATS1) to form lysophosphatidic acid (LPA), which is then exported to be utilized as a central precursor for the bulk phospholipid synthesis. Fatty acids and LPA could be used by an acylglycerol-3-phosphate acyltransferase (ATS2) to form phosphatidic acid (PA), whose role and function are yet to be determined. (Adapted from [26])

the parasite development during blood stages [31, 32]. Both studies confirmed that the pathway was only essential during the liver stage and thus a target for prophylactics rather than for treating patients. P. falciparum apicoplast was successfully purified and its lipid composition was determined. This study showed that the FASII pathway could indeed become metabolically active under limiting lipid resources, confirming previous transcriptomic

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observations from infected patients [10]. Furthermore, recent studies have given direct evidence that the parasite could modulate its lipid acquisition in response to changes in the host cell’s nutritional environment. It does that by either increasing the uptake of lipids from the host cell and its environment, when the latter is rich in nutrients, or increasing the de novo synthesis of lipids if the host’s nutritional environment is poor [24, 33–35]. Together, these important results demonstrate that the FASII can become essential when host lipid resources are scarce, and/or dispensable under high host nutrient content. Again, the current consensus was overturned, and it emphasized the importance of lipid neosynthesis and the metabolic flexibility of the parasite. Apicoplast fatty acid synthesis pathway begins with the import of PEP, glycolytic intermediates from the cytosol by the Triose phosphate transporters (TPTs). Thus, the apicoplast fatty acid synthesis is all depends on the glycolysis. PEP is then first converted to pyruvate and then to acetyl-CoA via the action of a PK and a multimeric pyruvate dehydrogenase complex (PDH). Acetyl-CoA is transformed into malonyl-Coa by the action of an acetyl-CoA carboxylase (ACCase). The malonyl group is then transferred to the acyl carrier protein (ACP), which is the core protein of the FASII that brings substrates from one enzyme to another. This transfer is catalyzed by a malonyl-CoA:ACP malonyltransferase (FabD). The fatty acid synthesis is initiated by the condensation of malonyl-ACP (3 carbons) and acetyl-CoA (2 carbons) by a keto-acyl-ACP synthase (FabD), which generates acetoacetyl-ACP (4 carbons). This four-carbon acyl group then enters the elongation cycle, which allows the addition of 2 carbons per cycle from a malonylACP and enables the formation of acyl-ACPs (i.e., fatty acid-ACP). The elongation cycle is a four-reaction cycle catalyzed by (i) an NADPH dependent β-keto acyl ACP reductase (FabB/F), (ii) an enoyl-ACP hydrase (FabZ), (iii) an enoyl-ACP reductase (FabI) and (iv) FabD again. Three cycles are required for the synthesis of octanoic acid (C8:0), six for myristic acid (C14:0), and seven for palmitic acid (C16:0). Metabolic labeling analyses suggest that the main products of FASII are octanoic acid (C8:0), myristic acid (C14:0), and palmitic acid (C16:0) [34]. Very interestingly, it has recently been shown that the Plasmodium falciparum apicoplast PK was capable of generating a broad range of nucleotide triphosphate, which might explain its importance during blood stages (new31). Interestingly, Niu et al. (new32) showed that the apicoplast PKII could be dispensable and not the source of energy of the apicoplast, unlike Plasmodium falciparum, but rather the apicoplast triose phosphate isomerase and the GAPDH [24]. The fate of those apicoplast neosynthesized FA has only been unraveled in T. gondii. Indeed, the neosynthesized lipoic acid from octanoic acid was only the characterized product from the FASII pathway. The apicoplast lipoic acid is essential for the modification

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(lipoylation) of the apicoplast PDH and, therefore, crucial for its function. Nevertheless, it seemed counterintuitive that the FASII pathway would only be useful to generate substrates for its own function such as lipoic acid. Furthermore, based on the analysis of FASII KOs, the pathway is essential for apicoplast biogenesis in late liver stage development and in T. gondii, as well as for the formation of plasma membrane and nucleus in the Plasmodium late liver stage. This suggested that these FA could be incorporated into membrane lipids and participate in membrane biogenesis, likely for the apicoplast membranes. By genome analysis, two acyltransferases, homologs to the chloroplast enzymes ATS1 and ATS2, have been predicted to be in the apicoplast. These enzymes catalyze the stepwise esterification of FA onto a glycerol-3-phosphate (G3P) backbone to form phosphatidic acid (PA). PA is the sole precursor for the neosynthesis of all phosphoglycerolipids, the major membrane lipids in Plasmodium. The apicoplast is also capable of synthesizing the G3P backbone of PA via a predicted glceraldehyde-3phosphate dehydrogenase (GapDH). This reaction requires the import of DHAP by the TPTs, which is then converted into glyceraldehyde-3-phosphate, the substrate of GapDH by a triose phosphate isomerase. Recent analyses, using inducible knockdowns of ATS1 combined with stable isotope labeling and mass spectrometry analysis, proved that the apicoplast used the FASII FA to assemble them into essential precursors, i.e., lysophosphatidic acid (LPA) [36, 37]. Specifically, ATS1 uses the major product of the FASII: myristic acid (C14:0) to make LPA (14:0), which is then used as a central precursor for the bulk phospholipid synthesis in both parasites. ATS1 then allows the proper intracellular organelle biogenesis and division during T. gondii tachyzoite and Plasmodium liver stage [36–38]. In the meantime, the apicoplast still exports some free FA for another lipid synthesis. The final destination for either FA or PA is the neighboring endoplasmic reticulum, where the parasite membrane lipid synthesis machinery resides. The role of the apicoplast ATS2 is yet to be determined. The apicoplast is also predicted to harbor part of the required machinery for FA and PA export to extra-plastidial compartments. However, direct evidence on the presence of putative transporters/cargo proteins for these lipids is extremely scarce, advocating for the search of likely metabolic transporters and exporters of the apicoplast, in order to fully grasp its function as a central metabolite provider for the parasite [1].

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Conclusion Apicoplast or the secondary plastid of the malaria parasite is an essential organelle for parasite survival throughout the parasite life cycle. It hosts important pathways that provide crucial metabolites

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for diverse functions of the parasitic cell. Its algal origin thus makes it an attractive target for the development of new drugs for treatments of malaria-infected patients, for prophylactic drugs preventing the spread of the parasite. Some apicoplast KO mutants, such as for the FASII and the heme pathways, are even considered potential vaccine candidates. Indeed, these mutant lines can be used as genetically attenuated strains capable of infecting the liver cells and generating an immune response but arresting their development at this asymptomatic stage. However, the exact role of the apicoplast is not fully understood and many questions remain on the cellular fate of its neosynthesized metabolites and the trafficking of the metabolites between the apicoplast and the other cellular compartments. Novel approaches combining conditional knockdowns and metabolomics have started highlighting new facets of the apicoplast metabolic pathways and their contribution to the parasite, which still remains to be fully understood.

Acknowledgments This work and their authors were supported by Agence Nationale de la Recherche, France (Project ApicoLipiAdapt grant ANR-21CE44-0010; Project Apicolipidtraffic grant ANR-23-CE15-000901), The Fondation pour la Recherche Me´dicale (FRM EQU202103012700), Laboratoire d’Excellence Parafrap, France (grant ANR-11-LABX-0024), LIA-IRP CNRS Program (Apicolipid project), the Universite´ Grenoble Alpes (IDEX ISP Apicolipid) and Re´gion Auvergne Rhone-Alpes for the lipidomics analyses platform (Grant IRICE Project GEMELI), Collaborative Research Program Grant CEFIPRA (Project 6003-1) by the CEFIPRA (MESRI-DBT). References 1. Kloehn J, Lacour CE, Soldati-Favre D (2021) The metabolic pathways and transporters of the plastid organelle in Apicomplexa. Curr Opin Microbiol 63:250–258 2. He CY, Shaw MK, Pletcher CH et al (2001) A plastid segregation defect in the protozoan parasite Toxoplasma gondii. EMBO J 20(3): 330–339 3. Botte´ CY, Dubar F, McFadden GI et al (2012) Plasmodium falciparum apicoplast drugs: targets or off-targets? Chem Rev 112(3): 1269–1283 4. Kohler S, Delwiche CF, Denny PW et al (1997) A plastid of probable green algal origin in Apicomplexan parasites. Science 275(5305):1485

5. McFadden GI, Reith ME, Munholland J et al (1996) Plastid in human parasites. Nature 381(6582):482 6. Wilson RJ, Denny PW, Preiser PR et al (1996) Complete gene map of the plastid-like DNA of the malaria parasite Plasmodium falciparum. J Mol Biol 261(2):155–172 7. Janouskovec J, Horak A, Obornik M et al (2010) A common red algal origin of the apicomplexan, dinoflagellate, and heterokont plastids. Proc Natl Acad Sci USA 107(24): 10949 8. Zhu G, Marchewska MJ, Keithly JS (2000) Cryptosporidium parvum appears to lack a plastid genome. Microbiology 146(2):315

Roles and Maintenance of the Apicoplast 9. Fichera ME, Roos DS (1997) A plastid organelle as a drug target in apicomplexan parasites. Nature 390(6658):407–409 10. Botte CY, Yamaryo-Botte Y, Rupasinghe TWT et al (2013) Atypical lipid composition in the purified relict plastid (apicoplast) of malaria parasites. Proc Natl Acad Sci USA 110(18): 7506 11. Ralph SA, van Dooren GG, Waller RF et al (2004) Tropical infectious diseases: metabolic maps and functions of the Plasmodium falciparum apicoplast. Nat Rev Microbiol 2(3): 203–216 12. Boucher MJ, Yeh E (2019) Plastidendomembrane connections in apicomplexan parasites. PLoS Pathog 15(6):e1007661 13. Gould SB, Waller RF, McFadden GI (2008) Plastid evolution. Annu Rev Plant Biol 59: 491–517 14. Tawk L, Dubremetz JF, Montcourrier P et al Phosphatidylinositol (2011) 3-monophosphate is involved in toxoplasma apicoplast biogenesis. PLoS Pathog 7(2): e1001286 15. van Dooren GG, Kennedy AT, McFadden GI (2012) The use and abuse of heme in apicomplexan parasites. Antioxid Redox Signal 17(4): 634–656 16. Kloehn J, Harding CR, Soldati-Favre D (2021) Supply and demand—heme synthesis, salvage and utilization by Apicomplexa. FEBS J 288(2):382–404 17. Nagaraj VA, Sundaram B, Varadarajan NM et al (2013) Malaria parasite-synthesized heme is essential in the mosquito and liver stages and complements host heme in the blood stages of infection. PLoS Pathog 9(8):e1003522 18. Renaud EA, Pamukcu S, Cerutti A et al (2022) Disrupting the plastidic iron-sulfur cluster biogenesis pathway in Toxoplasma gondii has pleiotropic effects irreversibly impacting parasite viability. J Biol Chem 298(8):102243 19. Balk J, Pilon M (2011) Ancient and essential: the assembly of iron-sulfur clusters in plants. Trends Plant Sci 16(4):218–226 20. Haussig JM, Matuschewski K, Kooij TWA (2013) Experimental genetics of Plasmodium berghei NFU in the apicoplast iron-sulfur cluster biogenesis pathway. PLoS One 8(6): e67269 21. Swift RP, Elahi R, Rajaram K et al (2023) The Plasmodium falciparum apicoplast cysteine desulfurase provides sulfur for both iron-sulfur cluster assembly and tRNA modification. elife 12:e84491. https://doi.org/10.7554/eLife. 84491

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22. Yeh E, DeRisi JL (2011) Chemical rescue of malaria parasites lacking an apicoplast defines organelle function in blood-stage plasmodium falciparum. PLoS Biol 9(8):e1001138 23. Sidik SM, Huet D, Ganesan SM et al (2016) A genome-wide CRISPR screen in toxoplasma identifies essential apicomplexan genes. Cell 166(6):1423–1435.e12. https://doi.org/10. 1016/j.cell.2016.08.019. Epub 2016 Sep 2 24. Swift RP, Rajaram K, Keutcha C et al (2020) The NTP generating activity of pyruvate kinase II is critical for apicoplast maintenance in Plasmodium falciparum. elife 9:e50807. https:// doi.org/10.7554/eLife.50807 25. Niu Z, Ye S, Liu J et al (2022) Two apicoplast dwelling glycolytic enzymes provide key substrates for metabolic pathways in the apicoplast and are critical for toxoplasma growth. PLoS Pathog 18(11):e1011009. https://doi.org/ 10.1371/journal.ppat.1011009. PMID: 36449552; PMCID: PMC9744290 26. Mombo-Ngoma G, Remppis J, Sievers M et al (2018) Efficacy and safety of Fosmidomycin– Piperaquine as Nonartemisinin-based combination therapy for uncomplicated Falciparum malaria: a single-arm, age De-escalation proofof-concept study in gabon. Clin Infect Dis 66(12):1823–1830 27. Okada M, Rajaram K, Swift RP et al (2022) Critical role for isoprenoids in apicoplast biogenesis by malaria parasites. elife 11:e73208 28. Suazo KF, Schaber C, Palsuledesai CC et al (2016) Global proteomic analysis of prenylated proteins in Plasmodium falciparum using an alkyne-modified isoprenoid analogue. Sci Rep 6:38615 29. Gisselberg JE, Zhang L, Elias JE et al (2017) The prenylated proteome of Plasmodium falciparum reveals pathogen-specific prenylation activity and drug mechanism-of-action. Mol Cell Proteomics 16(4):S54–S64 30. MacRae JI, Mare´chal E, Biot C et al (2012) The apicoplast: a key target to cure malaria. Curr Pharm Des 18(24):3490–3504 31. Yu M, Kumar TRS, Nkrumah LJ et al (2008) The fatty acid biosynthesis enzyme FabI plays a key role in the development of liver-stage malarial parasites. Cell Host Microbe 4(6): 567–578 32. Vaughan AM, O’Neill MT, Tarun AS et al (2009) Type II fatty acid synthesis is essential only for malaria parasite late liver stage development. Cell Microbiol 11(3):506 33. Amiar S, Katris NJ, Berry L et al (2020) Division and adaptation to host environment of apicomplexan parasites depend on apicoplast

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lipid metabolic plasticity and host organelle remodeling. Cell Rep 30(11):3778–3792.e9 34. Shunmugam S, Arnold CS, Dass S et al (2022) The flexibility of Apicomplexa parasites in lipid metabolism. PLOS Pathog 18(3):e1010313 35. Dass S, Shunmugam S, Berry L et al (2021) Toxoplasma LIPIN is essential in channeling host lipid fluxes through membrane biogenesis and lipid storage. Nat Commun 12(1):2813 36. Amiar S, MacRae JI, Callahan DL et al (2016) Apicoplast-localized lysophosphatidic acid precursor assembly is required for bulk phospholipid synthesis in Toxoplasma gondii and relies

on an algal/plant-like glycerol 3-phosphate acyltransferase. PLoS Pathog 12(8):e1005765 37. Shears MJ, MacRae JI, Mollard V et al (2017) Characterization of the Plasmodium falciparum and P. berghei glycerol 3-phosphate acyltransferase involved in FASII fatty acid utilization in the malaria parasite apicoplast. Cell Microbiol 19(1):e12633 38. Lindner SE, Sartain MJ, Hayes K et al (2014) Enzymes involved in plastid-targeted phosphatidic acid synthesis are essential for Plasmodium yoelii liver-stage development. Mol Microbiol 91(4):679

Chapter 4 Diversification of Plastid Structure and Function in Land Plants Henrik Aronsson and Katalin Solymosi Abstract Plastids represent a largely diverse group of organelles in plant and algal cells that have several common features but also a broad spectrum of morphological, ultrastructural, biochemical, and physiological differences. Plastids and their structural and metabolic diversity significantly contribute to the functionality and developmental flexibility of the plant body throughout its lifetime. In addition to the multiple roles of given plastid types, this diversity is accomplished in some cases by interconversions between different plastids as a consequence of developmental and environmental signals that regulate plastid differentiation and specialization. In addition to basic plastid structural features, the most important plastid types, the newly characterized peculiar plastids, and future perspectives in plastid biology are also provided in this chapter. Key words Proplastid, Etioplast, Chloroplast, Chromoplast, Leucoplast

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Introduction It was probably Anthony van Leeuwenhoek who first observed a plastid (i.e., the large, spiral chloroplast of a green alga, Spirogyra, that recalled him the copper or tin worms of distillers) in his microscope in 1674 [1]. The term plastid (from the Greek word “plastikos”, meaning formed or molded) was coined in 1883 by Andreas Franz Wilhelm Schimper in order to refer to the plasticity of this organelle, which includes its ability to transform from one kind of plastid into another one and was based on observations about the differentiation of small colorless plastids of meristems into large chloroplasts or chromoplasts in mature tissues (Schimper, 1883, cf. Gunning et al., 2006). Plastids cannot be formed de novo in the cytoplasm and are therefore inherited by the offspring and have a developmental continuity within the organism. As a consequence, the role(s) and metabolism and thus the structure of plastids may vary along with the differentiation of the organs,

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tissues, and specific cells of the plant body during the entire life cycle as part of the developmental program of the host cell that harbors them, but plastid differentiation is also strongly influenced by changes in the environmental conditions. Thus, intermediary plastid forms also exist and plastids with similar ultrastructure may have quite different metabolic activity. Plastids can be traced back to the event in the evolution (1.2–1.5 billion years ago) when an ancient cyanobacterium, considered as a photosynthetic free-living single prokaryotic cell, was engulfed and retained by a eukaryote, via a process named primary endosymbiosis [2–4]. The primary endosymbiosis gave rise to a primary plastid believed to be the common ancestor to all primary plastids found in the Archaeplastida, i.e., plastids in green algae and plants, in red algae (sometimes termed rhodoplasts), and in glaucophytes (also termed cyanelles or muroplasts) [3–5]. Photosynthetic eukaryotes with primary plastids were engulfed again resulting in secondary endosymbiosis, and organisms with plastids obtained by secondary endosymbiogenesis could be also absorbed by other eukaryotes by tertiary or serial endosymbiosis leading to a great diversification of photosynthetic organisms [4, 6, 7]. The engulfed free-living cyanobacterium originally had a large genome (more than 6 Mb) [8] as an endosymbiont; however, a large part of its genome was later transferred to the host genome (nucleus) or lost during the endosymbiogenesis process when it finally became a semi-autonomous organelle [9]. Mitochondria evolved similarly through gene transfer processes, the reduction of genome size, and nucleus-driven control of the organellar activity [10]. The chloroplast of most land plants has today around 110–190 kb in its plastome, although extremes exist as well [11– 13]. Although most of the original cyanobacterial endosymbiont’s genome (ca 95%) has been lost, the plastid evolved still has around 100 μm) [1] and recently has been performed in two model diatoms Thalassiosira pseudonana and Phaeodactylum tricornutum [2, 3]. Here we provide an improved isolation protocol for the diatom P. tricornutum. The integrity and purity of isolated plastids are verified by fluorescence images and western blot analysis.

2

Materials Prepare all solutions with analytical grade reagents and deionized water (electrical resistivity of 18 MΩ-cm at 25 °C), as described in Schober et al. (2018) [3].

2.1 Strain and Culture Medium

1. Phaeodactylum tricornutum CCMP2561, a gift from Prof. Chris Bowler (E´cole Normale Supe´rieure, Paris, France), is obtained from the culture collection of the Provasoli-Guillard National Center for Culture of Marine Phytoplankton, Bigelow Laboratory for Ocean Sciences (USA). 2. Grow algal cells in artificial seawater enriched with f/2 (nitrate concentration is increased to 4 times) [4]. 3. Sterilize liquid medium and 5-cm diameter glass culture tubes (60-cm long) by autoclaving, and then let the solution cool down to room temperature prior to cell culture.

2.2

Stock Solutions

Prepare the stock solution in advance, sterilize it by filtration through a filter membrane (0.2 μm pore size), and store it at 4 °C. It is better to use it up within two months. Prepare all the stock solutions according to Schober et al. (2018) [3]. 1. 2 M Sorbitol stock solution: Weigh 72.87 g Sorbitol and put it in a bottle. Add about 100 mL deionized water to the bottle and stir until the Sorbitol has dissolved completely. Transfer the solution to a measuring cylinder, and bring up the volume to 200 mL with deionized water. 2. 1 M HEPES-KOH stock solution, pH 8: Dissolve 47.67 g 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES) in 150 mL of deionized water, adjust the pH to 8 by slowly adding KOH, and bring up the volume to 200 mL with deionized water.

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3. 0.5 M EDTA stock solution, pH 8: Weigh 146.12 g EDTA and dissolve it in 300 mL of deionized water. Add KOH to adjust the solution pH to 8.0, and bring up the volume to 500 mL with deionized water. 4. 1 M MgCl2 stock solution: Dissolve 10.17 g MgCl2·6H2O in 40 mL deionized water. When dissolved completely, bring up to 50 mL. 5. 1 M KCl stock solution: Dissolve 3.73 g KCl in 40 mL deionized water. When dissolved completely, bring up to 50 mL. 6. 1 M MnCl2 stock solution: Dissolve 9.9 g MnCl2·4H2O in 40 mL deionized water. When dissolved completely, bring up to 50 mL. 7. 20 mM HEPES-KOH stock solution, pH 7.2: Dissolve 0.95 g HEPES in 150 mL deionized water. Adjust the pH to 7.2 by adding KOH, and then bring up to 200 mL. 2.3 Working Solutions

It is advisable to prepare working solutions at least one day before the actual isolation procedure and store them at 4 °C. Prepare all the working solutions according to Schober et al. (2018) [3]. 1. Isolation buffer: 0.5 M Sorbitol, 50 mM HEPES-KOH, 6 mM EDTA, 5 mM MgCl2, 10 mM KCl, 1 mM MnCl2, 1% (w/v) Polyvinylpyrrolidone 40 (PVP-40), 0.5% bovine serum albumin (BSA, defatted), 0.1% L-cysteine; pH 7.2–7.5. Weigh 1.0 g of PVP-40 and transfer it to a bottle. Add the following sterile substances while working under the clean bench: 25 mL 2 M Sorbitol stock solution, 5 mL 1 M HEPES-KOH stock solution pH 8, 1.2 mL 0.5 M EDTA stock solution pH 8, 0.5 mL 1 M MgCl2 stock solution, 1 mL 1 M KCl stock solution, 100 μL 1 M MnCl2 stock solution. Bring up to ~90 mL with deionized water. Lower the pH to 7.2–7.5 with HCl while stirring (see Note 1). Check the osmolality of the working solution using a calibrated Osmometer. The value should by default range between 700 and 750 mOsm/kg (see Note 2). If the osmolality is lower than 730–750 mOsm/kg, raise it by stepwise addition of small volumes of 2 M Sorbitol stock solution. Once the osmolality is adjusted, fill up to 100 mL with deionized water. On the day of plastid isolation, add 0.5 g BSA (defatted) and 0.1 g L-cysteine to 100 mL isolation buffer and precool it on ice. 2. Washing buffer: 0.5 M Sorbitol, 30 mM HEPES-KOH, 6 mM EDTA, 5 mM MgCl2, 10 mM KCl, 1 mM MnCl2, 1% (w/v) PVP-40; pH 7.2–7.5. Weigh 0.5 g of PVP-40 and transfer it to a 50 mL centrifuge tube. Add the following sterile substances

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while working under the clean bench: 12.5 mL 2 M Sorbitol stock solution, 1.5 mL 1 M HEPES-KOH stock solution pH 8, 0.6 mL 0.5 M EDTA stock solution pH 8, 0.25 mL 1 M MgCl2 stock solution, 0.5 mL 1 M KCl stock solution, and 50 μL 1 M MnCl2 stock solution. Bring up to ~45 mL with deionized water. Lower the pH to 7.2–7.5 with HCl while stirring (see Note 3). Check the osmolality of the working solution using a calibrated Osmometer. The value should by default range between 700 and 750 mOsm/kg (see Note 2). If the osmolality is lower than 730–750 mOsm/kg, raise it by stepwise addition of small volumes of 2 M Sorbitol stock solution. Bring up the volume to ~50 mL with deionized water. On the day of plastid isolation, divide the washing buffer evenly to two of 25 mL. Add 0.125 g BSA (defatted) to one of the two and precool both on ice. 3. 10%-, 20%- and 30%-Percoll Solution: 0.5 M Sorbitol, 30 mM HEPES-KOH, 6 mM EDTA, 5 mM MgCl2, 10 mM KCl, 1 mM MnCl2, 0.1% (w/v) PVP-40, 10%, 20% or 30% (v/v) Percoll; pH 7.2–7.5. Weigh 0.05 g of PVP-40 and transfer it to a 50 mL centrifuge tube. Add the following sterile substances while working under the clean bench: 12.5 mL 2 M Sorbitol stock solution, 1.5 mL 1 M HEPES-KOH stock solution pH 8, 0.6 mL 0.5 M EDTA stock solution pH 8, 0.25 mL 1 M MgCl2 stock solution, 0.5 mL 1 M KCl stock solution, 50 μL 1 M MnCl2 stock solution. Add 29.6 mL (for 10% Percoll), 24.6 mL (for 20% Percoll), or 19.6 mL (for 30% Percoll) deionized water. Lower the pH to 7.2–7.5 with HCl while stirring (see Note 3). Check the osmolality of the working solution using a calibrated Osmometer. If the osmolality is lower than 835 (±20), 935 (±20), or 1070 (±20) mOsm/kg for 10%-Percoll, 20%-Percoll, and 30%-Percoll solution, respectively (see Note 2), raise it carefully by stepwise addition of small volumes of 2 M Sorbitol stock solution. Once the osmolality is adjusted correctly, sterilize the solution by autoclaving. Then let the solution cool down to room temperature and add 5 mL (=10% v/v), 10 mL (=20% v/v), or 15 mL (=30% v/v) Percoll (sterile) under the clean bench and mix gently. 2.4 Main Laboratory Apparatus

1. Cryoscopic osmometer Osmomat 030 (Gonotec, Germany) 2. Ultracentrifuge Optima XPN-100 (Beckman, USA) with a SW41Ti-0765 rotor 3. Low Temperature Ultra-high Pressure Continuous Flow Cell Disrupter JN-02C (JNBIO, China)

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Methods Keep all working solutions, cells, and the isolated plastids on ice and carry out all steps at 4 °C, unless specified differently. Whenever possible, avoid exposition of crude extracts and plastids to light.

3.1 Cell Culture and Harvest

1. Incubate mid-exponential phase cells in glass culture tubes containing 600 mL of liquid medium for 5 days at 22 °C and 100 μmol photons m-2 s-1 with constant aeration. 2. Centrifuge cells after 4–5 days (see Note 4) of cultivation (to a concentration of 2–3 × 107 cells mL-1) at 2000 g for 10 min, 4 °C in a 50 mL centrifuge tube. Wash them twice with 30 mL of 20 mM HEPES-KOH stock solution (pH 7.2). Resuspend the resulting pellet gently, but thoroughly, in 40 mL of isolation buffer (see Note 5).

3.2 Cell Rupture and Plastid Isolation

1. Load the resulting cell solution to JN-02C Cell disrupter at 75–80 MPa. 2. Collect the eluate in a chilled falcon tube wrapped in aluminum foil to keep it dark. 3. Centrifuge the solution of broken cells at 300 g for 8 min (see Note 6). Collect the supernatant for centrifugation at 2000 g for 5 min. This step sediments the plastids. Resuspend the sediment cautiously with 10 mL of washing buffer (with BSA, defatted).

3.3 Purification of Plastids on a Discontinuous Percoll Gradient

1. Preparation of Percoll Gradient: Draw 3 mL of the 30%-, 3 mL of the 20%-, and 3 mL of the 10% Percoll solution to a thinwalled ultraclear centrifuge tube to gently layer discontinuous gradients from bottom to top. 2. Slowly pipette 2 mL of the resuspended plastid solution on one layered Percoll gradient. 3. Centrifuge the gradients for 60 min at 14400 g.

3.4 Extraction and Washing of Plastids

1. The plastid-enriched fraction accumulates at the interface between the 10% and 20% Percoll layers (see Note 7), as indicated by the red circle (see Fig. 1a). Carefully collect the highquality plastid fraction and transfer it into a new tube. 2. Fill up the tube with washing buffer (without BSA), and wash the plastids by centrifugation at 4000 g for 10 min. 3. Discard the supernatant and resuspend the pellet with washing buffer (without BSA) for centrifugation at 4000 g for 10 min. 4. Collect the pellet (the purified plastids) for protein extraction (see Note 8) or observation by microscope (see Note 9).

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Fig. 1 Analysis of composition and integrity of purified plastids from Phaeodactylum tricornutum. (a) Isolation workflow and position in Percoll gradient (indicated by a red circle) of plastids. (b) Fluorescence micrographs of isolated plastids

4 Notes 1. Usually, the pH value of the solution is about 7.4, so there is no need to adjust it. 2. Add 2 M Sorbitol stock solution slowly to adjust the osmolality of the solution. Every additional 500 μL 2 M Sorbitol stock solution raises the osmolality of the solution by about 20 mOsm/kg. 3. Usually, the pH value of the solution is about 7.35, so there is no need to adjust it 4. Ensure that exponential phase cells are used, and observe the cells under a microscope. 5. Make sure that the cell concentration of the resuspended solution is reasonably low prior to cell rupture. 6. Repeat the step if necessary. 7. The accumulation of plastid-enriched fraction may occur between different Percoll layers, depending on the Percoll provided by manufacturers and the cultivation condition of cells. 8. In order to determine the purity of the isolated plastids, plastid- (β-carbonic anhydrase: PtCA1 and PtCA2; PEPC1: periplastidial compartment localized phosphoenolpyruvate carboxylase 1), cytosol- (β-actin), and other organelles(nuclear protein AGO: argonaute protein; mitochondrial proteins: ME1, NAD-dependent malic enzyme; PEPC2; SHMT2, serine hydroxymethyltransferase 2; coatomer subunit alpha, COP1) specific antibodies are used for western blot analysis (see Fig. 2).

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Fig. 2 Western blot analysis of purified plastid. (a) Coomassie brilliant blue staining of whole cell and plastid protein extracts. (b) Analysis of protein extracts using antibodies against plastid proteins. (c) Analysis of protein extracts using antibodies against non-plastid proteins

9. Isolated plastids exhibited the same elongated shape as that observed in vivo; however, they tended to condense into rounded structures 2 h after the isolation (see Fig. 1b). References 1. Wittpoth C, Kroth PG, Weyrauch K et al (1998) Functional characterization of isolated plastids from two marine diatoms. Planta 206(1): 7 9 – 8 5 . h t t p s : // d o i . o r g / 1 0 . 1 0 0 7 / s004250050376 2. Schober AF, Rı´o Ba´rtulos C, Bischoff A, Lepetit B, Gruber A, Kroth PG (2019) Organelle studies and proteome analyses of mitochondria and plastids fractions from the diatom Thalassiosira pseudonana. Plant Cell Physiol 60(8):1811–1828. https://doi.org/10.1093/ pcp/pcz097

3. Schober AF, Flori S, Finazzi G, Kroth PG, Rı´o Ba´rtulos C (2018) Isolation of plastid fractions from the diatoms Thalassiosira pseudonana and Phaeodactylum tricornutum. In: Mare´chal E (ed) Plastids: methods and protocols. Springer, New York, pp 189–203 4. Guillard RL (1975) Culture of phytoplankton for feeding marine invertebrates. In: Smith W, Chanley M (eds) Culture of marine invertebrate animals. Springer US, pp 29–60. https://doi. org/10.1007/978-1-4615-8714-9_3

Chapter 11 Determining the Subcellular Localization of Proteins in the Different Membranes of Diatom Secondary Plastid Xiaojuan Liu and Yangmin Gong Abstract Diatoms such as Phaeodactylum tricornutum arose through a process termed secondary endosymbiosis, in which red alga–derived plastids are surrounded by a complicated membrane system. Subcellular marker proteins provide defined localizations on the compartmental and even sub-compartmental levels in the complex plastids of diatoms. Here we introduce how to use subcellular marker proteins and in vivo co-localization in the diatom P. tricornutum by presenting a step-by-step method allowing the determination of subcellular localization of proteins in different membranes of the secondary plastid. This chapter describes the materials required and the procedures of transformation and microscopic observation. Key words Secondary plastid, Co-localization, GFP, Phaeodactylum tricornutum, mRFP

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Introduction Plastids are the characteristic organelles of photosynthetic organisms and have a central position in oxygenic photosynthesis and primary metabolism. Land plants and algae have the primary plastids, which are evolved by engulfment and successive reduction of a cyanobacterial-like cell within an early eukaryote. Plastids of this origin are surrounded by two membranes. Different from the primary plastids, the secondary plastids of some diatoms and other heterokontophytes are originated by engulfment of a phototrophic eukaryote with a primary plastid by another eukaryotic cell, and they, thus, have three or four envelope membranes [1]. In diatoms, the outermost of the four envelope membranes might trace back to a phagotrophic membrane and appears to be continuous with the host endoplasmic reticulum (ER) system (cER, chloroplast ER). The second outermost membrane (periplastidal membrane, PPM) is probably derived from the plasma membrane of the eukaryotic endosymbiont (Fig. 1), and both innermost membranes are homologous to the two envelope membranes of the

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I. II. III. IV.

Chloroplast ER membrane cER Periplastidal membrane

PPC

Plastid outer envelope membrane

IMS

Plastid inner envelope membrane

Stroma

Fig. 1 Different membranes and compartments in the complex plastid of the diatom Phaeodactylum tricornutum. The plastid is surrounded by four membranes, and it is connected with the ER of the host cell. cER, chloroplast endoplasmatic reticulum; PPC, periplastidal compartment; IMS, inner membrane space

primary chloroplast [2, 3]. Although some plastid proteins are encoded by the plastid genome, most plastid proteins are encoded by the host nuclear genome and have to be transported across different plastid membranes in diatoms. Nuclear-encoded plastid proteins usually harbor a bipartite targeting sequence (BTS) at the N-terminus consisting of a signal peptide and a transit peptide. Within the signal peptide cleavage site, a conserved motif, namely ASAF or AFAP, is present in preproteins targeted into secondary plastids of diatoms [4]. Some important biochemical processes, including photosynthesis and de novo fatty acid synthesis, take place in the plastids of diatoms. Studying the biological function of a protein in diatom often requires the determination of its subcellular localization. Localization of diatom protein is complicated, especially for those proteins in the complex plastids. Currently, subcellular localization can be done immunohistochemically using transmission electron microscopy (TEM), and this method involves the preparation of cell samples, immunogold labeling, and the use of TEM, which requires advanced skills and experience, and thus, it is not often applied for most laboratories. In most cases, subcellular localization can be determined by expression of a C-terminal or N-terminal GFP (Green Fluorescence Protein)-fusion protein in P. tricornutum. This method requires the construction of GFP fusion plasmids, genetic transformation, and observation of

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Table 1 P. tricornutum marker proteins localized in the different membranes of plastid Compartment or organelle

Name

Protein ID

Predicted TS

cERM

Sec61

Phatr3_J41031 VPAF

Protein transport across the outermost membrane

PPM

ptE3P

Phatr3_J48034 KFAS

The ubiquitin ligase

OEM

Omp85 Phatr3_J46881 AFLP

Precursor protein translocation

IEM

Tic20

Protein import across the IEM

Phatr3_J45335 AFAP

Functional annotation

cERM, chloroplast ER membrane, PPM, periplastidal membrane, OEM, plastid outer envelope membrane, IEM, plastid inner envelope membrane

fluorescence signal within the cells of transformants using laserscanning confocal microscopy and has become the prevalent method for in vivo protein localization studies in diatoms. P. tricornutum contains a secondary plastid having plastid autofluorescence (PAF), which is detectable between 625 and 720 nm [5, 6]. The emission of enhanced version of GFP (eGFP) is detectable at 500–520 nm, and eGFP is a bright and photostable fluorescent protein. Thus, it has been extensively applied for subcellular localization studies in the diatom P. tricornutum. For determining the localization of proteins in the complex plastid of diatoms, it is very helpful to conduct co-localization studies using a second fluorescent protein fused to a protein marker localized to a certain compartment or subcompartment. Recently, the optimized mRuby3 was reported to be a second fluorescent protein suitable for in vivo co-localization studies [7], as its emission is detectable between 580 and 605 nm, which can be clearly discriminable from the emissions of eGFP and PAF. mRuby3 can be fused to the N-terminal or C-terminal of different marker proteins, including Sec61 localized in the cER membrane, ptE3P localized in the PPM, Omp85 localized in the plastid outer envelope membrane (OEM), and Tic20 localized in the plastid inner envelope membrane (IEM) (Table 1). In this chapter, we describe the detailed protocol for in vivo co-localization for proteins in the different membranes of plastids by simultaneous expression of eGFP and mRuby3 fused to marker protein in the diatom P. tricornutum.

2

Materials

2.1 Strains and Media

1. Escherichia coli DH5α, or similar. 2. P. tricornutum algal strain of choice (e.g., Culture Collection of Algae and Protozoa CCAP 1055/1).

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3. 100 μg/mL ampicillin LB plates: 10 g.L-1 (1%) Tryptone, 5 g. L-1 (0.5%) Yeast Extract, 10 g.L-1 (1%) NaCl, 15 g.L-1 (1.5%) Agar, 100 mg.L-1 Ampicillin. 4. Modified ESAW (Enriched Seawater, Artificial Water) medium: 362.7 mM NaCl; 25 mM Na2SO4; 8.03 mM KCl; 2.067 mM NaHCO3; 0.725 mM KBr; 0.372 mM H3BO3; 0.0657 mM NaF; 47.18 mM MgCl2; 9.134 mM CaCl2; 0.082 mM SrCl2; 21.8 μM Na2-glycerophosphate; 105.6 μM Na2SiO3; 14.86 μM disodium ethylenediaminetetraacetate dehydrate (Na2EDTA); 5.97 Fe(NH4)2(SO4)2 μM; FeCl3 0.592 μM; 2.42 μM MnSO4; 0.254 μM ZnSO4; 0.0569 CoSO4 μM; 0.52 μM Na2MoO4; 61.46 μM H3BO3; 10 nM Na2SeO3; 8.18 nM biotin (vitamin H); 2.94 nM cobalamin (vitamin B12); 0.594 μM thiamine (vitamin B1); and 5.49 mM NaNO3 and 0.224 mM NaH3PO4. 5. Stock solutions of pure paraformaldehyde and glutaraldehyde. 6. Stock solution of phosphate buffered saline buffer × 10: 1.37 M NaCl, 27 mM KCl, 100 mM Na2HPO4, and 18 mM KH2PO4. 2.2 Molecular Biology Reagents

1. Mighty TA-cloning Kit (Takara). 2. pPha-DUAL[2×NR]/pPha-2×NR (GenBank: JN180664). 3. pPha-mRuby3 (this work). 4. Phusion High-Fidelity PCR Master Mix (Thermo Scientific). 5. GeneJET Plasmid Miniprep Kit (Thermo Scientific). 6. MiniBEST Agarose Gel DNA Extraction Kit (Takara). 7. In-Fusion® Snap Assembly Master Mix (Takara). 8. RNeasy Mini Kit (Qiagen). 9. QuantiTect Reverse Transcription Kit (Qiagen). 10. NotI, EcoRI, SpeI, SacII, or other restriction enzyme. 11. Design primers for PCR amplification of genes encoding marker protein and mRuby3 using the online Primer Design tool (https://www.takarabio.com/learning-centers/cloning/ primer-design-and-other-tools) according to the In-Fusion Snap Assembly User Manual. In-Fusion allows one to join two or more DNA fragments (e.g., vector and insert or multiple fragments) as long as they share 15 bases of homology at each end. This homology is achieved through primer designed specifically for the experiment. Usually, the 5′ end of each primer must contain 15 bases that are homologous to 15 bases at one end of the DNA fragment to which it will be joined (i.e., the vector or another insert).

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12. Design primers for PCR amplification of genes encoding target protein and eGFP using designing methods, such as the online Primer Design tool according to the In-Fusion Snap Assembly User Manual. 2.3

Equipment

1. Micropipettes. 2. Micropipette tips. 3. Microcentrifuge tubes. 4. PCR tubes. 5. Petri dishes. 6. Thermocycler. 7. Microcentrifuge. 8. Incubator with shaking for liquid cultures. 9. Nanodrop or similar equipment for DNA quantification. 10. Access to Sanger sequencing services. 11. Confocal laser scanning microscope Leica TCS SP2. 12. Biolistic PDS-1000/He Particle Delivery System (Bio-Rad). 13. 1500 psi rupture discs. 14. Tungsten particles M17 (1.1 μm median diameter). 15. Water bath. 16. 2100 Bioanalyzer. 17. Clean bench. 18. Centrifuge. 19. Illumination incubator.

3

Methods

3.1 Construction of Plasmid Harboring Marker ProteinmRuby3 Fusion

1. Collect a total of 2 × 107 mid-exponentially growing algal cells from ESAW medium containing NaNO3 and NaH3PO4. Add RNA-protect cell reagent into cell solution with 1:1 volume ratio.

3.1.1 RNA Purification and cDNA Synthesis

2. Wash cell pellet with sterile distilled water twice to remove residual RNA-protect cell reagent and medium. 3. Suspend cell pellet in appropriate volume of lysis buffer supplied in Qiagen RNeasy Mini Kit. Add 250 μL acid-washed glass beads and mechanically disrupted cells using Mini-Beadbeater-16 Model 607. To avoid heat degradation of RNA, run beadbeating for 40 s and cooldown on ice for 1 min. Repeat this process until most cells are lysed. 4. From soluble fraction of above lysate, purify total RNA using Qiagen RNeasy Mini Kit. Aliquot 5–10 μL of purified RNA in

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RNase-free tube, and store at -80 °C until use. Check quantity and quality of purified RNA using Agilent’s Bioanalyzer. RNA samples with 260 nm/280 nm ratio between 1.8 and 2.0 will be used for the next experiments (see Note 1). 5. Synthesize cDNA using QuantiTect Reverse Transcription Kit. Measure the concentration of synthesized cDNA. Store cDNA at -20 °C until use. 3.1.2

PCR Amplification

1. Mix all the components according to Phusion High-Fidelity PCR Master Mix in PCR tube by vortexing vigorously. Mix the indicated volumes to reach the following concentrations in 20 μL final volume: x μL of cDNA/plasmid template, corresponding to 50 ng; 1 μL of Forward primer/reverse primers (from a stock solution of 10 μM each), corresponding to a final concentration of 0.5 μM; 10 μL of 2×Phusion Master Mix, 9-x μL of RNase/DNase-free water. Spin down at 300× g for 1 min to ensure solution settle at the bottom nice and even. To amplify gene encoding marker protein described in Table 1, use P. tricornutum cDNA as template. To amplify mRuby3 gene, use the plasmid harboring mRuby3 (e.g., pPha-mRuby3) as template (see Note 2). 2. Perform PCR on a thermal cycler with the following cycle: an initial step at 98 °C (30 s); 30 iterations at 98 °C (5–10 s), 60 °C (10–30 s), 72 °C (15–30 s per kilobase); a final step at 72 °C (5–10 min). 3. Spin-column purify PCR products of marker protein gene and mRuby3. 4. Set up the digestion reactions for the pPha-2×NR vector (Fig. 2) using the appropriate restriction enzyme (e.g., NotI) according to the multiple cloning site 1 of the pPha-2×NR vector as follows, and incubate for 60 min at 37 °C. Mix the indicated volumes to reach the following concentrations in 50 μL final volume: x μL pPha-2×NR, corresponding to 500 ng); 5 μL rCutSmart Buffer; 0.5 μL NotI-HF; 44.5-x μL of DNase-free water. 5. Run an electrophoresis gel to check the digestion results, and extract the desired fragments with the MiniBEST Agarose Gel DNA Extraction Kit to obtain the linearized pPha-2×NR vector.

3.1.3 Cloning of Marker Protein-mRuby3 Fusion into the pPha-2×NR Vector

1. Set up In-Fusion Snap Assembly cloning reaction by mixing the indicated volumes to reach the following concentrations in 10 μL final volume: 2 μL of 5×In-Fusion Snap Assembly Master Mix; x μL of linearized pPha-2×NR vector (NotI); y μL of marker protein gene (purified); z μL mRuby3 (purified); and 8-x-y-z μL of distilled water.

Localization of Plastid Membrane Proteins in Diatom (4270)

(4065)

(3946)

SspI

191

SpeI (452) NotI (459) SacII (470)

XmnI

ScaI MfeI (737)

(3865) TsoI (3836) PvuI

(3688)

(3506) (3466)

FspI

BmrI AhdI

EcoRI (1198) Eco53kI (1224) BanII - SacI (1226) Acc65I (1228) KpnI (1232) XbaI (1243) SalI (1249) SbfI (1259) SphI (1265) HindIII (1267)

pPha-DUAL[2xNR] 4635 bp

BspEI (1540)

(2989)

AlwNI

PvuII (1676) MscI (1735) MauBI (1768) SgrAI (1846) BmgBI (1866) SexAI* (1895) FseI (2008) (2457)

BbsI (2176) BspQI-SapI NcoI (2190) (2394) HpaI (2343) PaeR7I -TliI -XhoI BstAPI (2286)

Fig. 2 Plasmid map of the pPha-DUAL[2×NR] vector

2. Incubate the reaction for 15 min at 50 °C, then place on ice. 3. Transform E. coli DH5α competent cells with 2.5 μL of the above reaction mixture. Plate 200 μL of recovered cell culture on 100 μg/mL ampicillin LB plates. Incubate at 37 °C overnight. 4. Using the GeneJET Plasmid Miniprep Kit, extract plasmid DNA from the overnight cultures and elute plasmids from the column with 25 μL DNase-free water. Repeat this elution step once more (see Note 3). 5. Screen the plasmids by performing restriction digestion with restriction enzymes SpeI and SacII. The reaction mixture can be scaled down to 10 μL. 6. Run a DNA electrophoresis gel to screen the plasmids. Select plasmid samples that contain the correct fragment size. For positive clones, two fragments (~4.6 kb for the vector) should be obtained from pPha-2×NR_Marker-mRuby3 if the plasmids were digested with SpeI and SacII.

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3.2 Plasmid Construction for Target Protein-eGFP Fusion 3.2.1

PCR Amplification

1. Use the Phusion High-Fidelity PCR Master Mix to amplify target gene from P. tricornutum and eGFP, respectively. To amplify gene encoding full-length target protein, use P. tricornutum cDNA as template. To amplify eGFP gene, use the plasmid harboring eGFP (e.g., pPha-eGFP) as template. 2. Perform PCR as described in Subheading 3.1.2, then spincolumn purify PCR products of target protein gene and eGFP, respectively. Predict the presence of N-terminal targeting sequences of target protein (see Note 4). 3. Set up the digestion reactions for the plasmid of pPha2×NR_Marker-mRuby3 that is obtained from Subheading 3.1.3 using the appropriate restriction enzyme (e.g., EcoRI, KpnI, XbaI, SalI, HindIII) according to the multiple cloning site 2 of the pPha-2×NR vector. Mix the indicated volumes to reach the following concentrations in 50 μL final volume: x μL of pPha-2×NR_Marker-mRuby3, corresponding to 500 ng; 5 μL rCutSmart Buffer; 0.5 μL EcoRI-HF; 44.5-x μL DNasefree water. Incubate for 60 min at 37 °C. Choice of restriction enzyme should avoid cutting of marker protein gene and mRuby3 in the plasmid of pPha-2×NR_Marker-mRuby3. 4. Run an electrophoresis gel to check the digestion results and extract the desired fragments with the MiniBEST Agarose Gel DNA Extraction Kit to obtain the linearized pPha-2×NR_Marker-mRuby3.

3.2.2 Cloning of Target Protein-eGFP Fusion into the pPha-2×NR Vector

1. Set up In-Fusion Snap Assembly cloning reaction. Mix the indicated volumes to reach the following concentrations in 50 μL final volume: 2 μL 5×In-Fusion Snap Assembly Master Mix; x μL Linearized pPha-2×NR_Marker-mRuby3 (EcoRI), corresponding to 50–100 ng; y μL of the Target protein gene (purified); z μL of eGFP sequence (purified); 8-x-y-z μL of distilled water. 2. Incubate the reaction for 15 min at 50 °C, then place on ice. 3. Transform E. coli DH5α competent cells with 2.5 μL of the above reaction mixture. Plate 200 μL of recovered cell culture on 100 μg/mL ampicillin LB plates. Incubate at 37 °C overnight. 4. Screen the positive clones by performing colony PCR using two pairs of primers to amplify marker protein-mRuby3 and target protein-eGFP fusions, respectively. 5. Using the GeneJET Plasmid Miniprep Kit, extract plasmid DNA from the overnight cultures and elute plasmids from the column with 25 μL DNase-free water. Repeat this elution step once more.

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6. Digest the positive plasmids of pPha-2×NR_Marker-mRuby3_Targ-eGFP using appropriate restriction enzymes. Analyze results by agarose gel electrophoresis to verify the correct product size of digestion of the positive plasmids. 3.3 Transformation of P. tricornutum

The following protocol allows for efficient transformation of P. tricornutum with pPha_DUAL_2×NR vector-derived plasmids (see Note 5). 1. Tungsten particles M17 (1.1 μm median diameter) were coated with 0.8 μg of plasmid DNA in the presence of CaCl2 and spermidine. 2. Approximately 5 × 107 algal cells from the exponential phase (OD600 = 0.4~0.6) were spread, centrally covering one-third of a plate of solid F/2 medium 1 h prior to bombardment. 3. The plate was positioned at the second level within the Biolistic chamber for bombardment. 4. After bombardment, algal cells were illuminated for 24 h. 5. Cells were suspended in 0.5 mL of sterile F/2 medium. 6. 100 μL of cell suspension (~1 × 107 cells) was plated onto solid medium containing 100 μg/mL Zeocin. 7. The plates were placed under constant illumination for 3–4 weeks, and resistant colonies were restreaked on fresh solid medium containing at least 100 μg/mL Zeocin (see Note 6).

3.4 Confocal Microscopy

All P. tricornutum transformants are analyzed via confocal laser scanning microscope. 1. Leica TCS SP2 using an HCX PL APO 40×/1.25 to 0.75 oil CS objective after fixing with 4% paraformaldehyde – 0.0075% glutaraldehyde in 1× phosphate-buffered saline buffer. 2. The fluorescence of enhanced green fluorescent protein (eGFP) and plastid autofluorescence was excited with an argon laser at 488 nm and detected with two photomultiplier tubes at a bandwidth of 500–520 nm and 625–720 nm for eGFP and plastid autofluorescence, respectively (see Note 7). 3. The fluorescence of mRbuy3 was detected at a bandwidth of 580–605 nm. 4. Different dyes can be used to stain various subcellular organelles, including mitochondria, plasma membrane, and integrated vesicles (see Note 8). 5. Proteins localized in different membranes of P. tricornutum plastid usually have at least one transmembrane domain. To analyze the precise membrane topologies of these plastid membrane proteins, GFP-self assembling approach is recommended to be used (see Note 9).

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Notes 1. The RNA isolation containing potential genomic DNA should be treated by DNaseI to remove genomic DNA. 2. In addition to YFP, the fluorescence protein mRuby3 can also be used as tag for efficient in vivo protein localization studies with confocal laser scanning microscopy [7]. Its emission was detected efficiently between 580 and 605 nm, being distinguishable from the plastid autofluorescence (detectable between 625 and 720 nm) in vivo in P. tricornutum. 3. The plasmids can be constructed by using different methods, such as Gibson Assembly Master Mix Kit (New England Biolabs GmbH, Frankfurt, Germany) and EasyGeno Assembly Cloning Kit (Tiangen, China) [8, 9]. 4. Most plastid proteins encoded by the nucleus genome of diatom contain N-terminal bipartite targeting sequence, which is characterized by the presence of a conserved motif, namely ASAF or AFAP positioned at the cleavage site of the signal peptide. Before conducting the in vivo co-localization studies, the online tools of SignalP 3.0 (http://www.cbs.dtu.dk/ services/SignalP-3.0/) and TargetP 1.1 (http://www.cbs.dtu. dk/services/TargetP/) are recommended to search for the presence of putative N-terminal targeting sequences. 5. While we typically use biolistic bombardment, the plasmids for co-localization can also be transformed into P. tricornutum using electroporation with a Bio-Rad Gene Pulser Xcell Electroporation System as described by Zhang et al. [10]. For co-transformation, 2 μg of each linearized plasmid is recommended to be used. Colonies usually appear after 10~12 d, which is less than the method of biolistic bombardment. 6. PCR can be used to verify the presence of target protein-eGFP and marker protein-mRuby3 fusions in genomic DNA isolated from two antibiotic-resistant clones transformed with the constructed plasmid. Prior to confocal laser scanning microscopy, we often use general fluorescence microscopy to screen for the colonies that have strong mRuby3/GFP fluorescence signals. 7. During the fluorescence experiment, the bandwidths for plastid, eGFP, and mRuby3 should not be out of range; an argon laser at 488 nm should be in the suitable strength; otherwise, the false positive results might be observed. 8. For staining of the mitochondria, plasma membrane, or integrated vesicles, harvest P. tricornutum cells and wash once in 1×phosphate buffered saline (PBS, pH 7.5), then incubate them for 30 min in 0.5 mM MitoTracker Orange CMTMRos or for 25 min in 0.08 mM FM®4-64 in the dark, respectively.

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After staining, wash cells twice with 1×PBS and then fix with 4% paraformaldehyde-0.0075% glutaraldehyde in 1×PBS buffer for 20 min. After fixation, suspend cells in 30 μL PBS for microscopy. The cells from the staining of FM®4-64 dye will not be fixed and directly resuspended in 30 μL PBS for microscopy [11]. The nucleus can be stained using Hoechst 33342 (Hoechst AG, Frankfurt, Germany) and visualized using an excitation wavelength of 346 nm and an emission bandpass of 430–600 nm. ER-Tracker Blue-White DPX (Invitrogen) was used to visualize ER with an excitation wavelength of 375 nm and emission bandpass of 550–640 nm. Golgi-Tracker Red (Beyotime, Beijing, China) was used to locate Golgi apparatus with an excitation wavelength of 589 nm and emission spectrum of 610–650 nm. The plastid autofluorescence was visualized with an excitation wavelength of 488 nm and an emission wavelength of 650–750 nm [12]. 9. The proteins localized in the plastid membranes of diatom usually contain at least one transmembrane region. The TMHMM Ser ver (http://www.cbs.dtu.dk/ser vices/ TMHMM) is recommended to predict or search for the presence of potential transmembrane domains for target proteins. To further determine the orientation and topology of target protein, the self-assembling GFP system is recommended to be used as described previously [13–15]. In this system, a fusion protein consisting of the full-length sequence of target protein and the small self-assembling GFP fragment GFP_S11 will be simultaneously expressed with a large GFP_S1-10 fragment fused to marker protein with specific subcompartment such as cytosol, cER, PPC, IMS, or stroma.

Acknowledgments This work was supported by grants from the National Natural Science Foundation of China (41776175 and 31961133008 to YG, and 418068 to XL). References 1. Grosche C, Hempel F, Bolte K, Zauner S, Maier UG (2014) The periplastidal compartment: a naturally minimized eukaryotic cytoplasm. Curr Opin Microbiol 22:88–93 2. Cavalier-Smith T (2000) Membrane heredity and early chloroplast evolution. Trends Plant Sci 5:174–182 3. Kroth PG (2002) Protein transport into secondary plastids and the evolution of primary

and secondary plastids. Int Rev Cytol 221: 191–255 4. Kilian O, Kroth PG (2005) Identification and characterization of a new conserved motif within the presequence of proteins targeted into complex diatom plastids. Plant J 41:175– 183 5. Lamb JJ, Røkke G, Hohmann-Marriott MF (2018) Chlorophyll fluorescence emission

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spectroscopy of oxygenic organisms at 77 K. Photosynthetica 56:105–124 6. Lavaud J, Lepetit B (2013) An explanation for the inter-speciesvariability of the photoprotective non-photochemical chloro-phyll fluorescence quenching in diatoms. Biochim Biophys Acta 1827:294–302 7. Marter P, Schmidt S, Kiontke S, Moog D (2020) Optimized mRuby3 is a suitable fluorescent protein for in vivo co-localization studies with GFP in the diatom Phaeodactylum tricornutum. Protist 171(1):125715. https:// doi.org/10.1016/j.protis.2020.1257159 8. Hao X, Luo L, Jouhet J, Re´beille´ F, Mare´chal E, Hu H, Pan Y, Tan X, Chen Z, You L, Chen H, Wei F, Gong Y (2018) Enhanced triacylglycerol production in the diatom Phaeodactylum tricornutum by inactivation of a Hotdog-fold thioesterase gene using TALEN-based targeted mutagenesis. Biotechnol Biofuels 11:1–18. https://doi.org/10. 1186/s13068-018-1309-3 9. Huang W, Rı´o Ba´rtulos C, Kroth PG (2016) Diatom Vacuolar 1,6-β-Transglycosylases can functionally complement the respective yeast mutants. J Eukaryot Microbiol 63:536–546. https://doi.org/10.1111/jeu.12298 10. Zhang C, Hu H (2014) High-efficiency nuclear transformation of the diatom

Phaeodactylum tricornutum by electroporation. Mar Genomics 16:63–66 11. Liu X, Hempel F, Stork S, Bolte K, Moog D, Heimerl T, Maier UG, Zauner S (2016) Addressing various compartments of the diatom model organism Phaeodactylum tricornutum via sub-cellular marker proteins. Algal Res 20:249–257 12. You Y, Sun X, Ma M, He J, Li L, Porto FW, Lin S (2022) Trypsin is a coordinate regulator of N and P nutrients in marine phytoplankton. Nat Commun 13(1):4022. https://doi.org/10. 1038/s41467-022-31802-6 13. Cabantous S, Terwilliger TC, Waldo GS (2005) Protein tagging and detection with engineered self-assembling fragments of green fluorescent protein. Nat Biotechnol 23:102– 107 14. Moog D, Rensing SA, Archibald JM, Maier UG, Ullrich KK (2015) Localization and evolution of putative triose phosphate translocators in the diatom Phaeodactylum tricornutum. Genome Biol Evol 7(11):2955–2969 15. Lau JB, Stork S, Moog D, Sommer MS, Maier UG (2015) N-terminal lysines are essential for protein translocation via a modified ERAD system in complex plastids. Mol Microbiol 96(3): 609–620

Chapter 12 Monitoring of Lipid Fluxes Between Host and Plastid-Bearing Apicomplexan Parasites Sarah Charital, Amandine Lourdel, Nyamekye Quansah, Cyrille Y. Botte´, and Yoshiki Yamaryo-Botte´ Abstract Apicomplexan parasites are unicellular eukaryotes responsible for major human diseases such as malaria and toxoplasmosis, which cause massive social and economic burden. Toxoplasmosis, caused by Toxoplasma gondii, is a global chronic infectious disease affecting ~1/3 of the world population and is a major threat for any immunocompromised patient. To date, there is no efficient vaccine against these parasites and existing treatments are threatened by rapid emergence of parasite resistance. Throughout their life cycle, Apicomplexa require large amount of nutrients, especially lipids for propagation and survival. Understanding lipid acquisition is key to decipher host-parasite metabolic interactions. Parasite membrane biogenesis relies on a combination of (a) host lipid scavenging, (b) de novo lipid synthesis in the parasite, and (c) fluxes of lipids between host and parasite and within. We recently uncovered that parasite need to store the host-scavenged lipids to avoid their toxic accumulation and to mobilize them for division. How can parasites orchestrate the many lipids fluxes essential for survival? Here, we developed metabolomics approaches coupled to stable isotope labelling to track, monitor, and quantify fatty acid and lipids fluxes between the parasite, its human host cell, and its extracellular environment to unravel the complex lipid fluxes in any physiological environment the parasite could meet. Key words Apicomplexa, Apicoplast, Lipid fluxes, Metabolism, Stable isotope, GC-MS

1

Introduction Apicomplexan is a phylum of protist comprising intracellular parasites responsible for major infectious and chronic diseases, including toxoplasmosis. To ensure their rapid propagation and pathogenicity within the host, toxoplasma parasites require large amounts of lipids, needed for their membrane biogenesis. Precisely, these parasites’ membranes are mainly composed of phospholipids (PLs) like phosphatidylcholine (PC), phosphatidylethanolamine (PE), or phosphatidylinositol (PI) [1]. These FAs are obtained by the parasite through massive scavenging of host resources and de novo fatty acids (FAs) synthesis by the parasite, thanks to the type II

Eric Mare´chal (ed.), Plastids: Methods and Protocols, Methods in Molecular Biology, vol. 2776, https://doi.org/10.1007/978-1-0716-3726-5_12, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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FA synthesis (FASII) pathway found in its apicoplast [2]. Toxoplasma is able to adapt its metabolism to face different nutrient environments, by regulating the metabolic activities of FAs acquisition pathways: increasing scavenging in low nutrient conditions or increasing the storage in rich nutrient conditions [3]. These constant lipid fluxes between the parasite and the host can be monitored and analyzed using stable isotope labelling. To identify the origin of FAs used by the parasite for bulk membrane biogenesis (i. e., scavenged from host media, scavenged from host lipids, or made de novo by the parasite), our team developed novel lipid flux approaches combining stable isotope substrates-containing media (13C-glucose, 31d-C16:0) and mass spectrometry-based lipidomic analyses (i.e., fluxomic approaches) [4]: (1) monitoring the apicoplast FASII activity by tracking FASII-made FAs, (2) monitoring FAs directly scavenged from the host by tracking host-derived FAs, and finally (3) monitoring the extracellular host environment by tracking the scavenging of extracellular FAs uptake. Upon parasite grown in desired nutritional/stable isotope labelling conditions, followed by parasite collection, parasite metabolic quenching, and metabolite extraction on ice, labelled lipids are extracted. Extracted lipids are then either (i) derivatized for analysis by gas chromatography coupled with mass spectrometry (GC-MS) or (ii) directly prepared for LC/MSMS analysis.

2

Materials

2.1 Parasite Cell Lines and Cell Culture

1. Toxoplasma gondii RH-TIR tachyzoite lines are used in four biological replicates, with a similar cell number (which will be further used for normalization purposes): ~5 × 107 cells. For U-13C-glucose labelled samples, best results are achieved when using a concentration of 0.1 to 0.5 × 106 cells to be injected to the GC-MS. 2. The parasite host cell human foreskin fibroblasts (HFF) are cultured using Dulbecco’s Modified Eagle’s Medium (DMEM, Gibco) or glucose-free DMEM, supplemented with 10% fetal bovine serum (FBS, Gibco), 2 mM glutamine (Gibco), and 25 μg/mL gentamicin (Gibco) at 37 °C and 5% CO2. 3. Parasites are grown and routinely passaged in new HFF flasks on a regular basis (~twice/week) with the same inoculum size (~100 μL from the previous culture of freshly egressed parasites) for each biological replicate. Parasites then grow in host HFF complemented with a culture medium composed as such: DMEM supplemented with 1% fetal bovine serum (FBS, Gibco), 2 mM glutamine (Gibco), and 25 μg/mL gentamicin (Gibco) at 37 °C and 5% CO2.

Monitoring of Lipid Fluxes Between Host and Plastid-Bearing Apicomplexan. . .

2.2 Solvents, Reagents, and Material

199

1. Gas chromatograph coupled to a mass spectrometer (GC-MS). 2. Glass vials (1.5 mL) with sealed caps, complying with the use of solvents (such as N2, N8, and N9 from NPCI Analytics, or equivalent). 3. Pasteur pipettes. 4. 3-μm pore-size filter membrane with a syringe-adaptor. 5. 1.5 mL tubes with caps (e.g., from Eppendorf). 6. Sonicator bath, vortex. 7. Solvents: Chloroform, methanol, and a mix of chloroform and methanol, 1:1 (v/v). 8. Standards for metabolic extraction: Tridecanoic acid (C13:0) 1 mM in methanol (Sigma) and heneicosanoic acid (C21:0) 1 mM in methanol (Sigma). 9. Radiolabeled molecules: 31d-C16:0 U-13C-glucose, U-12C-glucose.

(palmitic

acid),

10. Antioxidant agent BHT: 1% butyl hydroxy toluene in methanol (Sigma). 11. Solutions: 0.1 M hydrochloric acid (HCl) in water, 0.2% potassium chloride (KCl) in water and DNase free H2O (Gibco). 12. 1x PBS: 137 mM NaCl, 2.7 mM KCl, 10 mM Na2HPO4, and 1.8 mM KH2PO4. 13. Derivatization reagents: Trimethylsulfonium hydroxide (TMSH, Macherey-Nagel) in chloroform/methanol, 7:25 (v/v).

3

Methods

3.1 Lipidomic Analysis: Total Lipid Analysis

The protocol needs to be performed on ice from the beginning unless mentioned otherwise. Decide which FA or lipid standard you want to use according to the analysis you want to perform; usually internal standards tridecanoic acid (C13:0) and heneicosanoic acid (C21:0) are used. Do not forget to vortex and warm samples before use. In this protocol, special labelling names are proposed for the tubes. 1. Aliquot solvents in N2 glass vials and prepare N8 vials for blank and one for internal standards. 2. Pre-heat BHT and standards for 10 min at 37 °C. 3. Take a parasite pellet in an Eppendorf tube stored on ice. 4. Add 10 μL of tridecanoic acid (C13:0) in 1 mM methanol, 10 μL of heneicosanoic acid (C21:0) in 1 mM methanol, and 1 μL of BHT in ethanol in each Eppendorf tube (see Note 1 and 2). Add 50 μL H2O.

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5. Add 250 μL methanol and sonicate in a sonicator bath to suspend the pellet (see Note 3 and 4). 6. Add 125 μL chloroform and sonicate. 7. Incubate for 15 min on ice and centrifuge at 2500 rcf for 5 min at 4 °C. 8. Transfer the supernatant into a N8 glass vial (A) to pool the organic phase (see Note 5). 9. To the original Eppendorf tube with sample pellet, add 10 μL of HCl 0.1 M and 125 μL of methanol and sonicate to suspend the pellet. 10. Add 250 μL chloroform and sonicate. 11. Incubate for 5–10 min on ice and centrifuge at 1000 rcf for 5–10 min at RT. 12. During the spin-down, add 375 μL of chloroform and 190 μL of KCl 0.2% in the N8 (A) glass vial. 13. Transfer the supernatant of the centrifuged Eppendorf tube to the N8 (A) glass vial (see Note 6). Vortex and centrifuge the N8 glass vial at 3000 rcf for 30 s at RT (see Note 7). 14. Transfer the bottom layer to a new N8 glass vial (B) (see Note 8). 15. Add 100 μL H2O to N8 (B) vials, vortex and centrifuge at 1200 rcf for 1 min at RT. Remove the top layer (see Note 9). 16. The final organic phase should be around 600–800 μL. 17. Insert 0.2 or 0.25 mL vial inserts in N9 glass vials and transfer an aliquot of the N8 (B) glass vial depending on the cell concentration obtained during parasite extraction (i.e., an aliquot of 250 μL will be transferred for a number of 1 × 107 cells; an aliquot of 100 μL will be transferred for a number of 5 × 107 cells). 18. Dry the N9 and the N8 (B) glass vials with the remaining sample in a speed vacuum at RT or 30 °C (45 °C maximum), (see Note 10). 19. Once completely dried, add 15–25 μL TMSH solution (7:25 in chloroform/methanol, v/v) according to sample amount, cell number, and concentration. Vortex each samples. 20. Prepare wash solvents on the GC-MS (2x hexane or dichloromethane and 2x methanol, depending on the sample number, can add more vials of wash solvents) and blank (hexane). Place samples on the sample trays, adjust the running sequence, and run the GC-MS.

Monitoring of Lipid Fluxes Between Host and Plastid-Bearing Apicomplexan. . .

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3.2 Stable Isotope Metabolic Labelling

Fatty acids required for lipid synthesis in T. gondii parasites are obtained from both the host, the host environment (i.e., medium), and/or can be synthesized de novo by the parasite, most often via the apicoplast FASII pathway for fatty acid/lipid precursor de novo synthesis. To identify the origin of FAs used by the parasite for bulk membrane biogenesis (i.e., scavenged from host media, scavenged from host lipids or made de novo by the parasite), novel lipid flux approaches combining stable isotope substrates-containing media (13C-glucose and 31d-C16:0) and mass spectrometry-based lipidomic analyses (i.e., fluxomic approaches) have been set up.

3.2.1 Monitoring the Apicoplast FASII Activity

To monitor de novo FA synthesis by the parasite apicoplast FASII, parasites are labelled with U-13C-glucose, which is added to a glucose-free medium on confluent host cells together with parasites as previously reported. 1. Grow HFF cells to confluence in a normal glucose containing DMEM (10% FBS). 2. When confluent monolayer of HFF is obtained, infect freshly egressed T. gondii tachyzoites with a glucose-free DMEM (1% FBS) supplemented with U-13C-glucose or U-12C-glucose, at a final concentration of 800 μM (you can use a 100x stock solution). 3. Harvest parasites after 48–72 h incubation at 37 °C and 5% CO2. 4. Metabolically quench parasites in a dry ice and ethanol slurry in a Falcon tube until the sample reaches 4 °C. From then on, all steps are carried on ice at 4 °C (see Note 11). 5. Syringe passage twice and then filter parasites with a 3-μm pore-size filter membrane. 6. Count parasite number in the Falcon tube using a Thoma or Mallassez hemocytometer (see Note 12). 7. Centrifuge parasites at 4 °C at 600 rcf for 10 min. 8. Wash parasite pellet with ice-cold phosphate-buffered saline (1X PBS) thrice. 9. Transfer the final parasite pellet to a microcentrifuge tube (see Note 13). 10. Extract total lipid from the parasite pellet, derivatize using TMSH (Macherey-Nagel), and analyze by GC-MS as described in Subheading 3.1: Lipidomic analysis. 11. All fatty acid methyl esters (FAMEs) are identified by comparison of retention time and mass spectra from GC-MS with authentic chemical standards. To quantify the concentration of FAMEs, normalize to different internal standards (C13: 0 and C21:0) and to parasite number.

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12. Calculate the natural abundance of 12C-glucose and the amount of 13C-carbon source in the culture medium. 13. After correction for natural abundance, calculate 13C incorporation to each FA as the percentage of the metabolite pool containing one or more 13C atoms. 14. Calculate the mass isotopomer distribution (MID) of each FAME from the shift in isotopic mass dependent on the amount of 12C carbons compared to the integration of 13C carbon atoms. 15. Use the MID of each FAMEs to determine the degree of the incorporation of 13C into FA (% carbon incorporation). 16. Finally, calculate the concentration of all isotopomers of 13 C-labeled FAMEs and normalize to authentic internal standards and parasite number to obtain the total abundance of 13 C-labeled Fas. 3.2.2 Tracking HostDerived Fatty Acids

To monitor FAs directly scavenged from the host, host cells are pre-labelled using U-13C-glucose in the absence of parasites and then grown as previously described to allow the labelling of all host glycerolipids at the level of their acyl chains. These 13C-pre-labeled host cells are then infected with parasites in normal culture medium, containing regular 12C-glucose. 1. Grow HFF cells to confluence in a glucose-free DMEM (10% FBS) complemented with stable isotope U-13C-glucose at a final concentration of 800 μM. 2. Once reached confluence, infect these pre-labelled HFF with freshly egressed parasites in the presence of normal glucose containing DMEM (1% FBS). 3. The host HFF and parasites are metabolically quenched separately, and harvest as described above (steps 3–9, Subheading 3.2.1). 4. Extract total lipid, derivatize them using TMSH (MacheryNagel), and quantify their lipid content by GC-MS analyses as described in Subheading 3.1: Lipidomic analysis. 5. As previously described, determine the degree of the incorporation of 13C into FA (% carbon incorporation) by the mass isotopomer distribution (MID) of each FAMES (steps 11–15, Subheading 3.2.1). 6. First, analyze the total abundance of 13C-labeled FA for HFF to check labelling of the metabolites (described step 16, Subheading 3.2.1). 7. Second, calculate the same for parasites to confirm direct uptake of 13C-labeled FA from the host.

Monitoring of Lipid Fluxes Between Host and Plastid-Bearing Apicomplexan. . . 3.2.3 Tracking the Scavenging of Extracellular Fatty Acid Uptake Using Deuterated Fatty Acid in the Extracellular Medium (Monitoring Extracellular Host Environment)

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To monitor the FAs sourced from the external environment, parasites and confluent host cells are grown in a medium complemented with deuterated palmitic acid (31d-C16:0). 1. Dissolve deuterated palmitic acid (31d-C16:0) in 10 mM in FA-free bovine serum albumin/PBS solution by sonication in a water bath for 30 min. 2. Incubate this dissolved 31d-C16:0 at 55 °C for 30 min. 3. Grow HFF cells to confluence in a normal glucose containing DMEM (10% FBS). 4. Infect confluent monolayer of HFF with freshly egressed parasites in the presence of normal glucose containing DMEM (1% FBS) and let invade for 2 h. 5. Following the invasion, wash off the uninvaded parasites with DMEM. 6. Add normal culture medium-DMEM containing 31d-C16:0 at a final concentration of 0.1 mM. Let grow for 24–48 h. 7. Harvest parasite by metabolic quenching as described previously (steps 3–9, Subheading 3.2.1). 8. Extract total lipid, derivate using TMSH, and further analyze by GC-MS as described in Subheading 3.1: Lipidomic analysis.

4

Notes 1. To avoid pipet contamination, drop off every standards on a different side of the tube each time. 2. For standards dissolved in methanol, use a positive displacement pipetman. BHT can be used with a normal pipetman. 3. Never reverse the sample tube to mix, use a sonicator bath. 4. Wear safety glasses when using methanol to protect the experimenter for droplets and splashes that could spill in your eyes. 5. To transfer in N8 vials, use Pasteur pipettes for each sample and keep them to re-use for next steps. 6. Pellet could be too soft or does not precipitate; in this case, transfer all including debris or leave. 7. N8 vials spin down can least until 5 min. 8. To transfer the bottom layer of the N8 (A) vial, use the same Pasteur pipette as above in steps 8 and 13. Create bubbles to pass the upper layer and thus avoiding taking this phase. 9. This step 15 could be omitted 10. Once dried, samples in N8 (B) or N9 with inserts can be stocked at -20 °C for later use (up to 6 months, 12 months maximum).

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11. Once quenched, the metabolism of the parasite is stopped; therefore, sample should be kept on ice and all steps should be conducted on ice or at 4 °C. 12. Label each tube during parasite harvest with the number of parasite that you count. 13. Parasites can be stored up to 12 months as a frozen parasite pellet at -80 °C until use.

Acknowledgments This work and their authors were supported by Agence Nationale de la Recherche, France (Project ApicoLipidAdapt grant ANR-21CE44-0010; Project Apicolipidtraffic grant ANR-23-CE15-000901), The Fondation pour la Recherche Me´dicale (FRM EQU202103012700), Laboratoire d’Excellence Parafrap, France (grant ANR-11-LABX-0024), LIA-IRP CNRS Program (Apicolipid project), the Universite´ Grenoble Alpes (IDEX ISP-IRGA Apicolipid), Re´gion Auvergne Rhone-Alpes for the lipidomics analyses platform (Grant IRICE Project GEMELI), the CEFIPRA via a Collaborative Research Program Grant (Project 6003-1). References 1. Welti R, Mui E, Sparks A, Wernimont S et al (2007) Lipidomic analysis of Toxoplasma gondii reveals unusual polar lipids. Biochemistry 46(48):13882–13890. https://doi.org/10. 1021/bi7011993 2. Mazumdar J, Striepen B (2007) Make it or take it: fatty acid metabolism of apicomplexan parasites. Eukaryot Cell 6(10):1727–1735. https://doi.org/10.1128/EC.00255-07 3. Amiar S, Katris NJ, Berry L et al (2020) Division and adaptation to host environment of

apicomplexan parasites depend on apicoplast lipid metabolic plasticity and host organelle remodeling. Cell Rep 30(11):3778–3792.e9. https://doi.org/10.1016/j.celrep.2020. 02.072 4. Dass S, Shunmugam S, Berry L et al (2021) Toxoplasma LIPIN is essential in channeling host lipid fluxes through membrane biogenesis and lipid storage. Nat Commun 12(1):2813. h ttps://d oi.org/10.1038/s414 67-02122956-w

Chapter 13 Extraction and Quantification of Lipids from Plant or Algae Vale´rie Gros, Josselin Lupette, and Juliette Jouhet Abstract In plants and algae, the glycerolipidome changes in response to environmental modifications. For instance, in phosphate starvation, phospholipids are degraded and replaced by non-phosphorus lipids, and in nitrogen starvation, storage lipids accumulate. In addition to the well-known applications of oil crops for food, algae lipids are becoming a model for potential applications in health, biofuel, and green chemistry and are used as a platform for genetic engineering. It is therefore important to measure accurately and quickly the glycerolipid content in plants and algae. Here we describe the methods to extract the lipid and quantify the fatty acid amount of the lipid extract and the different lipid classes that are present in these samples. Key words Lipid, Extraction, HPLC, Quantification, Gas chromatography, Mass spectrometry, Fatty acid methyl ester

1

Introduction The assessment of the lipid content and quality in plant and algae samples is critical in many studies addressing fundamental or applied questions. The literature on plant lipid biology is extremely abundant, for questions ranging from the response to environmental stresses to the improvement of the yield in oil crops. Recently, phytoplankton have become novel attractive models, firstly because they represent still poorly understood branches of the eukaryotic biodiversity; secondly, to address the question of the effect of climate change on oceanic ecosystems; and thirdly, since they are promising biotechnological models for the development of high value molecules. Characterization of glycerolipid profiles of several species of microalgae has been carried out in recent years in order to establish high value model species for biotechnology industries [1– 4]. The accurate and rapid determination of the glycerolipidome profile of plants or algae is therefore an important challenge. In

Eric Mare´chal (ed.), Plastids: Methods and Protocols, Methods in Molecular Biology, vol. 2776, https://doi.org/10.1007/978-1-0716-3726-5_13, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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photosynthetic organisms, fatty acids are synthesized within the stroma of the chloroplast, and photosynthetic lipids are specifically produced within the organelle itself. Whole cell lipid profiling gives therefore insights on chloroplast lipids as well [5]. The first quantification of total fatty acids dates back to the late of 1950s by Folch [6] and Bligh and Dyer [7] methods. Separation of glycerolipids is conventionally carried out by thin layer chromatography (TLC) and quantified by gas chromatography coupled to flame ionization detector (GC-FID) after methanolysis of fatty acids [1]. The progress made on lipid analyzes has since been considerable with the development of electrospray tandem mass spectrometry (ESI-MS/MS) in the late of 1980s allowing highthroughput lipid analyzes. However, differences in the quantification of lipids were observed between the GC-FID and the LC-MS/ MS methods. Our research team has developed recently a method based on an external standard per studied organism under the form of a known qualified and quantified lipid extract that corrects the bias between these two techniques [8]. In this chapter, we describe the key steps for the extraction, separation, and methanolysis of plant or microalgae lipids and their quantification by LC-MS/MS methods. This method relies on the knowledge of the glycerolipidome and on the production of a quantified lipid extract of the studied plant or algae. The glycerolipidome and the lipid extract can be established by the same method as the one described in chapter “Establishment and quantification of chloroplast lipidome” of this book.

2

Materials All reagents are prepared and stored at room temperature. Always use glass vessels and never plastic with organic solvents. Wash all vessels without detergent but with distilled water, then ethanol. All solvents must be for Analysis grade except Hexane that must be for GC grade. Chloroform should be ethanol stabilized. Diligently follow all waste disposal regulations when disposing waste materials.

2.1

Lipid Extractions

1. Corex 30 mL tubes. 2. Freeze dryer. 3. Liquid nitrogen. 4. Quartz wool (see Note 1). 5. Potters pestles (see Note 2). 6. Argon: Argon is used to dry samples by applying a flow of gas using thin glass Pasteur pipettes or metal needles (see Note 3).

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7. Ultrapure water (prepared by purifying deionized water, to attain a sensitivity of 18 MΩ-cm at 25 °C). 8. NaCl 1% (w/v) in ultrapure water. 9. Hemolysis glass tube for single use. 2.2

Methanolysis

1. Pyrex 20 mL glass tubes (160 mm × 16 mm tubes in borosilicate glass, with high thermal resistance) with screw top. Screw caps in phenolic resin with rubber seal lined with inert polytetrafluoroethylene (PTFE). 2. Standard C15 (fatty acid with 15 carbons) solution: Powder of pentadecanoic acid (99%), stored at RT. Prepare a solution at 0.5 mg/mL in chloroform/methanol (1/2, v/v) (see Note 4). 3. Dry-bath hot block. 4. Methanolysis buffer: Sulfuric acid (H2SO4 24N) 2.5% in methanol (v/v) (see Note 5). 5. Ultrapure water (prepared by purifying deionized water, to attain a sensitivity of 18 MΩ-cm at 25 °C). 6. Hexane for GC analysis 7. Hemolysis glass tube for single use.

2.3 Total Fatty Acid Quantification by Gas Chromatography – Flame Ionization Detector (GC-FID)

1. Vials for automatic sampler with an insert of 250 μL and screw caps 9 mm with polytetrafluoroethylene seal. 2. Hexane for gas chromatography (GC) analysis. 3. Column BPX70 (70% Cyanopropyl Polysilphenylene-siloxane) for GC: Length 30 m, internal diameter 0.22 mm, film thickness 0.25 μm. 4. Gas chromatography – Flame ionization detector. 5. Commercial fatty acid methyl ester (FAME) reference or standard solution to calibrate the GC-FID retention time.

2.4 Lipid Class Quantification by Liquid Chromatography– Mass Spectrometry (LC-MS)

1. Internal standard solution: Prepare 1 mL of stock solution for each internal standard at 1.25 mM in chloroform/methanol [2/1, (v/v)] (see Note 6). Internal standards were obtained from Avanti Polar Lipids Inc. for PE 18:0–18:0 and DAG 18: 0–22:6 and used directly. For SQDG, natural extract is purchased from Avanti Polar Lipids Inc. and hydrogenated to obtain SQDG 16:0–18:0 (see Note 7). To prepare the internal standard solution, in a volumetric flask of 50 mL, add 50 μL of each individual standard and complete to 50 mL with chloroform/methanol [2/1, (v/v)]. Aliquot the solution by 1 mL in hermetically sealed vials and store at -20 °C. 2. Hemolysis glass tube for single use.

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3. Argon (see Note 4). 4. High pressure liquid chromatography (HPLC) coupled to a triple quadripole mass spectrometer (see Note 8).

3

Methods

3.1 Lipids Extractions from Whole Plant (Rosettes) or from Plant and Algae Cell Suspension

1. For plant material: Harvest the rosettes or the cell in 50 mL Falcon pre-weighted and weigh the Falcon with the plant material to obtain the sample fresh weight. To obtain enough lipid extract, collect between 100 and 300 mg of plant material. 2. For algae material: Harvest the cells in 50 mL Falcon by centrifugating the culture until all the cells are pelleted and remove the culture media. To obtain enough lipid extract, collect around 100 million cells. 3. Drop the falcon in liquid nitrogen. Store the tube at -80 °C. 4. Make three holes with a needle in Falcon caps. Put the Falcon tubes in freeze-dry bottle and plug the bottle on the freeze dryer. Open the valve very slowly to avoid air movement. Wait until the pressure is stabilized before plugging another bottle. Leave overnight to freeze-dry (see Fig. 1a). 5. Remove the bottle by closing the valve very slowly to avoid air movement and sample loss. Weigh the Falcon tubes to obtain the sample dry weight and put back the Falcon tube in liquid nitrogen. 6. Pre-heat a water bath with a hot block inside. Experiment can start when the water is above 80 °C. In the same time, warm up an Erlenmeyer flask closed with double aluminum foil containing 200 mL of absolute ethanol until it boils (see Figs. 1b and 2). 7. Lipids extractions: Transfer plant material in 30 mL Corex tube in liquid nitrogen (see Note 9). With a Potter pestle, grind the frozen plants in powder inside the Corex tube. 8. Leave the tube outside the liquid nitrogen to warm it up a little bit with the pestle inside. 9. Add 4 mL of boiling ethanol, close the tube with double aluminum foil, and put it in the hot block in the water bath for 5 min with some shaking with the pestle (see Note 10). This step will inhibit phospholipase D activity for plant and lipase A1 activity for algae. 10. Remove the tube from the hot block, check the ethanol volume left in the tube and if necessary complete up to 4 mL with boiling ethanol. Add 2 mL of methanol to rinse the pestle directly in the tube. Remove the pestle and add 8 mL of chloroform. Apply a flow of argon in the solvent mixture to

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Fig. 1 Setup for lipid extraction. (a) Example of a freeze dryer. Tissue to freeze-dry is stored in a Falcon tube. The lid must be pierced at least 3 times to allow the water to get out of the tube. The Falcon tube is then put overnight in a specific container of the freeze-dryer and plugged into the machine. Freeze-drying process must be done at least overnight. (b) Example of setup for steps 5–9: prepared bottles of methanol and chloroform on the side, boiling ethanol in the middle and pre-warm water bath at 80 °C with a holding block that fits Corex tube. (c) Example of setup for steps 10 and 11: prepared bottles of chloroform/methanol 2/1 (v/v) and NaCl 1% (w/v) on the side, one tube from step 9 after 1 h waiting and one clean tube with the funnel containing the ethanol washed quartz wool

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Fig. 2 Procedure of the lipid extraction from steps 6 to 13. Step 6: Grind the frozen tissue with a pestle. Step 8: Boil for 5 min with the ethanol. Step 9: Complete the ethanol volume, add the chloroform and methanol and bubble argon for mixing the solvents and remove oxygen. Step 10: Filtrate the extract over the funnel in a new clean tube, rinse the corex tube with chloroform/methanol 2/1 (v/v) and pour the liquid over the funnel, and remove the funnel and add NaCl 1% (w/v). Step 11: Mix the solvent by bubbling argon at least 30 s and centrifugate. Step 12: Collect the bottom phase into a clean glass tube and evaporate under argon. Step 13: Resuspend the lipid extract in chloroform, transfer it into a clean hemolysis glass tube, and evaporate under argon

remove oxygen during 1 min (see Note 11 and Fig. 2). Close the tube with foil and leave it for 1 h at least at room temperature. 11. Filter the liquid in a new Corex tube with a funnel plug with ethanol washed glass wool to remove cell debris. Add 3 mL of chloroform:methanol (2:1, v:v) in the Corex tube containing the cell debris to rinse it and pour the liquid in the funnel (see Fig. 1c).

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12. Remove the funnel and add 5 mL of NaCl 1% to the filtrate mixture. Apply a flow of argon inside the tube to mix up solvents during 1 min and centrifuge 10 min at 1000× g to separate the organic and aqueous phase. 13. Collect the lower phase (organic phase) with a Pasteur pipette (see Note 12) and transfer it in a clean Corex glass tube. Dry all the solvents by applying a flow of argon at the top of the liquid. 14. Transfer the lipid extract in a hemolysis glass tube by rinsing the Corex glass tube with 3 × 1 mL of chloroform. Dry all the chloroform by applying a flow of argon at the top of the liquid, close the tube, and store the lipid extract at -20 °C. Lipid extract is stable for several months. 3.2 Production of Fatty Acid Methyl Esters (FAMEs)

1. Turn on the hot block at 100 °C and pre-warm an aliquot of standard C15 solution (see Note 13). 2. Resuspend the lipid extract in 1 mL of chloroform. Uptake between 10 and 100 μL of the lipid extract (see Note 14) and transfer it into the methanolysis tube. 3. With a Hamilton syringe, add 10 μL of C15 solution (5 μg/ tube) in the methanolysis tube. 4. Add 3 mL of methanolysis buffer in all the methanolysis tubes. In each tube, there are: C15 solution, some lipid extract, and the methanolysis buffer. Close tightly the glass tube. Incubate 1 h at 100 °C for the methanolysis reaction (esterification reaction) to occur. 5. Take the tube out of the hot block 5 min at room temperature to cool down the tube. Stop the reaction by adding 3 mL of water. 6. Add 3 mL of hexane to extract the FAMEs and vortex vigorously. Wait at least 20 min at room temperature to allow the biphase to separate correctly. At this step, biphase can be stored a few days at 4 °C if necessary. Take the upper phase (hexane phase) containing the FAMEs, transfer it in a hemolysis glass tube, and dry it under argon. 7. Repeat step 6 to re-extract FAMEs from the methanol-water phase by adding again 3 mL of hexane in the methanolysis tube. Pour the upper phase in the same hemolysis glass tube than the one used in step 6 and dry it under argon. 8. To concentrate the FAMEs at the bottom of the glass tube, rinse the tube wall with 200 μL of hexane. Allow the liquid to rest at the bottom of the tube and dry it gently under argon. 9. Store the FAMEs at -20 °C for few months or proceed with GC analysis.

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3.3 Quantification of FAMEs by Gas Chromatography– Flame Ionization Detector (GC-FID)

1. Resuspend the FAMEs in 45 μL of hexane and transfer the FAMEs in an insert vial. Seal the vial tightly to avoid evaporation. 2. Inject 2 μL of the sample in the GC-FID on a BPX70 column. Nitrogen is used as carrier gas with 3.5 mL/min constant flow compensation, split ratio is 13.3:1, injection temperature of 200 °C, detector temperature of 280 °C, and the oven temperature range start at 130 °C, hold for 7.5 min at 130 °C, ramp up to 180 °C at 3 °C/min, and hold 10 min at 180 °C. This will allow FAMEs separation from 12 C up to 24 C in function of the chain length and the number of desaturation. 3. Each FAME is identified by comparison of its retention times with those of standards. FID response is dependent on the mass of the FAME; therefore, each FAME will be quantified by the surface peak method using C15 surface peak for calibration with the following equation: Quantity in μg of FAME =

Area of FAME peak × Quantity in μg of C15 ð5 μgÞ : Area of C15 peak

4. By adding each FAME quantity and taking into account the volume used for the methanolysis, GC-FID analysis gives the total fatty acid content of the lipid extract as well as its fatty acid composition. 3.4 Quantification of Lipid Molecules by LC/ MS/MS

1. Prepare quantified control sample that will be used as an external standard (ES) for mass spectrometry quantification: Start from 4 g of fresh weight material of the organism to be analyzed. Extract the lipid as described in Subheading 3.1 but with volume multiply by 10. When the lipid extract is dry, resuspend it in 10 mL chloroform and take 3 × 2 μL to do the methanolysis and the GC-FID quantification in triplicate. Quantify each lipid class by TLC and GC-FID as described in chapter “Establishment and quantification of chloroplast lipidome” in this book. Aliquot the ES by 25 nmol. This ES will provide correction factor to quantify the lipid present in the sample to analyze ([8] and see Note 15). 2. Uptake a volume of the sample lipid extract corresponding to 25 nmol of total FA. Dissolve the lipid extract in 100 μL of chloroform/methanol [2/1, (v/v)] containing 125 pmol of PE 18:0–18:0, DAG 18:0–22:6, and SQDG 16:0–18:0. Do the same process with the ES. 3. Lipids are then separated by HPLC and quantified by ESI-MS/ MS. Lipid classes are separated using an HPLC system on a

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150 mm × 3 mm (length × internal diameter) 5 μm diol column at 40 °C. The mobile phases consisted of hexane/isopropanol/ water/ammonium acetate 1 M, pH 5.3 [625/350/24/1, (v/v/v/v)] (A) and isopropanol/water/ammonium acetate 1 M, pH 5.3 [850/149/1, (v/v/v)] (B). The injection volume is 20 μL, corresponding to 5 nmol of total fatty acid (see Note 16), and each sample is injected 3 times as technical replicates. After 5 min, the percentage of B was increased linearly from 0% to 100% in 30 min and stayed at 100% for 15 min. This elution sequence was followed by a return to 100% A in 5 min and an equilibration for 20 min with 100% A before the next injection, leading to a total runtime of 70 min. The flow rate of the mobile phase is 200 μL/min (see Note 17). 4. Mass spectrometric analysis is done on a triple quadripole mass spectrometer (see Note 18). The quadrupoles Q1 and Q3 were operated at widest and unit resolution, respectively. Specific MRM (Multiple Reaction Monitoring) scans used to define molecular species need to be known for each organism (see Note 19). In Table 1 is described as an example the acquisition settings for Arabidopsis glycerolipids. 5. To quantify each molecule of lipid, each chromatographic peak must be integrate by the mass spectrometer software. The area of the peak is proportional to the quantity of the molecule. Because mass spectrometry detection efficiency is always molecule dependent, the internal standard and the external standard are used to correct molecule bias quantification. Each molecular species of a lipid class (ms_LIP) is corrected by its corresponding internal standard (seg_is) to neglect the matrix effect (see Note 20) by applying the following formula with Q corresponding to the quantity in pmol: Qðms LIPÞ =

Area ðms LIPÞ  Qðseg isÞ: Area ðseg isÞ

The same process is done for the molecule species of the ES (ms_ES). Then all molecules of the same lipid class are summed to obtain the quantity of one lipid class: Q ðESÞ =

Q ðms ESÞ

The quantity of the lipid class of the ES measured by mass spectrometry is compared to the one measured by GC-FID to obtain a correction factor (CF) per lipid class: CF =

QGCFID ðESÞ Q ðESÞ

Compound name

DAG-18-1_18-1

DAG-18-2_18-1

DAG-18-2_18-1

DAG-18-3_18-1

DAG-18-3_18-1

DAG-18-3_18-2

DAG-18-3_18-2

DAG-18-3_18-3

DAG-16-0_18-1

DAG-16-0_18-1

DAG-16-0_18-2

DAG-16-0_18-2

DAG-16-0_18-3

DAG-16-0_18-3

DAG-16-1_18-3

DAG-16-1_18-3

DAG-16-0_16-1

DAG-16-0_16-1

DGDG-36-1

DAG A.t.

DAG A.t.

DAG A.t.

DAG A.t.

DAG A.t.

DAG A.t.

DAG A.t.

DAG A.t.

DAG A.t.

DAG A.t.

DAG A.t.

DAG A.t.

DAG A.t.

DAG A.t.

DAG A.t.

DAG A.t.

DAG A.t.

DAG A.t.

DGDG A.t.

Segment 1: from 0 to 7 min

Compound group

964

584

584

606

606

608

608

610

610

612

612

630

632

632

634

634

636

636

638

Prec ion

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

MS1 Res

623

311

313

311

335

313

335

313

337

313

339

335

335

337

335

339

337

339

339

Prod ion

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

MS2 Res

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

Dwell

Table 1 Transition and mass spectrometer parameters used to detect and quantify Arabidopsis lipid

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

Fragmentor

8

16

16

16

16

16

16

16

16

16

16

16

16

16

16

16

16

16

16

Collision energy

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

Cell accelerator voltage

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Polarity

214 Vale´rie Gros et al.

DGDG-36-2

DGDG-36-3

DGDG-36-4

DGDG-36-5

DGDG-36-6

DGDG-34-0

DGDG-34-1

DGDG-34-2

DGDG-34-3

DGDG-34-4

DGDG-34-5

DGDG-34-6

DGDG-32-1

DGDG-32-2

DGDG-32-3

MGDG-36-1

MGDG-36-2

MGDG-36-4

MGDG-36-5

MGDG-36-6

MGDG-34-0

MGDG-34-3

MGDG-34-4

MGDG-34-5

DGDG A.t.

DGDG A.t.

DGDG A.t.

DGDG A.t.

DGDG A.t.

DGDG A.t.

DGDG A.t.

DGDG A.t.

DGDG A.t.

DGDG A.t.

DGDG A.t.

DGDG A.t.

DGDG A.t.

DGDG A.t.

DGDG A.t.

MGDG A.t.

MGDG A.t.

MGDG A.t.

MGDG A.t.

MGDG A.t.

MGDG A.t.

MGDG A.t.

MGDG A.t.

MGDG A.t.

766

768

770

776

792

794

796

800

802

904

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

908 906

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

926

928

930

932

934

936

938

954

956

958

960

962

587

589

591

597

613

615

617

621

623

563

565

567

585

587

589

591

593

595

597

613

615

617

619

621

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

8

8

8

8

8

8

8

8

8

8

8

8

8

8

8

8

8

8

8

8

8

8

8

8

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

(continued)

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Extraction of Lipids 215

Compound name

MGDG-34-6

MGDG-32-0

MGDG-32-1

MGDG-32-2

MGDG-32-3

DAG-18-0_22-6

DAG-18-0_22-6

TAG-20-0_18-1_18-3

TAG-20-0_18-1_18-3

TAG-20-0_18-1_18-3

TAG-20-1_18-3_18-1

TAG-20-1_18-3_18-1

TAG-20-1_18-3_18-1

TAG-20-1_18-3_18-3

TAG-20-1_18-3_18-3

TAG-18-1_18-1_18-0

TAG-18-1_18-1_18-0

TAG-18-1_18-1_18-1

TAG-18-1_18-1_18-2

TAG-18-1_18-1_18-2

Compound group

MGDG A.t.

MGDG A.t.

MGDG A.t.

MGDG A.t.

MGDG A.t.

std DAG

std DAG

TAG A.t.

TAG A.t.

TAG A.t.

TAG A.t.

TAG A.t.

TAG A.t.

TAG A.t.

TAG A.t.

TAG A.t.

TAG A.t.

TAG A.t.

TAG A.t.

TAG A.t.

Table 1 (continued)

900.4

900.4

902.4

904.4

904.4

922.6

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

926.6 922.6

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

MS1 Res

926.6

926.6

928.6

928.6

928.6

686

686

742

744

746

748

764

Prec ion

601.4

603.4

603.4

603.4

605.4

595.4

627.4

599.4

627.4

631.4

599.4

629.4

633.4

341

385

563

565

567

569

585

Prod ion

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

MS2 Res

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

Dwell

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

Fragmentor

22

22

22

22

22

22

22

22

22

22

22

22

22

16

16

8

8

8

8

8

Collision energy

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

Cell accelerator voltage

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Polarity

216 Vale´rie Gros et al.

TAG-18-1_18-3_18-1

TAG-18-1_18-3_18-1

TAG-18-1_18-3_18-2

TAG-18-1_18-3_18-2

TAG-18-1_18-3_18-2

TAG-18-1_18-3_18-3

TAG-18-1_18-3_18-3

TAG-18-2_18-3_18-3

TAG-18-2_18-3_18-3

TAG-18-3_18-3_18-3

TAG-16-0_18-0_18-1

TAG-16-0_18-0_18-1

TAG-16-0_18-0_18-1

TAG-16-0_18-1_18-1

TAG-16-0_18-1_18-1

TAG-16-0_18-2_18-1

TAG-16-0_18-2_18-1

TAG-16-0_18-2_18-1

TAG-16-0_18-3_18-1

TAG-16-0_18-3_18-1

TAG-16-0_18-3_18-1

TAG-16-0_18-2_18-3

TAG-16-0_18-2_18-3

TAG-16-0_18-2_18-3

TAG-16-0_18-3_18-3

TAG A.t.

TAG A.t.

TAG A.t.

TAG A.t.

TAG A.t.

TAG A.t.

TAG A.t.

TAG A.t.

TAG A.t.

TAG A.t.

TAG A.t.

TAG A.t.

TAG A.t.

TAG A.t.

TAG A.t.

TAG A.t.

TAG A.t.

TAG A.t.

TAG A.t.

TAG A.t.

TAG A.t.

TAG A.t.

TAG A.t.

TAG A.t.

TAG A.t.

868.6

870.4

870.4

870.4

872.4

872.4

872.4

874.4

874.4

874.4

876.4

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

878.4 876.4

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

878.4

878.4

890.4

892.4

892.4

894.4

894.4

896.4

896.4

896.4

898.4

898.4

595.4

573.4

575.4

597.4

573.4

577.4

599.4

575.4

577.4

601.4

577.4

603.4

577.4

579.4

605.4

595.4

595.4

597.4

595.4

599.4

597.4

599.4

601.4

599.4

603.4

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

22

22

22

22

22

22

22

22

22

22

22

22

22

22

22

22

22

22

22

22

22

22

22

22

22

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

(continued)

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Extraction of Lipids 217

TAG-16-0_18-3_18-3

TAG-16-1_18-3_18-3

TAG-16-1_18-3_18-3

TAG-16-0_18-1_16-0

TAG-16-0_18-1_16-0

TAG-16-0_18-2_16-0

TAG-16-0_18-2_16-0

TAG-16-0_18-3_16-0

TAG-16-0_18-3_16-0

TAG-16-0_18-3_16-1

TAG-16-0_18-3_16-1

TAG-16-0_18-3_16-1

TAG-16-1_16-1_18-3

TAG-16-1_16-1_18-3

TAG-16-0_16-3_18-3

TAG-16-0_16-3_18-3

TAG-16-0_16-3_18-3

TAG A.t.

TAG A.t.

TAG A.t.

TAG A.t.

TAG A.t.

TAG A.t.

TAG A.t.

TAG A.t.

TAG A.t.

TAG A.t.

TAG A.t.

TAG A.t.

TAG A.t.

TAG A.t.

TAG A.t.

TAG A.t.

TAG A.t.

DGDG-36-0

DGDG-36-1

DGDG A.t.

DGDG A.t.

Segment 2: from 7 to 16 min

Compound name

Compound group

Table 1 (continued)

964

966

840.4

840.4

840.4

Widest

Widest

Widest

Widest

Widest

Widest

Widest

842.4 842.4

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

MS1 Res

844.4

844.4

844.4

846.4

846.4

848.4

848.4

850.4

850.4

866.4

866.4

868.6

Prec ion

623

625

545.4

567.4

573.4

547.4

571.4

547.4

571.4

573.4

551.4

573.4

551.4

575.4

551.4

577.4

571.4

595.4

573.4

Prod ion

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

MS2 Res

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

Dwell

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

Fragmentor

8

8

22

22

22

22

22

22

22

22

22

22

22

22

22

22

22

22

22

Collision energy

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

Cell accelerator voltage

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Polarity

218 Vale´rie Gros et al.

DGDG-36-2

DGDG-36-3

DGDG-36-4

DGDG-36-5

DGDG-36-6

DGDG-34-0

DGDG-34-1

DGDG-34-2

DGDG-34-3

DGDG-34-4

DGDG-34-5

DGDG-34-6

DGDG-32-1

DGDG-32-2

DGDG-32-3

MGDG-36-1

MGDG-36-2

MGDG-36-4

MGDG-36-5

MGDG-36-6

MGDG-34-0

MGDG-34-3

MGDG-34-4

MGDG-34-5

MGDG-34-6

DGDG A.t.

DGDG A.t.

DGDG A.t.

DGDG A.t.

DGDG A.t.

DGDG A.t.

DGDG A.t.

DGDG A.t.

DGDG A.t.

DGDG A.t.

DGDG A.t.

DGDG A.t.

DGDG A.t.

DGDG A.t.

DGDG A.t.

MGDG A.t.

MGDG A.t.

MGDG A.t.

MGDG A.t.

MGDG A.t.

MGDG A.t.

MGDG A.t.

MGDG A.t.

MGDG A.t.

MGDG A.t.

764

766

768

770

776

792

794

796

800

802

904

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

908 906

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

926

928

930

932

934

936

938

954

956

958

960

962

Unit Unit

585

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

587

589

591

597

613

615

617

621

623

563

565

567

585

587

589

591

593

595

597

613

615

617

619

621

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

8

8

8

8

8

8

8

8

8

8

8

8

8

8

8

8

8

8

8

8

8

8

8

8

8

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

(continued)

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Extraction of Lipids 219

MGDG-32-0

MGDG-32-1

MGDG-32-2

MGDG-32-3

PE-36-1

PE-36-2

PE-36-3

PE-36-4

PE-36-5

PE-36-6

PE-34-1

PE-34-2

PE-34-3

PE-34-4

PE-32-1

PE-32-2

PE-36-0

MGDG A.t.

MGDG A.t.

MGDG A.t.

MGDG A.t.

PE A.t.

PE A.t.

PE A.t.

PE A.t.

PE A.t.

PE A.t.

PE A.t.

PE A.t.

PE A.t.

PE A.t.

PE A.t.

PE A.t.

std PE

DPG_72-7

DPG_72-7

DPG A.t.

DPG A.t.

Segment 3: from 16 to 40 min

Compound name

Compound group

Table 1 (continued)

1450

1450

748

688

690

Widest

Widest

Widest

Widest

Widest

Widest

Widest

714 712

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

MS1 Res

716

718

736

738

740

742

744

746

742

744

746

748

Prec ion

695

697

607

547

549

571

573

575

577

595

597

599

601

603

605

563

565

567

569

Prod ion

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

MS2 Res

50

50

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

Dwell

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

Fragmentor

44

44

20

20

20

20

20

20

20

20

20

20

20

20

20

8

8

8

8

Collision energy

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

Cell accelerator voltage

Negative

Negative

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Polarity

220 Vale´rie Gros et al.

DPG_72-7

DPG_72-8

DPG_72-8

DPG_72-8

DPG_72-8

DPG_72-8

DPG_72-9

DPG_72-9

DPG_72-9

DPG_72-9

DPG_72-10

DPG_72-10

DPG_72-10

DPG_72-11

DPG_72-11

DPG_72-12

DPG_70-7

DPG_70-7

DPG_70-7

DPG_70-7

DPG_70-7

DPG_70-8

DPG_70-8

DPG_70-8

DPG A.t.

DPG A.t.

DPG A.t.

DPG A.t.

DPG A.t.

DPG A.t.

DPG A.t.

DPG A.t.

DPG A.t.

DPG A.t.

DPG A.t.

DPG A.t.

DPG A.t.

DPG A.t.

DPG A.t.

DPG A.t.

DPG A.t.

DPG A.t.

DPG A.t.

DPG A.t.

DPG A.t.

DPG A.t.

DPG A.t.

DPG A.t.

1420

1420

1420

1422

1422

1422

1422

1422

1440

1442

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

1444 1442

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

1444

1444

1446

1446

1446

1446

1448

1448

1448

1448

1448

1450

671

691

693

669

671

673

693

695

691

691

693

691

693

695

691

693

695

697

691

693

695

697

699

693

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

50

50

50

50

50

50

50

50

50

50

50

50

50

50

50

50

50

50

50

50

50

50

50

50

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

44

44

44

44

44

44

44

44

44

44

44

44

44

44

44

44

44

44

44

44

44

44

44

44

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

(continued)

Negative

Negative

Negative

Negative

Negative

Negative

Negative

Negative

Negative

Negative

Negative

Negative

Negative

Negative

Negative

Negative

Negative

Negative

Negative

Negative

Negative

Negative

Negative

Negative

Extraction of Lipids 221

Compound name

DPG_70-8

DPG_70-9

DPG_70-9

PA-36-1

PA-36-2

PA-36-3

PA-36-4

PA-36-5

PA-36-6

PA-34-0

PA-34-1

PA-34-2

PA-34-3

PA-34-4

PA-32-2

PC-36-1

PC-36-2

PC-36-3

PC-36-4

PC-36-5

Compound group

DPG A.t.

DPG A.t.

DPG A.t.

PA A.t.

PA A.t.

PA A.t.

PA A.t.

PA A.t.

PA A.t.

PA A.t.

PA A.t.

PA A.t.

PA A.t.

PA A.t.

PA A.t.

PC A.t.

PC A.t.

PC A.t.

PC A.t.

PC A.t.

Table 1 (continued)

780

782

784

786

788

662

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

688 686

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

MS1 Res

690

692

694

710

712

714

716

718

720

1418

1418

1420

Prec ion

184

184

184

184

184

547

571

573

575

577

579

595

597

599

601

603

605

669

691

669

Prod ion

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

MS2 Res

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

50

50

50

Dwell

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

Fragmentor

34

34

34

34

34

16

16

16

16

16

16

16

16

16

16

16

16

44

44

44

Collision energy

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

Cell accelerator voltage

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Negative

Negative

Negative

Polarity

222 Vale´rie Gros et al.

PC-36-6

PC-34-0

PC-34-1

PC-34-2

PC-34-3

PC-34-4

PC-32-0

PC-32-1

PC-32-2

PG-36-1

PG-36-2

PG-36-3

PG-36-4

PG-36-5

PG-36-6

PG-34-0

PG-34-1

PG-34-2

PG-34-3

PG-34-4

PG-32-0

PG-32-1

PI-36-1

PI-36-2

PC A.t.

PC A.t.

PC A.t.

PC A.t.

PC A.t.

PC A.t.

PC A.t.

PC A.t.

PC A.t.

PG A.t.

PG A.t.

PG A.t.

PG A.t.

PG A.t.

PG A.t.

PG A.t.

PG A.t.

PG A.t.

PG A.t.

PG A.t.

PG A.t.

PG A.t.

PI A.t.

PI A.t.

880

882

738

740

760

762

764

766

768

784

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

788 786

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

790

792

794

730

732

734

754

756

758

760

762

778

603

605

549

551

571

573

575

577

579

595

597

599

601

603

605

184

184

184

184

184

184

184

184

184

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

50

50

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

12

12

16

16

16

16

16

16

16

16

16

16

16

16

16

34

34

34

34

34

34

34

34

34

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

(continued)

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Extraction of Lipids 223

Compound name

PI-36-3

PI-36-4

PI-36-5

PI-36-6

PI-34-0

PI-34-1

PI-34-2

PI-34-3

PI-34-4

PI-32-0

PI-32-1

PI-32-2

PS-44-3

PS-44-4

PS-42-2

PS-42-3

PS-42-4

PS-42-9

PS-42-10

PS-42-11

Compound group

PI A.t.

PI A.t.

PI A.t.

PI A.t.

PI A.t.

PI A.t.

PI A.t.

PI A.t.

PI A.t.

PI A.t.

PI A.t.

PI A.t.

PS A.t.

PS A.t.

PS A.t.

PS A.t.

PS A.t.

PS A.t.

PS A.t.

PS A.t.

Table 1 (continued)

854

856

858

868

870

872

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

898 896

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

MS1 Res

824

826

828

848

850

852

854

856

872

874

876

878

Prec ion

669

671

673

683

685

687

711

713

547

549

551

571

573

575

577

579

595

597

599

601

Prod ion

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

MS2 Res

30

30

30

30

30

30

30

30

50

50

50

50

50

50

50

50

50

50

50

50

Dwell

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

Fragmentor

20

20

20

20

20

20

20

20

12

12

12

12

12

12

12

12

12

12

12

12

Collision energy

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

Cell accelerator voltage

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Polarity

224 Vale´rie Gros et al.

PS-40-1

PS-40-2

PS-40-3

PS-38-3

PS-36-1

PS-36-2

PS-36-3

PS-36-4

PS-36-5

PS-36-6

PS-34-0

PS-34-1

PS-34-2

PS-34-3

PS-34-4

PS-32-0

PS-32-1

PS-32-2

SQDG-36-1

SQDG-36-2

SQDG-36-3

SQDG-36-4

SQDG-36-5

PS A.t.

PS A.t.

PS A.t.

PS A.t.

PS A.t.

PS A.t.

PS A.t.

PS A.t.

PS A.t.

PS A.t.

PS A.t.

PS A.t.

PS A.t.

PS A.t.

PS A.t.

PS A.t.

PS A.t.

PS A.t.

SQDG A.t.

SQDG A.t.

SQDG A.t.

SQDG A.t.

SQDG A.t.

839

841

843

845

847

732

734

736

756

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

760 758

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

762

764

780

782

784

786

788

790

814

842

844

846

225

225

225

225

225

547

549

551

571

573

575

577

579

595

597

599

601

603

605

629

657

659

661

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

30

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

135

56

56

56

56

56

20

20

20

20

20

20

20

20

20

20

20

20

20

20

20

20

20

20

7

7

7

7

7

7

(continued)

Negative

Negative

Negative

Negative

Negative

Positive

Positive

Positive

7 7

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

Positive

7

7

7

7

7

7

7

7

7

7

7

7

7

7

7

Extraction of Lipids 225

SQDG-36-6

SQDG-34-1

SQDG-34-2

SQDG-34-3

SQDG-34-4

SQDG-34-5

SQDG-34-6

SQDG-32-1

SQDG-32-2

SQDG-32-3

SQDG-34-0

SQDG A.t.

SQDG A.t.

SQDG A.t.

SQDG A.t.

SQDG A.t.

SQDG A.t.

SQDG A.t.

SQDG A.t.

SQDG A.t.

SQDG A.t.

std SQDG

821

787

789

791

809

811

813

815

817

819

837

Prec ion

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

Widest

MS1 Res

225

225

225

225

225

225

225

225

225

225

225

Prod ion

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

Unit

MS2 Res

30

30

30

30

30

30

30

30

30

30

30

Dwell

135

135

135

135

135

135

135

135

135

135

135

Fragmentor

56

56

56

56

56

56

56

56

56

56

56

Collision energy

7

7

7

7

7

7

7

7

7

7

7

Cell accelerator voltage

Negative

Negative

Negative

Negative

Negative

Negative

Negative

Negative

Negative

Negative

Negative

Polarity

Prec Ion precursor ion, Res resolution, Prod Ion product ion, DAG diacylglycerol, DGDG digalactosyldiacylglycerol, MGDG monogalactosyldiacylglycerol, TAG triacylglycerol, PE phosphatidylethanolamine, DPG diphosphatidyldiacylglycerol, PA phosphatidic acid, PC phosphatidylcholine, PG phosphatidylglycerol, PI phosphatidylinositol, PS phosphatidylserine, SQDG sulfoquinovosediacylglycerol

Compound name

Compound group

Table 1 (continued)

226 Vale´rie Gros et al.

Extraction of Lipids

227

Fig. 3 Diagram representing the strategy to quantify glycerolipids by LC-MS/MS. One internal standard is used for each chromatographic segment: DAG 18:0–22:6 is used to adjust DAG, TAG, MGDG, and DGDG amounts, PE 18:0–18:0 for PE amount and SQDG 16:0–18:0 for SQDG, DGTA, DGTS, DGCC, PA, PC, PG, PS, DPG, and PI amounts

These correction factors determined by lipid class are then applied to the molecule species of the sample to determine a corrected quantity of lipid (see Fig. 3 for a summary of the whole quantification process) (see Note 21): Qcor ðms LIPÞ = Q ðms LIPÞ × CF

4

Notes 1. Quartz wool correspond to pure SiO2 wool with fiber thickness from 4 to 12 μm. It does not react with solvent and does not contain lipids. 2. Rinse potters pestles with distilled water, then ethanol. 3. If the evaporator bath could warm: Put the set temperature at 40 °C maximum. It will decrease the evaporation time.

228

Vale´rie Gros et al.

4. To prepare the standard solution at 0.5 mg/mL: Put 25 mg of C15 in an Eppendorf tube of 1 mL. Add 1 mL of chloroform/ methanol 1/2 (v/v) and make hand agitation. Transfer this solution in a graduated flask of 50 mL. Complete until 50 mL with chloroform/methanol 1/2 (v/v). Put a glass cap and parafilm around. Make some inversions. Put some milliliters of this C15 solution in a little beaker (to avoid evaporation) and realize aliquots of 500 μL and store them at -20 °C. Take out of the freezer at least 20 min before use. 5. Prepare the solution at 4 °C to avoid overheating of the solution. Put 400 mL of methanol in a glass bottle, under agitation. Then, add slowly 10 mL of sulfuric acid. Put a glass cap and store at RT. 6. All the stock standard solutions need to be quantified by GC-FID to be sure the concentrations are accurate. If necessary volume will be adjusted, to be sure the standard solution will be at 1.25 μM for each standard. 7. Solubilize 5 mg of SQDG in 1 mL of chloroform in a glass vial HPLC type. Add a small spatula of PtO2 powder (catalyzer). Add a small bar magnet (4 mm). Label the level of liquid on the vial. Close the vial and add at the center of the lid 2 thin needles to allow hydrogen to get in and out. Needles need to be above the liquid. Prepare an Erlenmeyer flask with a lid insensitive to chloroform with two holes, one for the entrance of hydrogen and the other one for a “safety valve” constituted of a Pasteur pipette with a finger of cut latex glove. Put the HPLC vial into the Erlenmeyer flask with the gaz entrance of the Erlenmeyer flask connected to the first needle of the vial (use silicon tube and junctions). Put the Elen above a magnet stirrer under the fume hood. Open the hydrogen bottle or generator to deliver 3.5 bars maximum to fill the Erlenmeyer vial with hydrogen then reduce the pressure to 1 bar. Leave overnight under small stirring. The following day recover the lipid by eventually adding chloroform and collecting the liquid. Quantify and check the SQDG composition by methanolysis and GC-FID (Subheadings 3.2 and 3.3) 8. Here, settings are described for an HPLC Agilent 1200 and a triple quadripole Agilent 6460. Parameters might vary for other instruments. 9. Put the Corex tube at least 10 s in the liquid nitrogen before receiving the freeze-dried material. 10. Be careful: Samples should not be dry. If there is almost no liquid left, take out the tube from the bath before 5 min of incubation and continue the protocol. 11. Open the argon very slowly after putting Pasteur pipette into the liquid.

Extraction of Lipids

229

12. To avoid any contamination with the upper phase, use a propipette on top of the Pasteur pipette. Suck a little bit of air into the pipette, plunge it at the bottom of the tube, eject one or 2 bubbles that will reject any upper phase that might have penetrate into the pipette, and suck up the lower phase. 13. Put the aliquot of standard C15 at least 20 min at room temperature before use. No overthrow but gently agitation. 14. For 100 mg of plant tissue, a volume of 50 μL will be adequate for the methanolysis. For 100 million of algae cells, prefer a volume of 100 μL for the methanolysis. 15. The ES lipid extract is run in the middle of the sample list within each series of analyses. These ES extracts corresponded to a known lipid extract from Phaeodactylum tricornutum, Nannochloropsis gaditana, or Arabidopsis thaliana cell cultures qualified and quantified by TLC and GC-FID as described by [8]. 16. We verify that the method is within a linear range up to 15 nmol of total fatty acid (S1 Fig). 17. The distinct glycerophospholipid classes were eluted successively as a function of the polar head group. Under these conditions, they were eluted in the following order: TAG (triacylglycerol), DAG (diacylglycerol), MGDG (monogalactosyldiacylglycerol), DGDG (digalactosyldiacylglycerol), PE (phosphatidylethanolamine), DGTS (diacylglycerylcarboxy-N-hydroxymethylhomoserine), DGTA (diacylglycerylhydroxymethyl-N,N,N-trimethyl-β-alanine), PG (phosphatidylglycerol), PI (phosphatidylinositol), SQDG (sulfoquinovosyldiacylglycerol), PS (phosphatidylserine), PC (phosphatidylcholine), DPG (diphosphatidylglycerol), and PA (phosphatidic acid). 18. For a 6460 triple quadrupole mass spectrometer (Agilent) equipped with a Jet stream electrospray ion source, the source parameters are the following settings: Drying gas heater: 230 ° C, Drying gas flow 10 L/min, Sheath gas heater: 200 °C, Sheath gas flow: 10 L/min, Nebulizer pressure: 25 psi, Capillary voltage: ±4000 V, Nozzle voltage: ±2000. Nitrogen was used as collision gas. 19. To use this method, all the different molecular species (with precise determination of fatty acids esterified to the glycerol backbone in each lipid class) of the studied organism glycerolipidome need to be known. Only lipid transitions that are entered in the method will be measured. It is a targeted method. 20. The corresponding internal standard is used in function of the retention time in order to have the molecule and the internal standard in roughly the same environment. In our conditions,

230

Vale´rie Gros et al.

DAG18:0/22:6 will be used for DAG, TAG, MGDG, and DGDG species, PE 18:0/18:0 for PE species, and SQDG 18: 0/18:0 for PG, PC, PA, PI, DPG, PS, DGTS, DGTA, and DGCC. 21. This quantification method does not give absolute quantification but gives corrected value that are close to the value measured by other method such as GC-FID that is known to be robust and reliable [9]

Acknowledgments This protocol was developed at the LIPANG (Lipid analysis in Grenoble) platform hosted by the LPCV and was supported by the Rhoˆne-Alpes-Auvergne Region, the FEDER fund, and Agence Nationale de la Recherche (ANR-10-LABEX-04 GRAL and ANR-17-EURE-0003 CBH-EUR-GS). References 1. Abida H, Dolch L-J, Meı¨ C et al (2015) Membrane glycerolipid remodeling triggered by nitrogen and phosphorus starvation in Phaeodactylum tricornutum. Plant Physiol l167:118– 136. https://doi.org/10.1104/pp.114. 252395 2. Degraeve-Guilbault C, Bre´he´lin C, Haslam R et al (2017) Glycerolipid characterization and nutrient deprivation-associated changes in the green Picoalga Ostreococcus tauri1. Plant Physiol 173:2060–2080. https://doi.org/10. 1104/pp.16.01467 3. Mu¨hlroth A, Winge P, El Assimi A et al (2017) Mechanisms of phosphorus acquisition and lipid class remodeling under P limitation in a marine microalga. Plant Physiol 175:1543–1559. https://doi.org/10.1104/pp.17.00621 4. Kumar Sharma A, Mu¨hlroth A, Jouhet J et al (2020) The Myb-like transcription factor phosphorus starvation response (PtPSR) controls conditional P acquisition and remodelling in marine microalgae. New Phytol 225:2380– 2395. https://doi.org/10.1111/nph.16248

5. Li-Beisson Y, Thelen JJ, Fedosejevs E et al (2019) The lipid biochemistry of eukaryotic algae. Progr Lipid Res 74:31–68. https://doi. org/10.1016/j.plipres.2019.01.003 6. Folch J, Ascoli I, Lees M et al (1951) Preparation of lipid extracts from brain tissue. J Biol Chem 191:833–841 7. Bligh EG, Dyer WJ (1959) A rapid method of total lipid extraction and purification. Can J Biochem Physiol 37:911–917. https://doi.org/10. 1139/o59-099 8. Jouhet J, Lupette J, Clerc O et al (2017) LC-MS/MS versus TLC plus GC methods: consistency of glycerolipid and fatty acid profiles in microalgae and higher plant cells and effect of a nitrogen starvation. PLoS ONE 12:e0182423. https://doi.org/10.1371/jour nal.pone. 0182423 9. Dodds ED, McCoy MR, Rea LD et al (2005) Gas chromatographic quantification of fatty acid methyl esters: flame ionization detection vs. electron impact mass spectrometry. Lipids 40:419–428. https://doi.org/10.1007/ s11745-006-1399-8

Chapter 14 Quantitative Assessment of the Chloroplast Lipidome Vale´rie Gros and Juliette Jouhet Abstract In plants and algae, photosynthetic membranes have a unique lipid composition. They differ from all other cellular membranes by their very low amount of phospholipids, besides some phosphatidylglycerol (PG), and high proportion of glycolipids. These glycolipids are the uncharged galactolipids, that is, mono- and digalactosyldiacylglycerol (MGDG and DGDG), and an anionic sulfolipid, that is, sulfoquinovosyldiacylglycerol (SQDG). In all photosynthetic membranes analyzed to date, from cyanobacteria to algae, protists, and plants, the lipid quartet constituted by MGDG, DGDG, SQDG, and PG has been highly conserved, but the composition in fatty acids of these lipids can vary a lot from an organism to another. To better understand the chloroplast biogenesis, it is therefore essential to know their lipid content. Establishing chloroplast lipidome requires first to purify chloroplast from plant or algae tissue. Here we describe the methods to extract the lipid, quantify the lipid amount of the chloroplast, and qualify and quantify the different lipid classes that might be present in these fractions. Key words Lipid, Extraction, 2D thin layer chromatography, Quantification, Gas chromatography, Mass spectrometry, Fatty acid methyl ester

1

Introduction Chloroplasts have emerged from a long and complex evolutionary history, originating from the engulfment of ancestral cyanobacteria within a eukaryotic host cell. Chloroplast membranes have an original lipid composition conserved from cyanobacteria to higher plants. They contain a high proportion of two uncharged glycolipids, namely, the galactoglycerolipids, that is, mono- and di-galactosyldiacylglycerol (MGDG and DGDG, respectively), and an anionic sulfolipid, that is, sulfoquinovosediacylglycerol (SQDG). A remarkable feature of the evolution from cyanobacteria to higher plants is the conservation of MGDG, DGDG, SQDG, and phosphatidylglycerol (PG), the major phospholipid of thylakoids. In higher plant, galactolipids result from the galactosylation of diacylglycerol (DAG) by UDP-galactose. This step of synthesis

Eric Mare´chal (ed.), Plastids: Methods and Protocols, Methods in Molecular Biology, vol. 2776, https://doi.org/10.1007/978-1-0716-3726-5_14, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

231

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occurs in the plastid envelope. DAG substrate is either de novosynthesized in the chloroplast by acylation of glycerol-3-phosphate or formed by a complex conversion of PC from the endoplasmic reticulum (ER) that requires a transfer of lipid between the ER and the chloroplast. The DAG backbone composition of the galactolipid reflects both its chloroplastic and its endomembrane origin (for a review, see [1, 2]). Thus, to understand the chloroplast lipid metabolism and its integration within the cell physiology, it is essential to be able to quantify and qualify, that is, position the fatty acid on the glycerol backbone, the chloroplast lipidome. Here, we describe how to quantify and qualify plastid lipidome starting from purified plastid fraction as described in this book in previous chapter. The procedure starts from the lipid extraction of the purified fractions, followed by the total fatty acid quantification. Lipid classes are then isolated by thin layer chromatography (TLC) and collected either to determine the DAG backbone by mass spectrometry or to quantify the amount of lipid by gas chromatography coupled to flame ionization detector (GC-FID). This complete procedure allows to establish the complete lipidome of plastid fraction.

2

Materials Prepare and store all reagents at room temperature. Always use glass vessels and never plastic with organic solvents. Wash all vessels without detergent but with distilled water, then ethanol. All solvents must be for Analysis grade except Hexane that must be for GC grade. Chloroform should be ethanol stabilized. Diligently follow all waste disposal regulations when disposing waste materials.

2.1

Lipid Extractions

1. Argon: Argon is used to dry samples by applying a flow of gas using thin glass Pasteur pipettes or metal needles (see Fig. 1 and Note 1). 2. Chlorofom, ethanol stabilized, purity ≥99.9% (see Note 2). 3. Methanol: purity ≥99.9%. 4. Chloroform:methanol 1:2 (v:v) 5. Ultrapure water (prepared by purifying deionized water, to attain a sensitivity of 18 MΩ-cm at 25 °C). 6. Hemolysis glass tube (volume 5 mL) for single use.

2.2

Methanolysis

1. Pyrex tubes of 20 mL (160 mm × 16 mm), tubes in borosilicate glass (high thermal resistance) with screw top. Screw caps in phenolic resin with rubber seal lined with inert polytetrafluoroethylene (PTFE).

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Fig. 1 Example of a system to remove solvents by applying a flow of argon. Argon is applied at the top of the tube, using a gas bottle, with a flow set at a maximum pressure of 2 bar. This system can warm the lipid extract to speed up the evaporation of solvents

2. Standard C15 (fatty acid with 15 carbons) solution: Powder of pentadecanoic acid (99%), stored at RT. Prepare a solution at 0.5 mg/mL in chloroform:methanol (1:2, v:v) (see Note 3). 3. Dry-bath heating block. 4. Methanolysis buffer: Sulfuric acid (H2SO4 24N) 2.5% in Methanol (v/v) (see Note 4). 5. Ultrapure water (prepared by purifying deionized water, to attain a sensitivity of 18 MΩ cm at 25 °C). 6. Hexane for gas chromatography (GC) analysis. 7. Hemolysis glass tube (volume 5 mL) for single use. 2.3 Quantification by Gas Chromatography– Flame Ionization Detector (GC-FID)

1. Vials for automatic sampler with an insert of 250 μL and screw caps 9 mm with PTFE seal. 2. Hexane for GC analysis. 3. Column BPX70 (70% Cyanopropyl Polysilphenylene-siloxane) for GC: length 30 m, internal diameter 0.22 mm, film thickness 0.25 μm. 4. Gas chromatography–flame ionization detector. 5. Commercial fatty acid methyl ester (FAME) standard solution to calibrate the GC-FID retention time.

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2D Thin Layer

1. Thin layer chromatography (TLC) plate, Silicagel 60 on glass 20 × 20 cm (see Note 5). 2. Glass tank for TLC migration. 3. Chloroform, ethanol stabilized purity ≥99.9% (see Note 2). 4. Argon. 5. Methanol, purity ≥99.9%. 6. ANS solution: Solubilize 8-anilinonaphthalene sulfonic acid (ANS) at 0.2% in methanol (w/v). Transfer this solution in a bottle with spray system such as Preval spray unit (aerosol) (see Note 6).

2.5 Lipid Class Identification by Mass Spectrometry

1. Chlorofom, ethanol stabilized, purity ≥99.9% (see Note 2). 2. Methanol: purity ≥99.9%. 3. Ultrapure water (prepared by purifying deionized water, to attain a sensitivity of 18 MΩ-cm at 25 °C). 4. MS buffer: 10 mM ammonium acetate in methanol. 5. Hemolysis glass tube (volume 5 mL) for single use. 6. Argon. 7. Ion trap mass spectrometer.

3

Methods

3.1 Lipid Extractions from Chloroplast Purified Fraction

1. Purify the chloroplast fraction as described in previous chapters of this book (see Note 7). Transfer the fraction into a hemolysis glass tube. Complete the volume of the fraction up to 2mL of water. Add 3.4 mL chloroform:methanol (1:2, v:v). Vortex 30 s. 2. Add 1.2 mL of chloroform. Blow argon carefully during 1 min inside the tube to mix up solvents and to remove oxygen. Incubate 10 min at room temperature. Centrifuge 10 min at 1000 g at room temperature to form the biphase. Collect the lower phase (organic phase) with a Pasteur pipette (see Note 8) and transfer it in a clean glass tube. 3. Repeat step 2 by adding 1.2 mL of chloroform in the glass tube containing the upper phase to re-extract the aqueous phase. Pull the two lower phases (organic phases) into the same glass tube of step 2. 4. Dry all the solvents by blowing argon on top of the liquid. Close the tube and store the lipid extract at -20 °C. Lipid extract is stable for several months.

Chloroplast Lipidome

3.2 Production of Fatty Acid Methyl Esters (FAMEs)

235

1. Set the heating block to 100 °C and pre-warm an aliquot of standard C15 solution (see Note 9). 2. Resuspend the lipid extract in 1 mL of chloroform. Transfer 10–100 μL of the lipid extract (see Note 10) into the methanolysis tube. 3. With a glass syringe, add 10 μL of C15 solution (5 μg/tube) in the methanolysis tube. 4. Add 3 mL of methanolysis buffer in methanolysis tubes. In each tube, there are C15 solution, lipid extract, and methanolysis buffer. Close tightly the glass tube. Incubate 1 h at 100 °C for the methanolysis reaction to occur (see Note 11). 5. Take the tube out of the hot block 5 min at room temperature to cool down the tube. Stop the reaction by adding 3 mL of water. 6. Add 3 mL of hexane to extract the FAMEs and vortex vigorously. Wait at least 20 min at room temperature to allow the biphase to separate correctly (see Note 12). Take the upper phase (hexane phase) containing the FAMEs, transfer it in a hemolysis glass tube, and dry it under argon (see Note 13). 7. Repeat step 6 to re-extract FAMEs from the methanol-water phase by adding again 3 mL of hexane in the methanolysis tube. Pour the upper phase in the same hemolysis glass tube than the one used in step 6 and dry it under argon. 8. To concentrate the FAMEs at the bottom of the glass tube, rinse the tube wall with 200 μL of hexane. Allow the liquid to rest at the bottom of the tube and dry it gently under argon. 9. Store the FAMEs at -20 °C for few months or proceed with GC analysis.

3.3 Quantification of FAMEs by Gas Chromatography– Flame Ionization Detector (GC-FID)

1. Resuspend the FAMEs in 45 μL of hexane and transfer the FAMEs in an insert vial. Seal the vial tightly to avoid evaporation. 2. Inject 2 μL of the sample in the GC-FID on a BPX70 column. Nitrogen is used as carrier gas with 3.5 mL/min constant flow compensation, split ratio is 13.3:1, injection temperature of 200 °C, detector temperature of 280 °C and the oven temperature range start at 130 °C, hold for 7.5 min at 130 °C, ramp up to 180 °C at 3 °C/min, and hold 10 min at 180 °C. This will allow FAMEs separation from 12 C up to 24 C in function of the chain length and the number of desaturation. 3. Each FAME is identified by comparison of its retention time with those of standards.

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4. FID response is dependent on the mass of the FAME; therefore, each FAME will be quantified by the surface peak method using C15 surface peak for calibration with the following equation: Quantity in μg of FAME =

Area of FAME peak × Quantity in μg of C15 ð5 μgÞ : Area of C15 peak

5. By adding each FAME quantity and taking into account the volume used for the methanolysis, GC-FID analysis gives the total fatty acid content of the lipid extract as well as its fatty acid composition. 3.4 Lipid Class Separation by 2D Thin Layer Chromatography (TLC)

1. Prepare migration tank with 99 mL of the solvent mix for the first dimension: chloroform:methanol:water 65:25:4 (v:v:v). Close the tank and wait 30 min to saturate the tank atmosphere. 2. Resuspend the lipid extract with chloroform. Take the volume corresponding to 300 μg of total fatty acid that was calculated by GC-FID analysis and transfer it in a new glass tube. Remove all the chloroform by applying a flow of argon to evaporate this solvent. 3. Resuspend the lipid extract corresponding to the 300 μg of total fatty acid in 40 μL of chloroform, and spot all the lipid extract with a Pasteur pipette on the TLC plate at 2.5 cm of both sides (see Fig. 2 and Note 14). Dry between each drop with argon. Do it again 3 times by rinsing the wall of the tube with 20 μL of chloroform and once with 10 μL to be sure all the sample is put down on the TLC plate. 4. Insert the TLC plate inside the tank in the right orientation: Maximum two plates per tank (see Note 15). Remove the plate when the migration front is at 2 cm from the edge (around 1.5 h). 5. Leave the TLC plate under the fume hood for 10 min to dry the solvent. Put it under a cloche saturated with argon for at least 10 min. Seal the cloche and leave the TLC plate overnight. 6. Prepare migration tank with 95 mL of the solvent mix for the second dimension: chloroform:acetone:methanol:acetic acid: water 50:20:10:10:5 (v:v:v:v:v). Close the tank and wait 30 min to saturate the tank atmosphere. 7. Insert the TLC plate inside the tank in the right orientation. Remove the plate when the migration front is at 2 cm from the edge (around 1.5 h). Store the TLC plate under the fume hood for 30 min to dry it.

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Fig. 2 Typical 2D TLC plates of Arabidopsis leave (a) and chloroplast (b) lipid extracts. The deposition spot is marked ( ), and dimensions are indicated by arrows on the side of the plates. Lipids are visualized after spraying with ANS and exposure under UV light. Glycerolipid spots are circled in black and named. MGDG: monogalactosyldiacylglycerol, DGDG: digalactosyldiacylglycerol, SQDG: sulfoquinovosediacylglycerol, PG: phosphatidylglycerol, DPG: diphosphatidylglycerol, PA: phosphatidic acid, PE: phosphatidylethanolamine, PC: phosphatidylcholine, PI: phosphatidylinositol, PS: phosphatidylserine

8. Stain the lipids by spraying ANS under the fume hood (see Note 16). Wait few minutes to dry the methanol and visualize the lipids under UV light. 9. Mark the lipid spots by drawing the spot contour with a pencil (see Fig. 2). Identify each lipid spot by mass spectrometry or quantify them by methanolysis and GC-FID. 10. To identify the lipid spot, proceed with Subheading 3.5. To quantify the lipid spot by methanolysis and GC-FID, scratch the silica containing the spot with a small spatula and pour the silica into a methanolysis tube and follow the same procedure from Subheading 3.2, steps 2 up to 4. By adding each FAME quantity and taking into account the quantity used for the TLC deposition, GC-FID analysis gives the total fatty acid content of each lipid class in the lipid extract as well as the fatty acid composition of each lipid class and therefore reveals the chloroplast lipidome. 3.5 Lipid Class Identification by Mass Spectrometry

1. Scratch the silica containing the spot with a small spatula and pour the silica into a hemolysis tube. 2. Extract the lipid from the silica: Add 1.35 mL of chloroform: methanol 1:2 (v:v), vortex, add 450 μL of chloroform, vortex, add 800 μL of water, vortex, wait 10 min, and centrifuge

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10 min at 1000 g. Take the bottom phase (organic phase) without collecting the silica (see Note 17) and pour it in a new hemolysis tube. Re-extract the aqueous phase by adding 1.2 mL of chloroform, vortex, wait 10 min, and centrifuge 10 min at 1000 g. Take the bottom phase (organic phase) without collecting the silica and pool it with the first organic phase in the hemolysis tube. Dry it under argon. Lipids can be stored at -20 °C or proceed with mass spectrometry analysis. 3. Resuspend the extracted lipid spot with 200 μL of MS buffer. Inject by direct infusion into the mass spectrometer at 0.7 μL/ min. Lipids are identified by MS/MS analysis with their precursor ion or by neutral loss analyses as indicated in Table 1 (see Note 18). For establishing the sn-1 and sn-2 position of the fatty acid on the glycerol backbone, lipid will be fragmented as described in Table 2. By comparing the height of the fragment described in Table 2, fatty acid positioning can be established.

Table 1 Precursor and neutral loss scans utilizing characteristic fragments generated by electrospray ionization for analysis of polar complex glycerolipids

Analyzed lipids

Ion Polarity analyzed

Scan mode

References

Phospholipids Phosphatidylcholines

+

[M + H]+

Neutral loss of m/z 59

[3]

Phosphatidylethanolamines

+

[M + H]+

Neutral loss of 141

[4]

+

+

Neutral loss of 185

[4]

Neutral loss of 189

[5]

Precursors of m/z 241

[6]

Phosphatidylserines Phosphatidylglycerols

+

[M + H]

+

[M + NH4] –

Phosphatidylinositols

+

[M – H]

Phosphatidic acids

+

[M + NH4]+ Neutral loss of 115

[7]

–

[M – H]–

Precursors of m/z 225

[8]

C20:5 acyl-Sulfoquinovosyldiacylglycerols –

[M – H]–

Precursors of m/z 509

[9]

Monogalactosyldiacylglycerols

[M + NH4]+ Neutral loss of 179

Nonphosphorous glycerolipid Sulfoquinovosyldiacylglycerols

Digalactosyldiacylglycerols Diacylglyceryl-N,N,Ntrimethylhomoserines

+ + +

+

[M + NH4] +

[M + H]

[7]

Neutral loss of 341

[10]

Precursors of m/z 236

[11]

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Table 2 Conditions for regiochemical assignment of polar complex lipids Ion Polarity Analyzed

Analyzed lipids

Regiochemical property

References

Phospholipids Phosphatidylcholines

–

[M-CH3]- [M-CH3– R2CHCO]-> [M-CH3– R1CHCO]-

[12]

Phosphatidylethanolamines

–

[M – H]–

[R2COO-] > [R1COO-]

[13]

Phosphatidylglycerols

–

[M – H]–

[M-H-R2COOH]- > [M-H-R1COOH]-

[14]

Phosphatidylinositols

–

[M – H]–

[M-H-R2COOH]- > [M-H-R1COOH]-

[6]

Sulfoquinovosyldiacylglycerols

–

[M – H]–

[M-H-R1COOH]- > [M-H-R2COOH]-

[15]

C20:5 Acylsulfoquinovosyldiacylglycerols

–

[M – H]–

[M-H-R1COOH]- > [M-H-R2COOH]-

[9]

Monogalactosyldiacylglycerols

+

[M + Na]+

[M+Na-R1COO-]+ > [M+Na-R2COO-]+

[16]

Digalactosyldiacylglycerols

+

[M + Na]+

[M+Na-R1COO-]+ > [M+Na R2COO-]+

[16]

Diacylglyceryl-N,N,Ntrimethylhomoserines

+

[M + H]+

[M+H-R2COOH]+ > [M+H-R1COOH]+

By analogy with phospholipid diacylglycerol backbone [17]

Nonphosphorous glycerolipid

4

Notes 1. If the drying device is able to heat, set the maximum temperature to 40 °C. The evaporation will be quicker, without degradation or evaporation of the lipids. 2. Chloroform needs to be stabilized with ethanol and not with amylene (be careful to the supplier choice). It will affect the TLC migration pattern. 3. Standard C15 solution will be the reference solution for quantification. The concentration of this solution needs to be accurate. To prepare the standard solution at 0.5 mg/mL: put

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25.0 mg of C15 in a graduated flask of 50 mL. Complete until 50 mL with chloroform:methanol 1:2 (v:v). Put a glass cap and parafilm around. Make some inversions until the C15 is fully dissolved. Aliquot the solution by 500 μL in a glass tube sealed with cap containing PTFE seal and store them at -20 °C for months. 4. For preparing the methanolysis buffer, put 400 mL of methanol in a glass bottle, under agitation in ice. Then, add slowly 10 mL of sulfuric acid 24N to avoid heating of the methanol. Once the temperature of the solution is stabilized, put a glass cap and store at room temperature. 5. TLC plates from different suppliers should be compared for the efficiency of the separation. 6. Powder of ANS is stored at room temperature. Dissolve 0.2 g in 100 mL of Methanol. Be careful to protect the ANS solution from light: use a dark bottle or wrap it with foil. 7. To have enough material for the whole analysis, it is safer to start with a fraction containing at least 1 mg of protein or 500 μg of chlorophyll. 8. To avoid pipetting the upper phase, suck up some air into the Pasteur Pipette, plunge the pipette at the bottom of the tube and blow until an air bubble get out of the pipette to remove any leftover of the upper phase that might have get into the pipette by capillarity. Start to collect the bottom phase. 9. Put the aliquot of standard C15 at least 20 min at room temperature before use. No overthrow but gentle agitation. 10. If protein content is used as reference, for a chloroplast fraction containing 1 mg of protein, a volume of 50 μL will be adequate for the methanolysis. If chlorophyll content is used as reference, for a chloroplast fraction containing 1 mg of chlorophyll, a volume of 25 μL will be adequate for the methanolysis. 11. Methanolysis reaction is a transesterification reaction: Fatty acids are hydrolyzed from the glycerol backbone and re-esterified with methanol, producing fatty acid methyl esters (FAMEs). 12. At this step, the biphasic mixture can be stored a few days at 4 ° C if necessary. 13. At this step, do not heat up the tube and be careful that the room temperature is below 30 °C to avoid volatilization of short chain FAMEs. 14. Choose a Pasteur pipette with a flat tip. Set down gently the pipette on the silica because it is very brittle.

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15. Maximum two plates per tank that will be back to back to avoid any cross contamination. Don’t move the tank after putting the plates. 16. Put the TLC straight over a cardboard to protect the fume hood. Hold the sprayer at 30 cm of the plate and spray uniformly all over the plates. 17. Silica will be present at the interface of the aqueous and the organic phase and can be quite thick. If there is too much silica, the volume of solvent can be increased as long as the ratio are respected. To avoid pipetting the upper phase and the silica, suck up some air into the Pasteur Pipette, plunge the pipette at the bottom of the tube, and blow until an air bubble get out of the pipette to remove any leftover of the upper phase and silica that might have get into the pipette by capillarity. Start to collect the bottom phase. 18. Parameters for the mass spectrometer source or fragmentation are dependent on the mass spectrometer. For an AmazonXL (Bruker), collision energy is setup to 50% and the isolation width is fixed to 1 Da in negative mode and to 2 Da and 3 Da for precursor ion [M + H]+ and [M + NH4]+, respectively. Adjustments should be made for different systems. References 1. Gue´guen N, Mare´chal E (2022) Origin of cyanobacterial thylakoids via a non-vesicular glycolipid phase transition and their impact on the Great Oxygenation Event. J Exp Bot 73:2721– 2734. https://doi.org/10.1093/jxb/erab429 2. Petroutsos D, Amiar S, Abida H et al (2014) Evolution of galactoglycerolipid biosynthetic pathways—from cyanobacteria to primary plastids and from primary to secondary plastids. Progr Lipid Res 54:68–85. https://doi.org/ 10.1016/j.plipres.2014.02.001 3. Domingues P, Amado FML, Santana-Marques MGO et al (1998) Constant neutral loss scanning for the characterization of glycerol phosphatidylcholine phospholipids. J Am Soc Mass Spectrom 9:1189–1195. https://doi.org/10. 1016/S1044-0305(98)00087-7 4. Bru¨gger B, Erben G, Sandhoff R et al (1997) Quantitative analysis of biological membrane lipids at the low picomole level by nanoelectrospray ionization tandem mass spectrometry. Proc Natl Acad Sci 94:2339–2344. https://doi.org/10.1073/pnas.94.6.2339 5. Taguchi R, Houjou T, Nakanishi H et al (2005) Focused lipidomics by tandem mass spectrometry. J Chromatogr B 823:26–36.

https://doi.org/10.1016/j.jchromb.2005. 06.005 6. Hsu F-F, Turk J (2000) Characterization of phosphatidylinositol, phosphatidylinositol-4phosphate, and phosphatidylinositol-4,5bisphosphate by electrospray ionization tandem mass spectrometry: a mechanistic study. J Am Soc Mass Spectrom 11:986–999. https:// doi.org/10.1016/S1044-0305(00)00172-0 7. Li-Beisson Y, Shorrosh B, Beisson F et al (2010) Acyl-lipid metabolism. Arabidopsis Book 8:e0133. https://doi.org/10.1199/ tab.0133 8. Welti R, Wang X, Williams TD (2003) Electrospray ionization tandem mass spectrometry scan modes for plant chloroplast lipids. Anal Biochem 314:149–152. https://doi.org/10. 1016/S0003-2697(02)00623-1 9. Naumann I, Klein BC, Bartel SJ et al (2011) Identification of sulfoquinovosyldiacyglycerides from Phaeodactylum tricornutum by matrix-assisted laser desorption/ionization QTrap time-of-flight hybrid mass spectrometry. Rapid Commun Mass Spectrom 25:2517– 2523. https://doi.org/10.1002/rcm.5137

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10. Moreau RA, Doehlert DC, Welti R et al (2008) The identification of mono-, di-, tri-, and tetragalactosyl-diacylglycerols and their natural estolides in oat kernels. Lipids 43:533–548. https://doi.org/10.1007/s11745-0083181-6 11. Benning C, Huang ZH, Gage DA (1995) Accumulation of a novel glycolipid and a betaine lipid in cells of Rhodobacter sphaeroides grown under phosphate limitation. Arch Biochem Biophys 317:103–111. https://doi.org/ 10.1006/abbi.1995.1141 12. Berdeaux O, Juaneda P, Martine L et al (2010) Identification and quantification of phosphatidylcholines containing very-long-chain polyunsaturated fatty acid in bovine and human retina using liquid chromatography/tandem mass spectrometry. J Chromatogr A 1217: 7738–7748. https://doi.org/10.1016/j. chroma.2010.10.039 13. Hsu FF, Turk J (2000) Characterization of phosphatidylethanolamine as a lithiated adduct by triple quadrupole tandem mass spectrometry with electrospray ionization. J Mass Spectrom 35:595–606. https://doi.org/10.1002/ (SICI)1096-9888(200005)

14. Hsu FF, Turk J (2001) Studies on phosphatidylglycerol with triple quadrupole tandem mass spectrometry with electrospray ionization: fragmentation processes and structural characterization. J Am Soc Mass Spectrom 12: 1036–1043. https://doi.org/10.1016/ S1044-0305(01)00285-9 15. Zianni R, Bianco G, Lelario F et al (2013) Fatty acid neutral losses observed in tandem mass spectrometry with collision-induced dissociation allows regiochemical assignment of sulfoquinovosyl-diacylglycerols. J Mass Spectrom 48:205–215. https://doi.org/10.1002/ jms.3149 16. Guella G, Frassanito R, Mancini I (2003) A new solution for an old problem: the regiochemical distribution of the acyl chains in galactolipids can be established by electrospray ionization tandem mass spectrometry. Rapid Commun Mass Spectrom 17:1982–1994. https://doi.org/10.1002/rcm.1142 17. Abida H, Dolch L-J, Meı¨ C et al (2015) Membrane glycerolipid remodeling triggered by nitrogen and phosphorus starvation in Phaeodactylum tricornutum. Plant Physiol 167:118– 136. https://doi.org/10.1104/pp.114. 252395

Chapter 15 The Use of Nanopore Sequencing to Analyze the Chloroplast Transcriptome Part I: Library Preparation Se´bastien Skiada, Alexandra Launay-Avon, Arnaud Liehrmann, Etienne Delannoy, and Benoıˆt Castandet Abstract Global understanding of plastid gene expression has always been impaired by its complexity. RNA splicing, editing, and intercistronic processing create multiple transcripts isoforms that can hardly be resolved using traditional molecular biology techniques. During the last decade, the wide adoption of RNA-seq-based techniques has, however, allowed an unprecedented understanding of all the different steps of chloroplast gene expression, from transcription to translation. Current strategies are nonetheless unable to identify and quantify full length transcripts isoforms, a limitation that can now be overcome using Nanopore Sequencing. We here provide a complete protocol to produce, from total leaf RNA, cDNA libraries ready for Nanopore sequencing. While most Nanopore protocols take advantage of the mRNA polyA tail we here first ligate an RNA adapter to the 3′ ends of the RNAs and use it to initiate the template switching reverse transcription. The cDNA is then prepared and indexed for use with the regular Oxford Nanopore v14 chemistry. This protocol is of particular interest to researchers willing to simultaneously study the multiple post-transcriptional processes prevalent in the chloroplast. Key words Nanopore sequencing, RNA-Seq, Chloroplast, RNA processing, RNA maturation

1

Introduction Plastid genomes still retain a strong bacterial identity. DNA is condensed with proteins in membrane bound structures called nucleoids and most genes are organized into clusters or polycistrons [1, 2]. Going from genes to proteins is however a very complex process as the different steps of gene expression are the result of a sophisticated interplay between nucleus and chloroplast encoded factors [3]. Transcriptomics data suggest that the full chloroplast genome is transcribed [4, 5], which, coupled to inefficient termination [6, 7], leads to a highly complex primary transcriptome. It is then heavily modified by a variety of posttranscriptional processing

Eric Mare´chal (ed.), Plastids: Methods and Protocols, Methods in Molecular Biology, vol. 2776, https://doi.org/10.1007/978-1-0716-3726-5_15, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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events like RNA splicing, editing, and trimming to produce the accumulating RNAs suitable for translation [3, 8]. Additionally, apart from the well-known genic transcripts, it is now clear that chloroplasts also possess abundant non-coding RNAs (ncRNAs) whose functions are still debated [9, 10]. This complexity is well illustrated by the psbB gene cluster where 6 genes lead to the accumulation of more than 20 mRNA isoforms that can be resolved on RNA blots [11] and more than 30 mRNA termini [12]. A major issue to understand this complexity is that traditional molecular biology techniques routinely used to analyze RNA phenotypes are tedious, labor intensive, and time consuming. They are therefore only suited to gene-by-gene experiments and offer limited perspectives where gene function is concerned as pleiotropic effects are common in chloroplast mutants [8]. These limitations have been the motivation for the development of RNA-seq-based approaches, leading to an unprecedented understanding of the different steps of chloroplast gene expression. For example, dedicated bioinformatics pipelines now allow the monitoring of RNA abundance, splicing, and editing [13–15], ribosome profiling can be used to monitor chloroplast translation rate [16, 17], and Terminome sequencing gives a single-nucleotide resolution map of the full set of chloroplast RNA termini [12]. One aspect that is however still resistant to RNA-seq analyses is the complexity and quantification of transcripts isoforms, mainly for technological reasons. Indeed, Illumina sequencing only sequences short fragments that are then bioinformatically assembled into longer transcripts. This makes it difficult to interpret the combination of genuine RNA processing events, a pressing problem when dealing with gene dense genomes like the chloroplast. With the development of Nanopore technology, it is now possible to directly sequence full length individual RNAs following conversion to cDNA [18, 19], a strategy that bears the promise to finally allow the identification and quantification of chloroplast transcripts isoforms. Another advantage of the long reads is that it allows the study of the coordination between different RNA processing events targeting a single transcript and separated by several hundreds of nucleotides. We recently demonstrated this possibility in the chloroplast [20]. A limitation, however, is that most Nanopore library preparation protocols published so far are based on the presence of a poly A tail in 3′ of the RNAs [18, 21] while this feature is largely absent in chloroplast transcripts where adenylation is a degradation signal [22]. To overcome this difficulty, we therefore had to adapt a protocol already developed and validated for RACE experiments where a RNA adapter is ligated to the 3′ ends of the RNA and serves as a template to initiate the template switching reverse transcription [20]. We provide here a step by step protocol to generate,

The Use of Nanopore Sequencing to Analyze the Chloroplast Transcriptome. . .

245

from total leaf RNA, Nanopore libraries amenable to Oxford Nanopore sequencing. A flowchart illustrating the key steps of the protocol is depicted in Fig. 1. The bioinformatic analysis of the sequencing results is detailed in a second manuscript (see Chap. 16 in this same issue).

2

Materials

2.1 DNA and RNA Oligonucleotides

1. RNA oligo ONT_LIG (10 μM): 5′/5Phos/rNrNrNrNrUrGrArArUrGrCrArArCrArCrUrUrCrUrGrUrArC/3InvdT/-3′ (obtained from providers such as IDT Technologies, Leuven, Belgium). 2. DNA oligo ONT_RT II (24 μM): 5′- GCAGGGGAAATCATCAGCGTATAACGTACAGAA GTGTTGCATTC -3′ (obtained from providers such as IDT Technologies, Leuven, Belgium). 3. DNA oligo ONT-TSO (48 μM): 5′-AAGCAGTGGTATCAACGCAGAGTACrGrG+G-3′ (obtained from providers such as IDT Technologies, Leuven, Belgium). 4. DNA oligo ONT-primer V2 (12 μM): 5′-AAGCAGTGGTATCAACGCAGAGTAC-3′ 5. DNA oligo ONT-primer II V2 (12 μM): 5′-GCAGGGGAAATCATCAGCGTATAAC-3′

2.2 Molecular Biology Kits and Enzymes

1. T4 RNA ligase kit (obtained from providers such as NEB). 2. SMARTScribe Reverse Transcriptase (obtained from providers such as TaKaRa). 3. QIAseq FastSelect-rRNA Plant kit (obtained from providers such as Qiagen). 4. SeqAmp DNA Polymerase kit (obtained from providers such as TaKaRa). 5. 10 mM dNTP mix. 6. RNase inhibitor, 40 U/μL (obtained from providers such as NEB). 7. Agencourt AMPure XP beads (from Beckman Coulter). 8. Agencourt RNAClean XP reagent (from Beckman Coulter) 9. NEB Ultra II End repair/dA-Tailing Module kit (obtained from providers such as NEB). 10. Native Barcoding Kit V14 (from Oxford Nanopore). 11. Blunt/TA Ligase Master Mix (obtained from providers such as NEB).

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Fig. 1 Workflow of the Nanopore sequencing library preparation protocol. The main steps are highlighted and numbered according to the methods section

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12. Quick Ligation Module (such as NEBNext obtained from NEB). 13. Short Fragment Buffer (SFB) (from Oxford Nanopore). 14. DNA HS kit (from Agilent). 2.3 Buffers and Solutions

1. Elution Buffer (EB): 1 M Trisodium Citrate (Na3C6H5O7), pH 6.4, 10% (v/v) Tween20. 2. Binding Buffer (BB): 1 M Trisodium Citrate (Na3C6H5O7), pH 6.4, 10% (v/v) Tween20, 5 M NaCl, 50% PEG8000 (see Note 1). 3. Reaction buffer 10x (RB): 1% (v/v) Tween20, RNase inhibitor, 40 U/20 μL. 4. 70% and 80% (v/v) Ethanol. 5. Ultrapure H2O.

2.4

Equipment

1. Incubator (4 °C, 25 °C or 37 °C). 2. Microcentrifuge. 3. Magnetic rack. 4. Thermocycler. 5. Agilent Technology 2100 Bioanalyzer.

3

Methods

3.1 Ligating RNA and RACE Oligonucleotide 3.1.1

RNA Ligation

1. Use 200 ng RNA diluted in 5.5 μL ultrapure H2O. Total leaf RNA purified with the NucleoZOL reagent is compatible with this protocol. 2. Add 1 μL of 10 μM ONT_LIG oligo and mix well by pipetting up and down. 3. Spin down briefly (see Note 2). 4. Incubate at 65 °C for 5 min to remove secondary structures, then immediately chill on ice for at least 2 min. 5. On a separate microcentrifuge tube, prepare the following mix at room temperature: – 2 μL of T4 RNA ligase buffer 10x. – 2 μL of 10 mM ATP (see Note 3). – 0.5 μL of RNase inhibitor. – 8 μL of 50% (w/v) PEG8000. – 1 μL of T4 RNA Ligase (10 U/μL) (see Note 4). 6. Gently mix all reagents by pipetting up and down and spin briefly.

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7. Add the 13.5 μL of this reaction mixture to the tube containing the RNA and ONT_LIG oligonucleotide previously placed on ice (total reaction volume is 20 μL). 8. Mix well by pipetting up and down and spin down briefly (see Note 2). 9. Incubate at 25 °C for 2 hours or 4 °C overnight (see Note 5). 3.1.2 Purification of RNA Ligation

This step uses the AMPure RNAClean XP reagent to purify the ligated RNA and remove reagents. Before starting, AMPure RNAClean XP reagent should be placed at room temperature for 30 min. All the following steps are carried out at room temperature. 1. After thoroughly vortexing the reagent, add 36 μL AMPure RNAClean XP reagent to the reaction tube. 2. Mix well by pipetting up and down at least ten times. 3. Incubate for 8 min. 4. Place on a magnetic rack to collect the beads and wait until the liquid is clear (approx. 5 min). 5. Remove and discard the supernatant from the reaction tube. 6. Elute the RNA with 20 μL fresh buffer EB. 7. Incubate for 2 min. 8. Add 40 μL of fresh buffer BB, then mix by pipetting up and down. 9. Incubate for 5 min. 10. Place on a magnetic rack to collect the beads and wait until the liquid is clear (approx. 5 min). 11. Remove and discard the supernatant from the reaction tube. 12. Wash three times with 70% ethanol: Keep the tube on the magnetic rack and add 200 μL fresh 70% ethanol. Incubate for 30 s and remove and discard all supernatant. 13. Spin down the reaction tube and place it back onto the magnetic rack to allow the beads to be completely bound. Remove any residual 70% ethanol with a 20 μL micropipette. 14. Air dry for 2–3 min. 15. Elute the RNA with 10 μL ultrapure H2O, then mix by pipetting up and down. 16. Incubate for 2 min. 17. Put the tube on the magnetic rack for 2 min. 18. Transfer 9.5 μL of the supernatant containing ligated RNA into a new microcentrifuge tube.

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Depletion

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This step uses the QIAseq FastSelect-rRNA Plant Kit to remove the overabundant rRNA. 1. On ice, combine in PCR tube: – 9.5 μL of ligated RNAs. – 1.5 μL of reaction buffer RB 10x. – 1 μL of Qiagen FastSelect 0.5x. – 3 μL of First-strand Buffer 5x. 2. Place the PCR tube on a thermocycler and run the following program (see Note 6): – Step 1: 72 °C for 3 min. – Step 2: 70 °C for 1 min. – Step 3: 65 °C for 1 min. – Step 4: 60 °C for 1 min. – Step 5: 55 °C for 1 min. – Step 6: 50 °C for 1 min. – Step 7: 37 °C for 1 min. – Step 8: 25 °C for 1 min. – Step 9: Hold at 4 °C. 3. Add 25 μL of ultrapure H2O to the PCR tube (final volume is 40 μL).

3.2.2 Purification of Depleted RNA

This step uses AMPure RNAClean XP reagent to purify the depleted RNA and remove reagents. Before starting, AMPure RNAClean XP reagent should be placed at room temperature for 30 min. All the following steps are carried out at room temperature unless otherwise specified. 1. After thoroughly vortexing the reagent, add 48 μL of AMPure RNAClean XP reagent to the reaction tube. 2. Mix well by pipetting up and down at least ten times. 3. Incubate for 8 min. 4. Place on a magnetic rack to collect the beads and wait until the liquid is clear (approx. 5 min). 5. Remove and discard the supernatant from the reaction tube. 6. Wash twice with 70% ethanol: keep the tube on the magnetic rack and add 200 μL fresh 70% ethanol. Incubate for 30 s and remove and discard all supernatant. 7. Spin down the reaction tube and place it back onto the magnetic rack to allow the beads to be completely bound. Remove any residual 70% ethanol with a 20 μL micropipette. 8. Air dry for 3 min or until beads look completely dry.

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9. Elute the RNA with 10 μL ultrapure H2O, then mix by pipetting up and down. 10. Incubate for 10 min at 37 °C (see Note 6). 11. Place on a magnetic rack for 1 min. 12. Transfer 9.5 μL of the supernatant containing the rRNA depleted ligated RNA into a new microcentrifuge tube. 3.3

cDNA Synthesis

1. On a PCR tube at room temperature, prepare the following mix using the SMARTScribe Reverse Transcriptase kit: – 2 μL of first-strand buffer 5x. – 0.25 μL of 0.1 M DTT. – 1 μL of 10 mM dNTP. – 0.5 μL of 24 μM ONT_RT II primer. – 1 μL of 48 μM ONT-TSO primer. – 0.25 μL of RNase inhibitor (40 U/μL). – 1 μL of SMARTScribe Reverse Transcriptase (100 U/μL) (see Note 4). 2. Gently mix all reagents by pipetting up and down and spin briefly. 3. Add 4 μL of the rRNA depleted ligated RNA to this reaction mix (total reaction volume is 10 μL). 4. Mix by pipetting up and down and spin down briefly (see Note 2). 5. Place the PCR tube on a thermocycler and run the following program (see Note 5): – Step 1: 42 °C for 90 min to allow reverse transcription. – Step 2: 70 °C for 10 min to stop the reaction. – Step 3: Hold at 10 °C.

3.4 Amplification of the cDNA by LD-PCR 3.4.1

PCR Amplification

1. On a microcentrifuge tube at room temperature, prepare the following mix using the SeqAmp DNA Polymerase kit: – 25 μL of SeqAmp PCR buffer 2x. – 1 μL of 12 μM ONT-primer V2 primer. – 1 μL of 12 μM ONT-primer II V2 primer. – 1 μL of SeqAmp DNA Polymerase (1.25 U/μL). – 12 μL ultrapure H2O. 2. Gently mix all reagents by pipetting up and down and spin down briefly. 3. Add the 40 μL of this reaction mix to the PCR tube containing the cDNA (10 μL) from the previous step (total reaction volume is 50 μL).

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4. Mix by pipetting up and down and spin down briefly (see Note 2). 5. Place the PCR tube on a thermocycler and run the following program (see Note 6): – Step 1: Denaturation at 95 °C for 1 min. – Step 2: Denaturation at 98 °C for 10 s. – Step 3: Annealing at 65 °C for 30 s. – Step 4: Elongation at 68 °C 3 min. – Repeat Steps 2–3–4, 19 times. – Step 5: Final elongation at 72 °C for 10 min. – Step 6: Hold at 10 °C. 3.4.2

Library Purification

This step uses AMPure XP beads to purify the depleted cDNA and remove reagents. Before starting, AMPure XP beads should be placed at room temperature for 30 min. All the following steps are carried out at room temperature unless otherwise specified. 1. Add 50 μL ultrapure H2O to the PCR tube. 2. After thoroughly vortexing the beads, add 60 μL AMPure XP beads to each reaction tube. 3. Mix well by pipetting up and down at least ten times. 4. Incubate for 8 min. 5. Place on a magnetic rack to collect the beads and wait until the liquid is clear (approx. 5 min). 6. Remove and discard the supernatant. 7. Wash twice with 80% ethanol: Keep the tube on the magnetic rack and add 200 μL fresh 80% ethanol. Incubate for 30 s and remove and discard all supernatant. 8. Spin down the reaction tube and place it back onto the magnetic rack to allow the beads to be completely bound. Remove any residual 80% ethanol with a 20 μL micropipette. 9. Air dry for 3 min or until beads look completely dry. 10. Elute the library with 20 μL ultrapure H2O, then mix by pipetting up and down. 11. Incubate for 10 min at 37 °C (see Note 6). 12. Place on a magnetic rack for 1 min. 13. Transfer 19 μL of the supernatant containing the amplified cDNA into a new microcentrifuge tube.

3.5 cDNA Quantification and Quality Control

This step is necessary to check the quality of the cDNA amplification and the quantification is required to ensure the different samples are pooled in equimolar amounts at Subheading 3.8.

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Fig. 2 A typical cDNA profile obtained on an Agilent Bioanalyzer with a DNA high sensitivity kit

1. Dilute the amplified cDNA at 1:5 in ultrapure H2O. 2. Run 1 μL of the dilution on an Agilent Technology 2100 Bioanalyzer using a High Sensitivity DNA chip. A typical library shows a broad size distribution (Fig. 2). 3. Determine the cDNA concentration in fmol/μL from the Bioanalyzer trace. 4. You can store the full-length cDNAs at 4 °C for up to 72h or at -80 °C for several months. 3.6 cDNA End Repair and dA-Tailing 3.6.1 End Repair and dATailing

This step uses the NEB Ultra II End repair/dA-Tailing Module kit to produce cDNA with 5′ phosphorylated and 3′ dA-tailed ends (referred to as end-prepped cDNA). 1. Transfer 100 fmol of the amplified cDNA into a new PCR tube (the volume depends on the concentration estimated in the previous step). 2. Complete to 11.5 μL with ultrapure H2O. 3. Add 1.75 μL of NEB Ultra II End-prep Reaction Buffer. 4. Add 0.75 μL NEB Ultra II End-prep Enzyme Mix. 5. Gently mix all reagents by pipetting up and down at least ten times. 6. Spin down briefly (total reaction volume is 14 μL) (see Note 2). 7. Place the PCR tube on a thermocycler and run the following program (see Note 6): – Step 1: 20°C for 5 min. – Step 2: 65°C for 5 min.

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This step uses AMPure XP beads to purify the end-prepped cDNAs and remove reagents. Before starting, AMPure XP beads should be placed at room temperature for 30 min. All the following steps are carried out at room temperature. 1. After thoroughly vortexing the beads, add 15 μL AMPure XP beads to the reaction tube. 2. Mix well by pipetting up and down at least ten times. 3. Incubate for 5 min. 4. Place on a magnetic rack to collect the beads and wait until the liquid is clear (approx. 5 min). 5. Remove and discard the supernatant. 6. Wash twice with 80% ethanol: Keep the tube on the magnetic rack and add 200 μL fresh 80% ethanol. Incubate for 30 s and remove and discard all supernatant. 7. Spin down the reaction tube and place it back onto the magnetic rack to allow the beads to be completely bound. Remove any residual 80% ethanol with a 20 μL micropipette. 8. Air dry for 3 min or until beads look completely dry. 9. Elute the cDNA with 11.5 μL ultrapure H2O, then mix by pipetting up and down. 10. Incubate for 2 min. 11. Place on a magnetic rack for 1 min. 12. Transfer 10 μL of the supernatant containing the end-prepped cDNA into a new microcentrifuge tube (see Note 7).

3.7

cDNA Indexing

3.7.1

Index Ligation

This step uses the Native Barcoding Kit V14 to ligate indexing adapters to the end-prepped cDNA. 1. Transfer 7.5 μL of the end-prepped cDNA from the previous step into a new microcentrifuge tube. 2. Add 2.5 μL of the Barcode Adapter of your choice from the Native Barcoding Kit V14. 3. Add 10 μL of Blunt/TA ligase Master Mix (total reaction volume is 20 μL). 4. Mix well by pipetting up and down at least ten times. 5. Spin down briefly (see Note 2). 6. Incubate for 20 min at room temperature. 7. Add 2 μL of the EDTA provided in the Native Barcoding Kit V14.

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3.7.2 Indexed cDNAs Purification

This step uses AMPure XP beads to purify the indexed cDNA and remove reagents. Before starting, AMPure XP beads should be placed at room temperature for 30 min. All the following steps are carried out at room temperature unless otherwise specified. 1. Add 8 μL of AMPure XP beads to the reaction tube. 2. Mix well by pipetting up and down at least ten times. 3. Incubate for 5 min. 4. Place on a magnetic rack to collect the beads and wait until the liquid is clear (approx. 5 min). 5. Remove and discard the supernatant. 6. Wash twice with 80% ethanol: keep the tube on the magnetic rack and add 200 μL fresh 80% ethanol. Incubate for 30 s and remove and discard all supernatant. 7. Spin down the reaction tube and place it back onto the magnetic rack to allow the beads to be completely bound. Remove any residual 80% ethanol with a 20 μL micropipette. 8. Air dry for 3 min or until beads look completely dry. 9. Elute the library with 21 μL ultrapure H2O, then mix by pipetting up and down. 10. Incubate for 10 min at 37 °C (see Note 6). 11. Place on a magnetic rack for 1 min. 12. Transfer 10 μL of the supernatant containing the library into a new microcentrifuge tube (see Note 7).

3.8 Sequencing Adapter Ligation of the Indexed cDNA 3.8.1

Adapter Ligation

This step uses the NEBNext Quick Ligation Module to ligate the sequencing adapters to the indexed cDNA. These adapters are necessary for the library to be recognized by the Nanopore sequencer. 1. Pool equal volumes of the different indexed cDNAs of interest into a single microcentrifuge tube. The final volume must be 30 μL. 2. Add 5 μL of Native Adapter. 3. Add 10 μL of the NEBNext Quick Ligation Reaction Buffer 5x. 4. Mix by pipetting up and down. 5. Add 5 μL of the Quick T4 DNA ligase (total reaction volume is 50 μL). 6. Mix by pipetting up and down. 7. Spin down at 500× g maximum (see Note 2). 8. Incubate for 20 min at room temperature.

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Final Purification

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This step uses AMPure XP beads to purify the final library and remove reagents as well as adapter dimers. Before starting, AMPure XP beads and the SFB buffer should be placed at room temperature for 30 min. All the following steps are carried out at room temperature unless otherwise specified. 1. Add 20 μL AMPure XP beads to each reaction tube. 2. Mix well by pipetting up and down at least ten times. 3. Incubate for 10 min. 4. Place on a magnetic rack to collect the beads and wait until the liquid is clear (approx. 5 min). 5. Remove and discard the supernatant. 6. Add 125 μL of the buffer SFB. 7. Incubate 30 s. 8. Place on a magnetic rack to collect the beads and wait until the liquid is clear (approx. 5 min). 9. Remove and discard the supernatant. 10. Spin down the reaction tube and place it back onto the magnetic rack to allow the beads to be completely bound. Remove any residual SFB with a 20 μL micropipette. 11. Elute the library with 15 μL fresh buffer EB, then mix by pipetting up and down. 12. Incubate for 10 min at 37 °C (see Note 5). 13. Place on a magnetic rack for 1 min. 14. Transfer 15 μL of the supernatant containing the library into a new microcentrifuge tube. 15. Analyze and quantify the library on an Agilent DNA Highsensitivity chip. An example of a good library profile is provided in Fig. 3. The library is ready to be sequenced. You can store it at 4 °C for 24h or at -80 °C for 10 days.

4

Notes 1. This buffer can be stored at room temperature for 3 months maximum. 2. Only spin down at low speed to prevent damaging the library during the process (shearing, fragmentation. . .). 3. Prepare aliquots to avoid multiple freeze-thaw cycles. 4. To get the best efficiency, add the enzyme just before use. 5. Both work equally well in our hands.

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Fig. 3 A typical Nanopore library profile on an Agilent Bioanalyzer with a DNA high sensitivity kit

6. We advise to pre-heat the incubator or the thermocycler to be sure that the tubes are immediately put at the required temperature. 7. Samples can be stored at 4 °C overnight. It is a potential stopping point.

Acknowledgments This work was supported by the Agence Nationale de la Recherche through the grant ANR-20-CE20-0004 JOAQUIN to BC. AL was supported by a PhD fellowship from the French ministe`re de l’enseignement supe´rieur et de la recherche. The IPS2 benefits from the support of Saclay Plant Sciences-SPS (ANR-17-EUR0007). References 1. Majeran W, Friso G, Asakura Y et al (2012) Nucleoid-enriched proteomes in developing plastids and chloroplasts from maize leaves: a new conceptual framework for nucleoid functions. Plant Physiol 158(1):156–189 2. Sakai A, Takano H, Kuroiwa T (2004) Organelle nuclei in higher plants: structure, composition, function, and evolution. Int Rev Cytol 238(SPEC. ISS.):59–118 3. Small I, Melonek J, Bohne AV et al (2023) Plant organellar RNA maturation. Plant Cell 35(6):1727–1751 4. Zhelyazkova P, Sharma CM, Forstner KU et al (2012) The primary transcriptome of barley chloroplasts: numerous noncoding rnas and

the dominating role of the plastid-encoded RNA polymerase. Plant Cell 24(1):123–136 5. Lima MS, Smith DR (2017) Pervasive, genome-wide transcription in the organelle genomes of diverse plastid-bearing protists. Genes Gen Genet 7(11):3789–3796 6. Mullet JE, Klein RR (1987) Transcription and RNA stability are important determinants of higher plant chloroplast RNA levels. EMBO J 6(6):1571–1579 7. Stern DB, Gruissem W (1987) Control of plastid gene expression: 3′ inverted repeats act as mRNA processing and stabilizing elements, but do not terminate transcription. Cell 51(6):1145–1157

The Use of Nanopore Sequencing to Analyze the Chloroplast Transcriptome. . . 8. Germain A, Hotto AM, Barkan A et al (2013) RNA processing and decay in plastids. Wiley Interdiscip Rev RNA 4(3):295–316 9. Hotto AM, Schmitz RJ, Fei Z, Ecker JR et al (2011) Unexpected diversity of chloroplast noncoding RNAs as revealed by deep sequencing of the Arabidopsis transcriptome. Genes Gen Genet 1(7):559–570 10. Hotto AM, Germain A, Stern DB (2012) Plastid non-coding RNAs: emerging candidates for gene regulation. Trends Plant Sci 17(12): 737–744 11. Stoppel R, Meurer J (2013) Complex RNA metabolism in the chloroplast: an update on the psbB operon. Planta 237(2):441–449 12. Castandet B, Germain A, Hotto AM et al (2019) Systematic sequencing of chloroplast transcript termini from Arabidopsis thaliana reveals >200 transcription initiation sites and the extensive imprints of RNA-binding proteins and secondary structures. Nucleic Acids Res 47(22):11889–11905 13. Castandet B, Hotto AM, Strickler SR et al (2016) ChloroSeq, an optimized chloroplast RNA-Seq bioinformatic pipeline, reveals remodeling of the organellar transcriptome under heat stress. Gen Gen Genet 6(9):2817–2827 14. Michel EJSS, Hotto AM, Strickler SR et al (2018) A guide to the chloroplast transcriptome analysis using RNA-Seq. Methods Mol Biol 1829:295–313

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15. Malbert B, Rigaill G, Brunaud V et al (2018) Bioinformatic analysis of chloroplast gene expression and RNA posttranscriptional maturations using RNA sequencing. Methods Mol Biol 1829:279–294 16. Chotewutmontri P, Stiffler N, Watkins KP et al (2018) Ribosome profiling in maize. Methods Mol Biol 1676:165–183 17. Chotewutmontri P, Barkan A (2016) Dynamics of chloroplast translation during chloroplast differentiation in maize. PLOS Genet 12(7): e1006106 18. Parker MT, Knop K, Sherwood AV et al (2020) Nanopore direct RNA sequencing maps the complexity of arabidopsis mRNA processing and m6A modification. Elife 9:e49658 19. Gru¨nberger F, Ferreira-Cerca S, Grohmann D (2022) Nanoore sequencing of RNA and cDNA molecules in Escherichia coli. RNA 28(3):400–417 20. Guilcher M, Liehrmann A, Seyman C et al (2021) Full length transcriptome highlights the coordination of plastid transcript processing. Int J Mol Sci 22(20):11297 21. Cozzuto L, Liu H, Pryszcz LP et al (2020) MasterOfPores: a workflow for the analysis of oxford nanopore direct RNA sequencing datasets. Front Genet 11:211 22. MacIntosh GC, Castandet B (2020) Organellar and secretory ribonucleases: major players in plant RNA homeostasis. Plant Physiol 183(4): 1438

Chapter 16 The Use of Nanopore Sequencing to Analyze the Chloroplast Transcriptome Part II: Bioinformatic Analyzes and Virtual RNA Blots Etienne Delannoy, Arnaud Liehrmann, and Benoıˆt Castandet Abstract Nanopore sequencing of full-length cDNAs offers unprecedented details of the plastid RNA metabolism. After the generation of the nanopore reads, several bioinformatic steps are required to analyze the data. In this chapter, we describe in a few simple command lines the processing and mapping of the reads as well as the generation of virtual Northern blots as a simple and familiar way to visualize Nanopore sequencing data. Key words Nanopore sequencing, RNA-Seq, Chloroplast, Virtual Northern blot

1

Introduction Over the last decade, RNA-seq has been a major tool helping to better understand the various steps of plastid gene expression. It has now become the method of choice for deep analysis of chloroplast transcript patterns, not only because of its constant decreasing cost but also because of its exceptional flexibility and adaptability [1]. Various library preparation protocols and analytical bioinformatic pipelines have indeed been developed to study virtually all aspects of plastid gene expression, going well beyond differential gene expression analysis. Historically, RNA-seq has been used first to discover a hundred novel plastid noncoding RNAs (pncRNA) in Arabidopsis thaliana [2] and to show that their expression was modified upon heat stress in Brassica [3]. In another application, size selected RNAs were identified as footprints of RNA binding proteins and shed light on the sequence specificity of RNA processing [4, 5]. Strategies designed to specifically capture transcripts termini revealed a

Eric Mare´chal (ed.), Plastids: Methods and Protocols, Methods in Molecular Biology, vol. 2776, https://doi.org/10.1007/978-1-0716-3726-5_16, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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surprisingly large number of transcription initiation sites and more than a thousand mature 5′ and 3′ ends [6, 7]. RNA editing and splicing were also extensively studied through RNA-seq [8–12], and finally, sequencing of mRNA fragments protected by the ribosome gave new insights into plastid translation [13, 14]. Almost all these strategies are based on Illumina sequencing that only produces short sequence fragments or reads, a major concern when the biological question of interest is to identify or quantify the different transcripts isoforms expressed from a gene [15]. These fragments are indeed bioinformatically assigned to a specific transcript isoform using probabilistic models [16–18], but the accuracy of these approaches decreases as the number of transcripts isoforms increases [19]. This is particularly problematic when studying plastid gene expression as one transcriptional unit can produce dozens of different transcripts isoforms [20]. The advent of long read sequencing technology like PacBio or Oxford Nanopore is a unique opportunity to overcome these difficulties[1]. It is theoretically now possible to identify and quantify all of the plastid transcripts isoforms as the reads may indeed cover an entire isoform, from start to end. One issue however is that most Nanopore library preparation protocols use the mRNA polyA tail to initiate a template switching reverse transcription. Because polyadenylation of chloroplastic RNAs acts as a degradation signal, we recently developed an alternative protocol using the ligation of an RNA adapter at the 3′ end of the RNA to initiate the cDNA synthesis [21] and Skiada et al. (Chap. 15, this volume). Alternative approaches based on A tailing of the RNAs have also recently been used in bacteria where short poly A tails similarly act as a degradation signal [22]. Because Nanopore sequencing to study plastid or bacterial gene expression is still in its infancy, there are no standardized or user friendly pipeline to analyze the data. Moreover, visualization of long reads data is a challenge of its own and new tools are being developed to tackle this issue [21, 23]. We describe here the bioinformatic steps to analyze the reads generated using the protocol proposed in Skiada et al. (Chap. 15, this volume). After the conversion of the raw Nanopore data into fastq files, the proper orientation of the reads is identified and restored before they are mapped on the reference genome. Once mapped, the reads can be analyzed using classical RNA-seq tools as described in [24], but full-length cDNA reads offer many other possibilities among which the generation of virtual Northern blots. We describe how to generate these Northern blots with a few command lines.

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Materials 1. Although no coding knowledge is necessary to apply the methods explained here, an understanding of basic Unix commands is helpful. 2. Commands given in the Linux shell are denoted with a ‘$’ prefix. 3. The analysis can be run on a 64-bit Linux system, and at least 8 GB of memory is recommended. The following programs should be installed (a) Last [25] that can be downloaded from https://gitlab. com/mcfrith/last (b) Cutadapt [26] that can be downloaded from https:// cutadapt.readthedocs.io/en/stable/ (c) Samtools [27] that can downloaded from http://www. htslib.org/ (d) Minimap2 [28] that can be downloaded from https:// github.com/lh3/minimap2 (e) R with the GenomicRanges, GenomicAlignments ([29] both available on Bioconductor, https://bioconductor. org/), ggplot2 (https://ggplot2.tidyverse.org/) and reshape2 ([30] https://cran.r-project.org/web/ packages/reshape2/index.html) packages. (f) Python with the matplotlib (https://matplotlib.org/) and numpy (https://numpy.org/) packages and the math and csv modules. 4. The files required are: (a) The fasta file of the genome on which to map the reads. It will hereafter be named “genome_seq.fasta”. The file used in this example can be downloaded from https://doi.org/ 10.57745/UWAHNR (b) The fastq files generated for each sample by the Nanopore basecalling and demultiplexing. They are usually grouped per barcode in a folder called pass/barcodexx. The example files C_final.fastq.gz and P_final.fastq.gz corresponding to stranded and trimmed fastq reads can be downloaded from https://doi.org/10.57745/ UWAHNR (c) The set_strand.sh, virtualNorthern.R and vBNv2.py scripts can be downloaded from https://doi.org/10. 57745/UWAHNR

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Methods Setting Strand

The Nanopore basecalling and demultiplexing can be performed either live or after the sequencing. It generates multiple fastq files containing only unstranded reads. The set_strand.sh bash script identifies the correct orientation of each reads using the presence of the 5′ end and 3′ end adapters ligated to the reads during the library construction (Chap. 15, see Note 1). When needed, it reverse-complements the reads and finally removes the sequences corresponding to the end adapters. 1. Copy the set_strand.sh script in the “pass” folder containing the “barcode” folders 2. Run the command: $ find barcode* -maxdepth 0 -type d -exec bash -c ’cd "{}"; pwd;../set_strand.sh’ \; This command will run the set_strand.sh script in each barcode folder. The stranded reads are in the final_stranded_reads.fastq.gz files in each barcode folder.

3.2 Mapping of the Stranded Reads

The mapping of the stranded reads is done using minimap2 that requires an index file of the genome sequence. 1. Generate the genome index file genome_seq.mmi by running the command: $ minimap2 -d genome_seq.fasta genome_seq.mmi 2. Run the command: $ minimap2 -a -x splice -t 20 --secondary=no genome_seq.mmi C_final.fastq.gz | samtools sort - -o C.bam This is an example for the sample C. The results are written in the C.bam file that is specified by the –o option. The -x splice option indicates that we are mapping RNA-seq reads on a genome sequence. Twenty processing cores (-t 20) are used for the process, but this can be adapted according to the hardware available. Only primary mappings are provided (-secondary=no). The mapping is then sorted with samtools (| samtools sort - ). The same command must be adapted and run for the other data points (see Note 2). 3. Run the command: $ samtools index C.bam This command generates the index file of the C.bam file. It is required for several downstream analyses.

3.3 Visualization Using Virtual Northern Blots

Once the mapping is done, many different analyses can be performed depending on the questions. However, full length RNA-seq presents unique opportunities and challenges in terms of visualization. A classical approach is to use the genome browser

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IGV [31], but this viewer displays the diversity of reads rather than the full set of reads mapping at a particular locus. This means that it provides a biased picture of the amount of the different isoforms. Here, using the virtualNorthern.R R script and the vBNv2.py python script, we propose a simple way to generate virtual Northern blots that are a more faithful representation of the diversity of the transcripts isoforms. For more complex normalizations and visualizations, we however advise you to use the Nanoblot R package that uses bam files to generate customizable virtual Northern blots ([23], available at https://github.com/SamDeMario-lab/ NanoBlot). 1. In the folder containing the indexed bam files, copy the virtualNorthern.R R script and the vBNv2.py python script. 2. Run the command $ Rscript --vanilla virtualNorthern.R Pt 75700 76000 plus 2000 In this example, all bam files in the folder will be processed to extract the size of the reads overlapping the region from the start position 75700 to the end position 76000 on the plus strand of the Pt chromosome (exon2 of petB in Arabidopsis thaliana). The strand parameter can be either “plus” or “minus.” The maximum size of reads to display in the virtual Northern blot is 2000 bases (see Note 3 for another example). This command generates two files: – Northern_intensities.pdf (Fig. 1) corresponding to the plot of the abundance of each read size for all bam files. The intensities are normalized within each bam file to a value of 100 for the most abundant read size. – size_distrib.txt corresponding to the input required for the vBNv2.py python script 3. Run the command $ python vBNv2.py This command will generate the file vNB.png (Fig. 2) from the size_distrib.txt input file corresponding to the same locus as Fig. 1.

4

Notes 1. IMPORTANT: The reads must come from sequencing libraries constructed exactly as described in part 1 (Chap. 15). The bioinformatic processing of the fastq reads relies on the sequence of the DNA oligos ONT_RT II and ONT-TSO. Any change in their sequence should be translated in the set_strand. sh script.

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100

intensity

75

samples C

50

P

25

0 0

500

1000

length

1500

2000

Fig. 1 Density plot of the RNA abundance at the petB exon2 locus of A. thaliana in the samples C and P. The intensity of the most abundant isoform in each sample is set to 100

2. For example, for the P sample, the command is $ minimap2 -a -x splice -t 20 --secondary=no genome_seq.mmi P_final.fastq. gz | samtools sort - -o P.bam 3. The following command: $ Rscript --vanilla virtualNorthern.R Mt 97700 98100 minus 2000 will extract the size of the reads overlapping the region from the start position 97700 to the end position 98100 on the minus strand of the Mt chromosome (exons 1, 2, and 3 of mitochondrial nad2 in Arabidopsis thaliana).

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Fig. 2 Virtual Northern blot emulated from the same data as Fig. 1

Acknowledgments This work was supported by the Agence Nationale de la Recherche through the grant ANR-20-CE20-0004 JOAQUIN to BC. The IPS2 benefits from the support of Saclay Plant Sciences-SPS (ANR-17-EUR-0007).

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References 1. Stark R, Grzelak M, Hadfield J (2019) RNA sequencing: the teenage years. Nat Rev Genet 20(11):631–656 2. Hotto AM, Schmitz RJ, Fei Z et al (2011) Unexpected diversity of chloroplast noncoding RNAs as revealed by deep sequencing of the Arabidopsis transcriptome. Genes Gen Genet 1(7):559–570 3. Wang L, Yu X, Wang H, Lu YZ et al (2011) A novel class of heat-responsive small RNAs derived from the chloroplast genome of Chinese cabbage (Brassica rapa). BMC Genomics 12(1):289 4. Ruwe H, Schmitz-Linneweber C (2012) Short non-coding RNA fragments accumulating in chloroplasts: footprints of RNA binding proteins? Nucleic Acids Res 40(7):3106–3116 5. Ruwe H, Wang G, Gusewski S et al (2016) Systematic analysis of plant mitochondrial and chloroplast small RNAs suggests organellespecific mRNA stabilization mechanisms. Nucleic Acids Res 44(15):7406–7417 6. Zhelyazkova P, Sharma CM, Forstner KU et al (2012) The primary transcriptome of barley chloroplasts: numerous noncoding RNAs and the dominating role of the plastid-encoded RNA polymerase. Plant Cell 24(1):123–136 7. Castandet B, Germain A, Hotto AM et al (2019) Systematic sequencing of chloroplast transcript termini from Arabidopsis thaliana reveals >200 transcription initiation sites and the extensive imprints of RNA-binding proteins and secondary structures. Nucleic Acids Res 47(22):11889–11905 8. Ruwe H, Castandet B, Schmitz-Linneweber C et al (2013) Arabidopsis chloroplast quantitative editotype. FEBS Lett 587(9):1429–1433 9. Castandet B, Hotto AM, Strickler SR et al (2016) ChloroSeq, an optimized chloroplast RNA-Seq bioinformatic pipeline, reveals remodeling of the organellar transcriptome under heat stress. Genes Gen Genet 6(9):2817–2827 10. Malbert B, Rigaill G, Brunaud V et al (2018) Bioinformatic analysis of chloroplast gene expression and rna posttranscriptional maturations using RNA sequencing. Methods Mol Biol 1829:279–294 11. Guillaumot D, Lopez-Obando M, Baudry K et al (2017) Two interacting PPR proteins are major Arabidopsis editing factors in plastid and mitochondria. Proc Natl Acad Sci U S A 114(33):8877–8882 12. Michel EJSS, Hotto AM, Strickler SR et al (2018) A guide to the chloroplast

transcriptome analysis using RNA-Seq. Methods Mol Biol 1829:295–313 13. Chotewutmontri P, Barkan A (2016) Dynamics of chloroplast translation during chloroplast differentiation in maize. PLOS Genet 12(7): e1006106 14. Chotewutmontri P, Stiffler N, Watkins KP et al (2018) Ribosome profiling in maize. Methods Mol Biol 1676:165–183 15. Wang Y, Zhao Y, Bollas A et al (2021) Nanopore sequencing technology, bioinformatics and applications. Nat Biotechnol 39(11): 1348–1365 16. Li B, Dewey CN (2011) RSEM: accurate transcript quantification from RNA-Seq data with or without a reference genome. BMC Bioinformatics 12(1):323 17. Trapnell C, Roberts A, Goff L et al (2012) Differential gene and transcript expression analysis of RNA-seq experiments with TopHat and Cufflinks. Nat Protoc 7(3):562–578 18. Patro R, Duggal G, Love MI et al (2017) Salmon provides fast and bias-aware quantification of transcript expression. Nat Methods 14(4):417–419 19. Zhang C, Zhang B, Lin L et al (2017) Evaluation and comparison of computational tools for RNA-seq isoform quantification. BMC Genomics 18(1):583 20. Stoppel R, Meurer J (2013) Complex RNA metabolism in the chloroplast: an update on the psbB operon. Planta 237(2):441–449 21. Guilcher M, Liehrmann A, Seyman C et al (2021) Full length transcriptome highlights the coordination of plastid transcript processing. Int J Mol Sci 22(20):11297 22. Gru¨nbergern F, Ferreira-Cerca S, Grohmann D (2022) Nanopore sequencing of RNA and cDNA molecules in Escherichia coli. RNA 28(3):400–417 23. DeMario S, Xu K, He K et al (2023) NanoBlot: an R-package for visualization of RNA isoforms from long read RNA-sequencing data. RNA. rna.079505.122 24. Baudry K, Delannoy E, Colas des Francs-Small C (2022) Analysis of the Plant Mitochondrial Transcriptome. Methods Mol. Biol 2363:235– 262 25. Kiełbasa SM, Wan R, Sato K et al (2011) Adaptive seeds tame genomic sequence comparison. Genome Res 21(3):487–493

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Chapter 17 Targeted Gene Editing of Nuclear-Encoded Plastid Proteins in Phaeodactylum tricornutum via CRISPR/Cas9 Ce´cile Giustini, Jhoanell Angulo, Florence Courtois, and Guillaume Allorent Abstract Genome modifications in microalgae have emerged as a crucial and indispensable tool for research in fundamental and applied biology. In particular, CRISPR/Cas9 has gained significant recognition as a highly effective method for genome engineering in these photosynthetic organisms, enabling the targeted induction of mutations in specific regions of the genome. Here, we present a comprehensive protocol for generating knock-out mutants in the model diatom Phaeodactylum tricornutum using CRISPR/Cas9 by both biolistic transformation and bacterial conjugation. Our protocol outlines the step-by-step procedures and experimental conditions required to achieve successful genome editing, including the design and construction of guide RNAs, the delivery of CRISPR/Cas9 components into the algae cells, and the selection of the generated knockout mutants. Through the implementation of this protocol, researchers can harness the potential of CRISPR/Cas9 in P. tricornutum to advance the understanding of diatom biology and explore their potential applications in various fields. Key words Phaeodactylum tricornutum, CRISPR/Cas9, Genome editing, Biolistic transformation, Conjugation

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Introduction Phaeodactylum tricornutum, a marine unicellular microalga, holds significant ecological importance and has garnered attention as a promising model organism and valuable research tool in both academic and industrial sectors [1, 2]. The complete sequencing of its genome [3] facilitates targeted genetic modifications and opens up new avenues for scientific research. Historically, the creation of gene-specific knock-out mutants in Phaeodactylum tricornutum first relied on labor-intensive and expensive protocols, such as meganucleases or TAL effector endonucleases (TALENs) [4, 5]. While effective, these techniques presented challenges for routine application in laboratories, limiting their practicality and

Eric Mare´chal (ed.), Plastids: Methods and Protocols, Methods in Molecular Biology, vol. 2776, https://doi.org/10.1007/978-1-0716-3726-5_17, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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widespread adoption. The emergence of CRISPR/Cas9 genome editing tools suitable in Phaeodactylum tricornutum few years ago [6] has revolutionized genetic manipulation in this diatom species by dramatically improving the ability to generate stable mutants of interest with increased efficiency, while significantly reducing costs and simplifying experimental procedures [7]. The CRISPR/Cas9 system, originally derived from bacterial immune mechanisms [8], has been successfully adapted across various organisms [9]. It comprises two main components: the Cas9 DNA endonuclease, responsible for generating DNA double-strand breaks, and a guide RNA (sgRNA) that is bound to the Cas9 protein and acts as a recognition module to ensure the delivery of the Cas9 endonuclease to the desired genomic location. The recognition of the DNA sequence by sgRNA is facilitated by a specific sequence referred to as “target sequence.” By modifying this region, researchers can precisely determine the gene and the location within the gene where DNA cleavage and subsequent genetic modifications will occur with high accuracy. After Cas9 cuts the DNA, the DNA repair machinery comes into play to fix these breaks, often resulting in the insertion or deletion of nucleotides. This repair process alters the protein sequence upon translation and may result in the formation of a premature stop codon, thereby modifying gene function. This ability for precise genetic modification has been a revolution in the field of genome editing. In recent years, significant advancements have been made in modifying the Cas9 nuclease, enabling its application beyond traditional genome editing. Researchers have harnessed the power of CRISPR/Cas9 to manipulate gene transcription, enabling precise control over the level of gene expression. In addition, innovative techniqus [10] have emerged that leverage the Cas9 protein to visualize specific DNA sequences in cells, providing valuable insights into genome dynamics and organization. Additionally, the Cas9 protein can be used to target and purify specific DNA regions, enabling the efficient isolation and analysis of genomic fragments. Here, we describe a protocol for creating stable targeted gene knock-out (KO) mutants by CRISPR/Cas9. This protocol allows the generation of mutants of interest within two months using biolistic transformation or bacterial conjugation. Biolistic transformation is a widely recognized and efficient method for transforming living cells, including unicellular microalgae, with DNA molecules. More recently, bacterial conjugation has been shown to be a highly effective protocol for the transformation of P. tricornutum [11, 12]. In this approach, the transfer of exogenous DNA takes place through physical interaction between Escherichia coli and diatoms, facilitating the transfer of genetic material. This approach offers several benefits: (i) the inserted DNA (episome) can be readily extracted from the algae, which can be a key

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advantage for biotechnological applications; (ii) a slight reduction in the time required to obtain transformants; and (iii) significantly reduced experiment costs, as no gene gun apparatus or related consumables are necessary. This protocol describes the procedure to select the CRISPR/Cas9 target sequence in the gene of interest, prepare the plasmids for transformation, transform the cells, and analyze transformants.

2

Materials

2.1 Identification of Targets Sequences for CRISPR/Cas9 Editing in the Gene of Interest

1. Phire Plant Direct PCR Master Mix (from ThermoFisher Scientific) or equivalent. This kit contains the dilution buffer used for DNA extraction.

2.1.1 Solutions and Consumables 2.1.2

Equipment

1. Vortex mixer. 2. Table top centrifuge. 3. Thermal cycler.

2.2 Design and Synthesis of the Adapter

1. T4 DNA ligase buffer (from providers such as New England Biolabs).

2.2.1 Solutions and Consumables 2.2.2

Equipment

1. Thermal cycler.

2.3 P. tricornutum Transformation by Gene Gun

1. pKSdiaCas9_sgRNA plasmid (from Addgene).

2.3.1 Digestion of the pKSdiaCas9_sgRNA Plasmid by BsaI

3. CutSmart Buffer from (New England Biolabs).

2. BsaI-HFv2 endonuclease, 20,000 U/mL (from providers such as New England Biolabs). 4. Agarose and buffer for DNA electrophoresis. 5. Agarose gel DNA extraction kit.

Solutions and Consumables Equipment

1. Water or dry bath. 2. DNA gel electrophoresis system. 3. Table top centrifuge. 4. Nanodrop/spectrophotometer for DNA quantitation.

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2.3.2 Ligation of the Adapter into the Linearized pKSdiaCas9_sgRNA Plasmid Solutions and Consumables

1. T4 DNA ligase (from providers such as New England Biolabs). 2. Bacteria DH5α or Top10 competent cells. 3. Lysogeny Broth (LB) liquid medium. 4. LB agar plates supplemented with ampicillin (100 μg/mL, from providers such as ThermoFisher Scientific). 5. Ampicillin (from providers such as ThermoFisher Scientific). 6. Plasmid mini-prep extraction kit.

Equipment

1. Water or dry bath. 2. Sterile hood. 3. 37
C-heated incubator and shaker for bacteria plates and liquid cultures. 4. Table top centrifuge.

2.3.3 P. tricornutum Cells Liquid Culture and Plating Before Biolistic Transformation

1. ESAW liquid medium. 2. ESAW plates with 1% (w/v) agar.

Solutions and Consumables Equipment

1. Sterile hood. 2. Table top centrifuge. 3. Incubator and shaker for P. tricornutum cells culture.

2.3.4 Gold Beads Preparation Solutions and Consumables Equipment

1. 0.6 μm gold microcarriers (ref 1652262). 2. Absolute ethanol 70% (v/v). 3. Glycerol 50% (v/v).

1. Table top centrifuge. 2. Vortex mixer.

2.3.5 Coating Gold Beads with Plasmids Solutions and Consumables

1. pAF6 plasmid [11] or any plasmid conferring to P. tricornutum cells antibiotic resistance for selection of the transformants. 2. 2.5 M CaCl2 sterile solution. 3. 0.1 M Spermidine (from providers such as Sigma Aldrich) prepared in sterile water, aliquoted in 1.5 mL microfuge tubes and stored at 80
C. 4. Absolute ethanol and 70% (v/v) ethanol in 1.5 mL centrifuge tubes and stored at 20
C.

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1. Table top centrifuge. 2. Vortex mixer.

2.3.6 Cells Bombardment Solutions and Consumables

1. Consumables for the PDS-1000/He system (from Biorad) or equivalent: 1550 psi rupture disks, stopping screens, macrocarriers, macrocarrier holders. 2. Enriched artificial seawater (ESAW) liquid medium: use appropriate composition as described is articles such as [13] or from a provider. A large number of compositions are available and should be tested for the capacity to grow diatom cells in the designed laboratory conditions. 3. ESAW plates with 1% (w/v) agar supplemented with Zeocin (from providers such as ThermoFisher Scientific).

Equipment

1. PDS-1000/He system (from Biorad) or equivalent. 2. Sterile hood. 3. Incubator for P. tricornutum cells culture.

2.4 P. tricornutum Transformation by Bacterial Conjugation

1. PtPuc3_diaCas9_sgRNA plasmid (from Addgene).

2.4.1 Digestion of the PtPuc3_diaCas9_sgRNA Plasmid by BsaI

3. CutSmart Buffer (from New England Biolabs).

2. BsaI-HFv2 endonuclease, 20,000 U/mL (from providers such as New England Biolabs). 4. Agarose and buffer for DNA electrophoresis. 5. Agarose gel DNA extraction kit.

Solutions and Consumables Equipment

1. Water or dry bath. 2. DNA gel electrophoresis system. 3. Table top centrifuge. 4. Nanodrop/spectrophotometer for DNA quantitation, or equivalent.

2.4.2 Ligation of the Adapter into the Linearized PtPuc3_diaCas9_sgRNA Plasmid Solutions and Consumables

1. T4 DNA ligase (from providers such as New England Biolabs). 2. E. coli competent cells (preferentially DH10B, see Note 9). 3. Lysogeny Broth (LB) liquid medium. 4. LB agar plates supplemented with kanamycin (50 μg/mL, from providers such as ThermoFisher Scientific). 5. Plasmid mini-prep extraction kit.

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Equipment

1. Water or dry bath. 2. Sterile hood. 3. 37
C-heated incubator and shaker for bacteria plates and liquid cultures. 4. Table top centrifuge.

2.4.3 Bacteria Preparation for Conjugation Solutions and Consumables

1. pTAmob plasmid (from Addgene). 2. E. coli competent cells (preferentially DH10B, see Note 9). 3. LB agar plates supplemented with kanamycin (50 μg/mL, from providers such as ThermoFisher Scientific) and gentamycin (10 μg/mL, from providers such as Roth). 4. Lysogeny Broth (LB) liquid medium.

Equipment

1. Water or dry bath. 2. Sterile hood. 3. 37
C-heated incubator and shaker for bacteria plates and liquid cultures. 4. Table top centrifuge. 5. Spectrophotometer.

2.4.4 P. tricornutum Cells Liquid Culture and Plating Before Conjugation Solutions and Consumables Equipment

1. ESAW 1/2 plates with 1% (w/v) agar supplemented with 5% LB (v/v). 2. ESAW liquid medium. 3. Six-well plate (from providers such as Greiner Bio-One).

1. Sterile hood. 2. Incubator for P. tricornutum cells culture. 3. Table top centrifuge.

2.4.5 Transfer of the PtPuc3_diaCas9_sgRNA Plasmid into P. tricornutum Cells by Bacterial Conjugation

1. ESAW liquid medium. 2. ESAW 1/2 plates with 1% (w/v) agar supplemented with 5% LB (v/v), 100 μg/mL Zeocin, and 100 μg/mL ampicillin (from providers such as ThermoFisher Scientific).

Solutions and Consumables Equipment

1. Incubator for P. tricornutum cells culture. 2. 30
C-heated incubator. 3. Sterile hood.

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2.5 Selection of Mutants

1. ESAW plates with 1% (w/v) agar supplemented with Zeocin (100 μg/mL).

2.5.1 Solutions and Consumables

2. Phire Plant Direct PCR Master Mix (from ThermoFisher Scientific) or equivalent. This kit contains the Dilution Buffer used for DNA extraction.

2.5.2

1. Sterile hood.

Equipment

2. Table top centrifuge. 3. Vortex mixer. 4. Thermal cycler. Please ensure to diligently adhere to all waste disposal regulations when disposing of waste materials, particularly when handling materials and solutions that contain gold beads. It is essential to always wear gloves when manipulating tubes and working with reagents.

3

Methods Unless otherwise instructed, perform all procedures at room temperature.

3.1 Identification of Target Sequences for CRISPR/Cas9 Editing in the Gene of Interest

1. Pour 20 μL of dilution buffer into a 1.5 mL microfuge tube.

3.1.1 Extraction of Genomic DNA (gDNA) and PCR Amplification of the Gene of Interest

3. Thoroughly resuspend the cells in the dilution buffer until no aggregates are visible, resulting in a pale brown solution.

2. Transfer a small amount of P. tricornutum cells, equivalent to the volume of two pinheads, from a plated colony into a microfuge tube.

4. Vortex vigorously for 30 s. 5. Centrifuge at 16,000 g for 5 min at 22
C. 6. Transfer the supernatant containing gDNA into a new microfuge tube. 7. Design two primers for PCR amplification of the gene of interest, utilizing the DNA sequence retrieved from the database (http://protists.ensembl.org/Phaeodactylum_tri cornutum/Info/Index) (see Note 1). 8. Amplify the gene of interest by PCR using 0.5 μL of gDNA solution as template and primers designed in step 7 (see Note 2). 9. Proceed with amplicon sequencing by utilizing the services of a DNA sequencing company, using the primers from step 7. Employ the resulting sequence to identify the CRISPR/Cas9 target sequence in the subsequent phase (Subheading 3.1.2).

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Fig. 1 Example of design and synthesis of the adapter. Primers for the creation of the adapter are designed by removing the NGG terminal part and by adding TCGA or AAAC overhangs in the CRISPR/Cas9 target sequence. Synthesis of the adapter is realized by annealing the two designed primers 3.1.2 Identification of CRISPR/Cas9 Target Sequences

1. Identify CRISPR/Cas9 target sequences in the gene of interest using the PhytoCRISP-Ex (https://www.phytocrispex. biologie.ens.fr/CRISP-Ex/) or CRISPOR (http://crispor. tefor.net/) web tools: use the sequence obtained in Subheading 3.1.1 and follow the procedure described in [14] (see Notes 3 and 4).

3.2 Design and Synthesis of the Adapter

1. Follow the instructions in Fig. 1 to design the two primers necessary for creating the adapter that will be inserted into the plasmid in Subheading 3.3 (gene gun transformation) or in Subheading 3.4 for the bacterial conjugation protocol. In summary, the adapter involves annealing two 20-mer complementary oligonucleotides with 50 overhangs.

3.2.1 Bioinformatic Design of the Adapter 3.2.2 Synthesis of the Adapter

1. The adapter is generated by annealing the two primers designed in Subheading 3.2.1—mix 1 μg of each primer with 1x of T4 ligase buffer in a total volume of 50 μL in PCR tubes. 2. Incubate the mixture in a thermal cycler at 85
C for 10 min, followed by gradually decreasing the temperature by 5
every 5 min until reaching 25
C. Keep on ice or store at 20
C until Subheading 3.3.2 for gene gun transformation or 3.4.2 for transformation by bacterial conjugation.

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Fig. 2 Simplified map of the pKSdiaCas9_sgRNA and the PtPuc3_diaCas9_sgRNA plasmids. A detailed and complete overview of the features of these two plasmids can be found on the Addgene website. (a) The pKSdiaCas9_sgRNA plasmid, used for gene gun transformation, enables the expression of a P. tricornutum codon-optimized Cas9 protein and the sgRNA containing the CRISPR/Cas9 target sequence between the two BsaI restriction sites. In bacteria, this plasmid provides ampicillin resistance (AmpR). (b) The pKSdiaCas9_sgRNA plasmid, used for transformation via bacteria conjugation, facilitates the expression of a P. tricornutum codon-optimized Cas9 protein and the sgRNA containing the CRISPR/Cas9 target sequence between the two BsaI restriction sites. This plasmid confers Zeocin resistance to P. tricornutum cells (BleoR) and kanamycin resistance (KanR) to bacteria. For both plasmids, the M13 reverse primer (M13 rev) is used for sequencing the target sequence 3.3 P. tricornutum Transformation by Gene Gun 3.3.1 Digestion of the pKSdiaCas9_sgRNA Plasmid by BsaI (Fig. 2, See Note 4)

The subsequent steps outline the procedure for the gene gun transformation protocol. For transformation using the bacterial conjugation approach, refer to Subheading 3.4. 1. Mix 1 μg of the pKSdiaCas9_sgRNA plasmid, 1 μL of BsaI-HF restriction enzyme, and 1x of the CutSmart buffer in a total volume of 25 μL. 2. Incubate at 37
C for 2 h to liberate the 20 bp fragment corresponding to the previous target sequence from the plasmid. 3. To ensure the separation of the previous target sequence from the linearized plasmid and to prevent religation, load and run the digestion mix on a 1% (w/v) agarose gel according to usual laboratory procedure. 4. Excise the agarose band corresponding to the linearized plasmid (expected size: 8500 bp). Note that the band representing the 20 bp fragment of the previous target sequence may not be visible on the gel due to its extremely small size.

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5. Extract DNA from the agarose gel using a commercial kit following the supplier’s recommendations. 6. Quantify the linearized plasmid using Nanodrop or spectrophotometer approaches. Keep on ice until Subheading 3.3.2. 3.3.2 Ligation of the Adapter into the Linearized pKSdiaCas9_sgRNA Plasmid

1. Mix 5 μL of the adapter obtained in Subheading 3.2.2, 100 ng of the linearized pKSdiaCas9-sgRNA plasmid obtained in Subheading 3.3.1, 2 μL of T4 DNA ligase, and 1x of T4 DNA ligase buffer in a total volume of 20 μL. 2. Incubate at room temperature for 10 min or overnight at 16
C to generate the pKSdiaCas9_sgRNA plasmid containing the CRISPR/Cas9 target sequence of the gene of interest. 3. Use 4 μL of the ligation mix to transform E. coli competent cells using standard protocols. 4. Proceed to plating on LB agar plates supplemented with 100 μg/mL ampicillin and incubate overnight at 37
C. 5. Pick up a colony and amplify in LB medium supplemented with ampicillin (100 μg/mL) under agitation overnight at 37
C. 6. Purify plasmid using mini-prep extraction kit following the supplier’s recommendations. 2.5 μg of plasmid is required for each bombardment (Ssubheading 3.4). 7. Submit the plasmid for sequencing to a DNA sequencing company to verify the correct insertion of your target sequence. Use the universal M13-rev primer (GTCATAGCTGTTTCCTG) for the sequencing process (Fig. 2a).

3.3.3 P. tricornutum Cells Liquid Culture and Plating Before Biolistic Transformation

1. Grow P. tricornutum cells (12 h light/dark cycles, 30/40 μmol photons.m2.s1, 19
C) in ESAW liquid medium so that the culture reaches exponential phase (between 1.5106 and 2.5106 cells/mL) 24 h before transformation (Subheading 3.4 below). 2. Under a sterile hood, harvest 108 cells from the liquid culture. 3. Centrifuge cells (2250 g, 10 min, 19
C) and gently discard the supernatant. 4. Resuspend the pellet containing 108 cells in 500 μL of fresh ESAW medium. 5. Immediately spread 500 μL of cell suspension on ESAW plate containing 1% (w/v) agar without antibiotics (see Note 5). 6. Allow the plate to dry under the sterile hood until the liquid is evaporated (around 30 min). 7. Incubate the plate under dim light (20/30 μmol of photons. m2.s1, 19
C) for 24 h.

CRISPR/Cas9 in P. tricornutum 3.3.4 Gold Beads Preparation (Adapted from the BIORAD PDS-1000/He System Protocol)

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This step can be anticipated and realized at any time. 1. Weigh 30 mg of gold beads into a 1.5 mL sterile microfuge tube. 2. Add 1 mL of 70% (v/v) ethanol. 3. Vortex vigorously for 5 min. 4. Let the beads settle for 15 min. 5. Pellet the beads by short centrifugation (5 s, 6000 g, 22
C). 6. Remove and discard the supernatant. 7. Wash the beads: add 1 mL of water, vortex vigorously 1 min, let the beads settle for 1 min, pellet beads by short centrifugation (10 s, 3000 rpm, 22
C), remove and discard the supernatant. 8. Repeat step 7 two times. 9. Resuspend beads in 500 μL sterile 50% (v/v) glycerol by thoroughly vortexing. 10. Prepare aliquots of 50 μL in 1.5 mL microfuge tubes and store at 20
C for up to 6 months.

3.3.5 Coating Gold Beads with Plasmids (Adapted from the BIORAD PDS1000/He System Protocol)

1. In a 1.5 mL microfuge tube, combine 2.5 μg of the pKSdiaCas9_sgRNA plasmid prepared in Subheading 3.3.2 with 2.5 μg of the pAF6 plasmid conferring Zeocin resistance (see Note 4). Ensure that the total volume of the mixture does not exceed 10 μL. 2. Consider one 50 μL aliquot of gold beads prepared in Subheading 3.3.4. 3. While vortexing, add the following to the beads in order: the mix of both plasmids, 50 μL of 2.5 M CaCl2, and 20 μL of 0.1 M spermidine (see Note 6). 4. Vortex for 2–3 min and let settle for 1 min at room temperature. 5. Pellet beads by short centrifugation (10 s, 6000 g, 22
C), remove and discard the supernatant. 6. Add 140 μL of ice-cold 70% (v/v) ethanol. Do not vortex or mix and directly remove and discard ethanol. 7. Repeat the previous step with ice-cold 100% ethanol. 8. Add 48 μL of 100% ethanol to the sample. Keep the sample on ice and proceed promptly with cell bombardment.

3.3.6 Cells Bombardment

1. Use the PDS-1000/He system under a sterile hood according to the manufacturer’s instructions. 2. Switch on the laminar flow hood and carefully clean it and the PDS-1000/He system with ethanol 70% (v/v).

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3. Wash macrocarrier holders, macrocarrier, and stopping screens in a 70% ethanol bath. Allow them to dry on a filter paper under the hood. Do not wash the rupture disks. 4. Carefully resuspend the beads coated with plasmids prepared in Subheading 3.3.5 and load 10 μL onto the macrocarrier. Allow it to dry for 2 min under the hood until the ethanol evaporates. 5. Insert the 1550-psi rupture disk and assemble the microcarrier launch (including the stopping screen, the macrocarrier, and macrocarrier holder prepared earlier) following the manufacturer’s instructions. 6. Position the previously plated P. tricornutum cells (refer to Subheading 3.3.3) on shelf n
2, ensuring a 6 cm distance between the microcarrier launch and the plate. 7. Proceed to the bombardment as described in the manufacturer’s instructions. 8. Repeat steps 5–7 three times on the same plate, using a new macrocarrier prepared as described in step 4 above for each repetition. Rotate the plate by 45
between each bombardment to ensure that a larger surface area is targeted. 9. Allow the cells to recover on plate in an incubator under dim light (30/40 μmol of photons.m2.s1, 19
C) for 24–48 h. 10. After the recovery period, add 1 mL of fresh sterile ESAW liquid medium to the plate and gently resuspend the transformed cells using an inoculation loop. 11. Transfer and spread the cells onto a new ESAW plate containing 100 μg/mL of Zeocin for selection (see Note 7). 12. Allow the plate to dry under the hood until the liquid has evaporated, which typically takes around 30 min. 13. Place the plate in an incubator under dim light (30/40 μmol of photons.m2.s1, 19
C). 14. The first transformants should appear as colonies with a brownish color within a timeframe of 3 weeks. 15. Go to Subheading 3.5 for the screening procedure of the transformants. 3.4 P. tricornutum Transformation by Bacterial Conjugation

1. Mix 1 μg of the PtPuc3_diaCas9_sgRNA plasmid, 1 μL of BsaI-HF restriction enzyme, and 1x of the CutSmart buffer in a total volume of 25 μL (see Note 4).

3.4.1 Digestion of the PtPuc3_diaCas9_sgRNA Plasmid by BsaI (Fig. 2)

2. Incubate at 37
C for 2 h to liberate the 20 bp fragment corresponding to the previous target sequence from the plasmid. 3. To ensure the separation of the previous target sequence from the linearized plasmid and to prevent religation, load and run the digestion mix on a 1% (w/v) agarose gel according to usual laboratory procedure.

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4. Excise the agarose band corresponding to the linearized plasmid (expected size ~12800 bp). Note that the band representing the 20 bp fragment of the previous target sequence may not be visible on the gel due to its extremely small size. 5. Extract DNA from the agarose gel using a commercial kit following the supplier’s recommendations. 6. Quantify the linearized plasmid using Nanodrop or spectrophotometer approaches. Place it on ice and proceed to Subheading 3.4.2, or store it at 20
C for long-term preservation. 3.4.2 Ligation of the Adapter into the Linearized PtPuc3_diaCas9_sgRNA Plasmid

1. Mix 5 μL of the adapter obtained in Subheading 3.2.2, 100 ng of the linearized PtPuc3_diaCas9_sgRNA plasmid obtained in Subheading 3.3.1, 2 μL of T4 DNA ligase, and 1x of T4 DNA ligase buffer in a total volume of 20 μL. 2. Incubate at room temperature for 10 min or overnight at 16
C to generate the PtPuc3_diaCas9_sgRNA plasmid containing the CRISPR/Cas9 target sequence of the gene of interest. 3. Use 4 μl of the ligation mix to transform E. coli competent cells using standard protocols (see Note 8). 4. Proceed to plating on LB agar plates supplemented with 50 μg/mL kanamycin and incubate overnight at 37
C. 5. Pick up a colony and amplify in LB medium supplemented with kanamycin (50 μg/mL) under agitation overnight at 37
C. 6. Purify plasmid using mini-prep extraction kit following the supplier’s recommendations. 7. Submit the plasmid for sequencing to a DNA sequencing company to verify the correct insertion of your target sequence. Use the universal M13-rev primer (GTCATAGCTGTTTCCTG) for the sequencing process (Fig. 2b). The following Subheadings, 3.4.3 and 3.4.4, need to be carried out in parallel and require prior planning.

3.4.3 Bacteria Preparation for Conjugation

1. Mix 100 ng of the PtPuc3_diaCas9_sgRNA plasmid prepared in Subheading 3.4.2 with 100 ng of the pTAmob plasmid. 2. Use the mix to transform E. coli competent cells using standard protocols (see Note 9). 3. Proceed to plating on LB agar plates supplemented with 50 μg/mL kanamycin and 10 μg/mL gentamycin. Incubate overnight at 37
C. 4. Store the plate at 4
C. The plates can be stored in these conditions for at least one month. 5. At the end of the day before the conjugation, inoculate a 5 mL culture of LB supplemented with 50 μg/mL kanamycin and

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10 μg/mL gentamicin with the bacteria containing the two plasmids from step 4. Incubate the culture overnight at 37
C with agitation at 200 rpm. 6. On the morning of the day of conjugation, use 600 μL–1.2 mL of this culture to inoculate a 30 mL culture of LB supplemented with 50 μg/mL kanamycin and 10 μg/mL gentamicin. Keep at 37
C with agitation at 200 rpm. 7. Regularly monitor the optical density of the culture at 600 nm. When it reaches a range of 0.8–1.2, centrifuge the bacterial culture for 5 min at 3500 g at 22
C. 8. Discard the supernatant and carefully resuspend the pellet in 200 μL of LB. 3.4.4 P. tricornutum Cells Liquid Culture and Plating Before Conjugation

1. Prepare a 50 mL solution of ESAW supplemented with 1% agar (w/v) and 5% LB (v/v). Autoclave the solution. Do not add antibiotics at this stage. Add 8 mL of the solution to one well of a six-well plate. One well is needed for each conjugation event. It is necessary to reserve some wells for conducting the positive and negative control of the experiment (see Note 10). This step can be performed a few days before the experiment. In this case, seal the plate with a parafilm and keep the plate in the dark at 4
C. 2. Grow P. tricornutum cells (12 h light/dark cycles, 30/40 μmol photons.m2.s1, 19
C) in ESAW liquid medium so that the culture reaches exponential phase (between 1.5106 and 2.5106 cells/mL). 3. Under a sterile hood, harvest 2108 cells from the liquid culture. 4. Centrifuge cells (2250 g, 10 min, 19
C) and gently discard the supernatant. 5. Resuspend the pellet containing 2108 cells in 2 mL of fresh ESAW medium. 6. Spread 300 μL of the cell culture in each well of the plate prepared in step 1. Allow it to dry for 30 min under the hood until the liquid evaporates. 7. Seal the plate with parafilm and incubate for 4 days under 30/40 μmol photons.m2.s1 at 19
C.

3.4.5 Transfer of the PtPuc3_diaCas9_sgRNA Plasmid into P. tricornutum Cells by Bacterial Conjugation

1. Drop 100 μL of the bacteria prepared in Subheading 3.4.3 in one well of the six-well plate with algae cells prepared in Subheading 3.4.4. 2. Gently mix using an inoculating loop and allow to dry for 5 min under the hood.

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3. Seal the plate with parafilm and incubate for 90 min in the dark at 30
C. 4. Incubate the plate under continuous light (30/40 μmol photons.m2.s1) at 19
C for 48 h. 5. Add 1 mL of ESAW in the well and gently resuspend cells using inoculating loop. 6. Spread 500 μL of the cells onto two ESAW plates supplemented with 1% agar (w/v) and 100 μg/mL of Zeocin for selection. 7. Allow the plates to dry under the hood until the liquid has evaporated, which typically takes around 30 min. 8. Place the plate in an incubator under dim light (30/40 μmol of photons.m2.s1, 19
C). 9. The first transformants should appear as colonies with a brownish color within a timeframe of 2 weeks. 3.5 Selection of Mutants

1. Pick up transformants and spread them on a new plate of ESAW medium supplemented with Zeocin (100 μg/mL) to increase their size. 2. Extract gDNA from cells as described in Subheading 3.1.1. 3. Amplify by PCR the CRISPR/Cas9 targeted region of the gene of interest using appropriate primers. 4. Proceed with amplicon sequencing by utilizing the services of a DNA sequencing company. 5. Analyze sequencing chromatograms to identify and select mutants of interest (refer to Fig. 3 and see Note 11). All transformants possess Zeocin resistance. However, mutations are expected to occur near the CRISPR/Cas9 target sequence only if Cas9 has cleaved at the desired location. The efficiency of mutation occurrence varies significantly among genes and even between different target sequences within the same gene. Therefore, it is highly recommended to screen several dozen transformants. Different molecular phenotypes at the targeted locus can be observed. – Wild-type (WT)-like transformants, indicating no modification of the targeted locus, are generated in three different scenarios: first, when only the Zeocin resistance has been integrated; second, if the mutated gene induces a lethal phenotype; and third, when the DNA repair system is highly efficient. – Insertions and/or deletions can occur in the targeted sequence, with the type and length of these events varying among transformants, ranging from single modified

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Fig. 3 Molecular analysis of mutants. Sequencing chromatograms were generated by sequencing the PCR product of the region where the CRISPR/Cas9 target sequence was designed. Mutations compared to the wild type (WT) are indicated by red squares in the nucleotide sequence. The presence of mixed peaks, as observed in the last chromatogram, suggests a population containing a mixture of WT and/or mutant cells

nucleotides to dozens of nucleotides. It should be noted that in rare cases, the insertion of hundreds of nucleotides can also occur, originating from the plasmids used for transformation or from other parts of the P. tricornutum genome. – A mixture of modified targeted sequences is observed, as indicated by the overlapping peaks in the sequencing chromatogram (Fig. 3). This phenomenon occurs when different strains (mutants and/or wild-type cells) are present within the colony picked up during step 1. In such cases, streak the cells onto a new plate to isolate individual colonies once again and repeat steps 1–5. It may be necessary to repeat this procedure multiple times before achieving pure isolation of the desired mutant. 6. Check for the presence of a premature stop codon in mutants that have undergone insertion or deletion events. Specifically select and analyze the phenotype of 2 or 3 mutants (see Note 3) where the stop codon is positioned as close as possible to the ATG start codon to minimize the chance of generating a truncated yet potentially functional mutated protein. If possible, the absence of the protein can be verified through immunodetection using the appropriate antibody.

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Notes 1. Prior to identifying target sequences for CRISPR/Cas9, it is important to check for the occurrence of polymorphism in the gene of interest. This refers to differences between the DNA sequence present in the database (http://protists.ensembl. org/Phaeodactylum_tricornutum/Info/Index) and the sequence in the strain being used. Polymorphisms can potentially cause errors in determining the target sequence, resulting in the failure to cleave DNA at the intended locus. 2. The CRISPR/Cas9 target sequences should be positioned as closely as possible to the ATG start codon, preferably within the first exon. This approach reduces the likelihood of generating a truncated yet potentially functional protein in the mutant. 3. To minimize off-target effects, it is advisable to generate mutants by targeting, whenever feasible, at least two or three distinct regions within the gene of interest. Subsequently, phenotypic comparisons should be made among these mutants. Alternatively, mutants should be complemented by introducing a wild-type copy of the mutated gene to validate the observed phenotypes. When performing complementation, it is crucial to introduce silent mutations in the target sequence of the inserted gene copy. These mutations should not alter the protein sequence but serve to prevent further Cas9 cleavage. 4. The plasmid utilized for gene gun transformation in this protocol is pKSdiaCas9_sgRNA (Fig. 2). This plasmid enables the expression of a codon-optimized Cas9 nuclease and the sgRNA [6]. The CRISPR/Cas9 target sequence identified in subheading 3.1.2 must be inserted between the two BsaI sites. Additionally, it should be noted that this plasmid does not contain a resistance gene for selecting transformants on plates supplemented with the corresponding antibiotics. Therefore, a second plasmid carrying the resistance (pAF6, [15]) is required for transformation. Please note that during this procedure, there is no regeneration of the two BsaI sites, and the plasmid cannot be reused for another target. To work with a different target, you will need to start again from the original pKSdiaCas9_sgRNA plasmid. The same holds true for the PtPuc3_diaCas9_sgRNA plasmid used for conjugation except that the antibiotic resistance is already present in the plasmid. 5. To mitigate bacterial contamination that may occur during transformation, ampicillin (100 μg/mL) can be added to the plates.

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6. Spermidine should be aliquoted and stored at 80
C. Repeated freeze-thaw cycles of spermidine will significantly reduce transformation efficiencies. Thaw an aliquot on ice shortly before use. 7. It is possible to combine this step with Subheading 3.5 by picking up a colony, spreading it on a new plate, and utilizing the remaining cells on the inoculating loop for DNA extraction and PCR. In this scenario, reduce the volume of dilution buffer used in the gDNA extraction protocol described in Subheading 3.1.1. 8. The PtPuc3_diaCas9_sgRNA plasmid is a large-sized plasmid. Ensure to use the suitable bacteria for the transformation process like the DH10B competent cells from Thermofisher Scientific (ref EC0113). 9. At this stage, it is important for the bacteria to express both plasmids, which can potentially impact the transformation efficiency. To address this, increase the recovery time of the bacteria after transformation before plating them on the LB agar plate supplemented with kanamycin and gentamycin. 10. As conjugation requires both PtPuc3_diaCas9_sgRNA and pTAmob plasmids, proceed with conjugation using bacteria harboring only one of these two plasmids for negative control. Alternatively, use LB medium instead of bacteria culture at the Subheading 3.4.5. It would be necessary to adapt Subheading 3.4.3 to prepare bacteria used for control. 11. Commercial kits can be used in Subheading 3.5 to facilitate the detection of locus-specific double strand break formation. The presence of Cas9 in cells can also be checked by PCR. These approaches limit the number of samples sent for sequencing. References 1. Chisti Y (2007) Biodiesel from microalgae. Biotechnol Adv 25(3):294–306 2. Ambrust EV (2009) The life of diatoms in the world’s ocean. Nature 459(7244):185–192 3. Bowler C, Allen AE, Badger JH et al (2008) The Phaeodactylum genome reveals the evolutionary history of diatom genomes. Nature 456(7219):239–244 4. Daboussi F, Leduc S, Mare´chal A et al (2014) Genome engineering empowers the diatom Phaeodactylum tricornutum for biotechnology. Nat Commun. 5:3831 5. Weyman PD, Beeri K, Lefebvre SC et al (2015) Inactivation of Phaeodactylum tricornutum urease gene using transcription activator-like effector nuclease-based targeted mutagenesis. Plant Biotechnol J 13(4):460–470

6. Nymark M, Sharma AK, Sparstad T et al (2016) A CRISPR/Cas9 system adapted for gene editing in marine algae. Sci Rep 6:24951 7. Kroth PG, Bones AM, Daboussi F et al (2018) Genome editing in diatoms: achievements and goals. Plant Cell Rep 37(10):1401–1408 8. Jinek M, Chylinski K, Fonfara I et al (2012) A programmable dual-RNA-guided DNA endonuclease in adaptive bacterial immunity. Science 337(6096):816–821 9. Sander JD, Joung JK (2014) CRISPR-Cas systems for editing, regulating and targeting genomes. Nat Biotechnol 32(4):347–355 10. Wang H, La Russa M, Qi LS (2016) CRISPR/ Cas9 in genome editing and beyond. Annu Rev Biochem. 85:227–264

CRISPR/Cas9 in P. tricornutum 11. Karas BJ, Diner RE, Lefebvre SC et al (2015) Designer diatom episomes delivered by bacterial conjugation. Nat Commun 6:6925 12. Diner RE, Bielisnki VA, Dupont CL et al (2016) Refinement of the diatom episome maintenance sequence and improvement of conjugation-based DNA delivery methods. Front Bioeng Biotechnol 4:65 13. Villanova V, Fortunato AE, Singh D et al (2017) Investigating mixotrophic metabolism in the model diatom Phaeodactylum

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tricornutum. Philos Trans R Soc Lond B Biol Sci 372(1728) 14. Rastogi A, Murik O, Bowler C et al (2016) PhytoCRISP-Ex : a web-based and standalone application to find specific target sequences for CRISPR/CAS editing. BMC Bioinformatics 17(1):261 15. Falciatore A, Casotti R, Leblanc C et al (1999) Transformation of nonselectable reporter genes in marine diatoms. Mar Biotechnol (NY) 1(3):239–251

Chapter 18 Isolation of Cytosolic Ribosomes Associated with Plant Mitochondria and Chloroplasts Laura Dimnet, Thalia Salinas-Giege´, Sara Pullara, Lucas Moyet, Chloe´ Genevey, Marcel Kuntz, Anne-Marie Ducheˆne, and Norbert Rolland Abstract Excluding the few dozen proteins encoded by the chloroplast and mitochondrial genomes, the majority of plant cell proteins are synthesized by cytosolic ribosomes. Most of these nuclear-encoded proteins are then targeted to specific cell compartments thanks to localization signals present in their amino acid sequence. These signals can be specific amino acid sequences known as transit peptides, or post-translational modifications, ability to interact with specific proteins or other more complex regulatory processes. Furthermore, in eukaryotic cells, protein synthesis can be regulated so that certain proteins are synthesized close to their destination site, thus enabling local protein synthesis in specific compartments of the cell. Previous studies have revealed that such locally translating cytosolic ribosomes are present in the vicinity of mitochondria and emerging views suggest that localized translation near chloroplasts could also occur. However, in higher plants, very little information is available on molecular mechanisms controlling these processes and there is a need to characterize cytosolic ribosomes associated with organelles membranes. To this goal, this protocol describes the purification of higher plant chloroplast and mitochondria and the organelle-associated cytosolic ribosomes. Key words Cytosolic ribosomes, Organelle, Chloroplast, Mitochondria

1

Introduction Chloroplasts and mitochondria are two organelles involved in energy (ATP) production, through photosynthesis and cellular respiration, respectively. Chloroplast imports CO2, phosphate, and H2O for photosynthetic reactions. In turn, it produces O2, reducing power (NADPH) and ATP, which are used to fuel the cellular metabolism. Mitochondrion converts pyruvate into acetylCoA, which is used in TCA cycle. The generated reducing power (NADH, FADH2) allows ATP synthesis through oxidative phosphorylation. Both organelles are also involved in the synthesis of other essential organic compounds (vitamins, lipids, sugars, amino acids, pigments, hormones. . .) [1, 2].

Eric Mare´chal (ed.), Plastids: Methods and Protocols, Methods in Molecular Biology, vol. 2776, https://doi.org/10.1007/978-1-0716-3726-5_18, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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Chloroplasts and mitochondria both originate from an endosymbiotic event of a bacterium, respectively, a cyanobacterium or an α-proteobacterium, into a eucaryote. During evolution, numerous genes from the ancestral bacterium were lost or transferred to the nucleus. Nowadays, chloroplasts and mitochondria still have a genome, which codes for a limited number of proteins. A large majority of the ~2–3000 proteins in each organelles are nucleusencoded and therefore synthesized by cytosolic ribosomes before their post-translational import into the organelles, most of them via canonical import machineries relying on chloroplast transit peptide (cTP) or mitochondrial targeting sequence (MTS) at the N-terminal end of the imported polypeptides [3–5]. However, alternative mechanisms also exist for both chloroplasts and mitochondria (e.g., [6–10]). During the last decade, proteomic analyses targeting the chloroplast envelope from higher plants [11, 12] repeatedly identified subunits of cytosolic ribosomes associated with purified chloroplast envelope membranes, suggesting that localized translation might occur at the chloroplast surface. In the green algae Chlamydomonas reinhardtii, evidence was also obtained that localized translation occurs near plastids [13], and a plastid translation zone was proposed where both nuclear- and plastid-encoded mRNAs involved in photosynthetic complex biogenesis are translated [14]. Similarly, it has been shown that cytosolic ribosomes near the mitochondrial outer membrane synthesize hundreds of mitochondria proteins. Factors involved in such a localized translation were identified in yeast, Drosophila, and human cells [15, 16], and the presence of cytosolic ribosomes on the surface of mitochondria could be visualized by electron cryo-tomography [17]. Messengers RNA targeting to the surface of mitochondria has been also shown in plants [18, 19], and a protein named Friendly (FMT) was identified as a cytosolic RNA binding protein that binds cytosolic ribosomes at the surface of mitochondria [20]. It has been suggested [21] that localized translation occurs at most intracellular organelles. However, very little information is available on mechanisms controlling organelle-coupled translation in higher plants, and the in depth characterization of specific cytosolic ribosomes associated with organelles membranes remains to be performed. To this aim, this protocol describes the purification of pure and intact chloroplast and mitochondria from higher plant and the procedures available to isolate cytosolic ribosomes associated to these organelles.

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Materials

2.1 Plant Lines and Growth

Arabidopsis thaliana Columbia wild type, as well as the RPL18FLAG line [22] in the Columbia ecotype, were used for organelle purification and immunoprecipitation of cytosolic ribosomes (see Note 1).

2.1.1 Growth Condition for Chloroplast Purification

1. Prepare large (30 cm × 45 cm) plastic trays filled with compost and generously watered. 2. Sow Arabidopsis thaliana seeds onto the surface of the compost by scattering them carefully at a high density (around 30 mg of seeds for a whole tray). 3. Grow up to a 60-day-old rosette stage in a standard growth chamber under a short-day light period (12 h light cycle, with a light intensity of ca. 150 μmol.m-2.s-1) and a temperature of 23 °C during the day and 18 °C at night. One tray containing such Arabidopsis plantlets is expected to provide 100–150 g of rosette leaf material and is considered enough to prepare one sample of plastid-associated cytosolic ribosomes.

2.1.2 Growth Condition for Mitochondria Purification

1. Sow Arabidopsis thaliana seeds onto a pot and transplant them at 2–3 rosette leaves stage in trays (30 cm × 45 cm). Prepare 4–6 trays with 60 plants in each tray. 2. Grow up plants to flowering stage in long-day conditions (16: 8 h light/dark photocycle at 21 °C and 18 °C cycles; LED tubes Philips 1500 mm SO 20 W 840 T8; Philips, Eindhoven, the Netherlands; photon flux density of 120 μmol.m-2.s-1 at the plant level). Four trays can provide around 30 g of inflorescences to obtain 1–2 mg of purified mitochondria.

2.2 Purification of Chloroplasts from Arabidopsis Leaves

1. Muslin or cheesecloth, 80 cm large.

2.2.1

3. Beakers (5 L).

Material

2. Nylon blutex (50 μm aperture) (from providers such as Tripette et Renaud, Sailly Saillisel, France). 4. Ice and ice buckets. 5. Pipettes (1 and 10 mL). 6. Percoll (from providers such as GE Healthcare or SigmaAldrich ).

2.2.2

Equipment

1. Motor-driven blender, three speeds, 1 gallon (3.785 L) capacity (from providers such as Waring blender) (see Note 2). 2. Refrigerated high-speed centrifuge (such as Aventi JE, from Beckman Coulter ).

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3. Rotors: a JLA 10.500 rotor (6 × 500 mL plastic bottles), fixedangle rotors JA-20 (6 × 36 mL tubes), swinging-bucket rotor JS-13.1 (6 × 36 mL tubes). 2.2.3

Stock Solutions

1. 0.5 M Ethylenediaminetetraacetic acid (EDTA) (from providers such as Euromedex). 2. 1 M Tricine-KOH (from providers such as Sigma-Aldrich), pH 8.4. 3. 1 M 3-(N-morpholino) propane sulfonic acid (MOPS)-NaOH, pH 7.8 (from providers such as Sigma-Aldrich). 4. Phenylmethylsulfonyl fluoride: 100 mM PMSF (from providers such as Sigma-Aldrich). 5. 100 mM Benzamidine (from providers such as Sigma-Aldrich). 6. 50 mM ε-amino caproic acid (from providers such as SigmaAldrich). 7. 1 M MgCl2 (from providers such as ROTH).

2.2.4

Working Solutions

1. Leaf grinding medium: 20 mM Tricine-KOH, pH 8.4, 0.4 M sorbitol, 10 mM EDTA, 10 mM NaHCO3, 0.1% (w/v) bovine serum albumin (BSA, defatted). 2. Chloroplast washing medium: 20 mM Tricine-KOH, pH 7.6, 0.4 M sorbitol, 5 mM MgCl2, 2.5 mM EDTA. 3. Hypotonic medium for chloroplast lysis: 10 mM 3-(N-morpholino) propane sulfonic acid (MOPS)-NaOH, pH 7.8, 1 mM PMSF, 1 mM benzamidine, and 0.5 mM ε-amino caproic acid. 4. Bradford dye-binding solution (Bio-Rad protein assay) to determine protein concentration.

2.3 Purification of Mitochondria from Arabidopsis Inflorescence

1. Sucrose.

2.3.1

4. Polyvinylpyrrolidone-40 (PVP-40) (Sigma-Aldrich).

Material

2. Tetrasodium pyrophosphate (10.H2O) (from providers such as Sigma-Aldrich). 3. BSA (from providers such as Euromedex). 5. MOPS (from providers such as Sigma-Aldrich). 6. Ethylenediaminetetraacetic acid (EDTA) (from providers such as Euromedex). 7. L-Cysteine (from providers such as Sigma-Aldrich). 8. Potassium dihydrogen phosphate (KH2PO4) (from providers such as Carlo Erba). 9. L-Ascorbic acid.

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10. Ethyleneglycol-bis(β-aminoethyl)-N,N,N′,N′-tetraacetic acid (EGTA) (from providers such as Sigma-Aldrich). 11. Percoll (from providers such as GE Healthcare or SigmaAldrich). 12. Miracloth (from Millipore). 13. Phosphoric acid 85% (from providers such as Sigma-Aldrich). 14. Potassium hydroxide (KOH). 15. Bradford dye-binding solution (from Bio-Rad protein assay). 2.3.2

Equipment

1. Corning PYREX Tissue Grinder Dounce of 15 mL and 2 mL. 2. Commercial blender with a bucket of 200 mL capacity. 3. AVANTI-JE Beckman Coulter centrifuge or equivalent. 4. Rotors: JA25.5 fixed-angle rotor 8 × 50 mL Beckman Coulter and JLA10.500 fixed-angle rotor 6 × 500 mL Beckman, or equivalent. 5. Round Centrifuge Tube Polycarbonate, 50 mL (Nalgene or equivalent). 6. Centrifuge Bottle with SealingCap, PPCO, 500 mL (Nalgene or equivalent). 7. Bench centrifuge (e.g., the 5415R microcentrifuge from Eppendorf or equivalent). 8. 1.5 mL and 2 mL Eppendorf tubes or similar. 9. A round paint brush (n°10).

2.3.3

Stock Solutions

2.3.4

Working Solutions

1. 5 M KOH. The working solutions are prepared in advance and conserved at 4 ° C except for the gradient solutions prepared during the mitochondria purification. 1. Extraction buffer: 0.3 M sucrose, 15 mM tetrasodium pyrophosphate (10.H2O), 2 mM EDTA, 10 mM potassium phosphate (KH2PO4), 1% (w/v) polyvinylpyrrolidone-40, 1% (w/v) BSA, 20 mM L-ascorbic acid, and 5 mM L-cysteine, pH 7.5. To prepare this solution, sucrose, tetrasodium pyrophosphate (10.H2O), EDTA, potassium phosphate, and PVP-40 powders are mixed, and osmosis water is added to less than the desired volume. The pH is adjusted to 7.5 with phosphoric acid 85% and stored at 4 °C. The solution is completed to the desired volume and conserved at 4 °C. The BSA, L-ascorbic acid, and L-cysteine powders are added to the solution on the day of the mitochondria purification. 2. Washing buffer: 0.3 M sucrose, 10 mM MOPS, 1 mM EGTA, pH 7.2. The pH is adjusted with a 5 M KOH solution.

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3. Gradient buffer 5x: 1.5 M sucrose, 50 mM MOPS, pH 7.2. The pH is adjusted with a 5 M KOH solution. 4. Gradient solution 50%: 1x Gradient buffer, 50% (v/v) of Percoll. 5 mL per gradient. 5. Gradient solution 25%: 1x Gradient buffer, 25% (v/v) of Percoll. 25 mL per gradient. 6. Gradient solution 18%: 1x Gradient buffer, 18% (v/v) of Percoll. 5 mL per gradient. 2.4 Coimmunoprecipitation of Cytosolic Ribosomes

1. M columns (from Miltenyi Biotec).

2.4.1 Material and Equipment

3. MACS Multistand magnetic rack (from Miltenyi Biotec).

2. μMACS DYKDDDDK Isolation kit (from Miltenyi Biotec): Only the supplied FLAG-tagged beads and elution buffer are used. 4. Rotating mixer. 5. Benchtop refrigerated centrifuge for Eppendorf tubes. 6. Benchtop heating block for Eppendorf tubes. 7. Liquid nitrogen. 8. Ice and ice bucket. 9. Free Protease Inhibitor Cocktail (from providers such as Roche). 10. Equipment and solutions for SDS-PAGE and western blot analyses. 11. ANTI-FLAG antibody (from providers such as Merck) to analyze (western blot) the eluates of the co-immunoprecipitation experiments.

2.4.2

Stock Solutions

1. 0.2 M HEPES-KOH, pH 7.6 (from providers such as Euromedex). 2. 2 M KCl. 3. 1 M MgCl2. 4. 100 mM DTT (freshly prepared). 5. 25% Triton X-100 (from providers such as ROTH). 6. 100 mM PMSF for chloroplast samples (from providers such as Sigma-Aldrich) or Free Protease Inhibitor Cocktail 25x for mitochondria samples. 7. 50 mg.mL-1 Cycloheximide (from providers such as SigmaAldrich, catalog number: C7698-5G).

2.4.3

Working Solutions

1. Lysis buffer: 20 mM HEPES-KOH, 100 mM KCl, 20 mM MgCl2, 1 mM DTT, 1% (w/v) Triton X-100, 1 mM PMSF for

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chloroplast samples or 1x Free Protease Inhibitor Cocktail for mitochondria samples, 20 μg/mM cycloheximide. 2. Washing buffer: 20 mM HEPES-KOH, 100 mM KCl, 20 mM MgCl2, 1 mM DTT, 0.1% (w/v) Triton X-100, 1 mM PMSF for chloroplast samples or Free Protease Inhibitor Cocktail for mitochondria samples, 20 μg/mM cycloheximide.

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Methods For organelles purification, all the steps should be carried out at 0–5 °C, either on ice or in a cold room. Preferentially use pre-cooled tubes and glassware.

3.1 Purification of Chloroplasts from Arabidopsis Leaves

The protocol used to purify intact chloroplasts was adapted from a previously described protocol [23]. Before the experiment, prepare two tubes (each containing 30 mL of a 50% Percoll/chloroplast washing medium. Pre-form Percoll gradients for further chloroplast purification by centrifugation at 18,000 rpm (40,000 g) for 55 min (JA-20 rotor). Store the tubes containing preformed Percoll gradients in the cold room until use. Preparation of Percollpurified chloroplasts should take 2.5 h knowing that harvesting of the rosette leaves is performed during the centrifugation required to pre-form the Percoll gradients (55 min).

3.1.1 Preparation of a Crude Chloroplast Suspension

1. Harvest 100–150 g of rosette leaves and transfer them into a cold room for the next step. 2. Homogenize the 100–150 g of leaf material with 1 L of Leaf Grinding Medium three times, for 1 s each time, in a blender. Filter the homogenate rapidly through four layers of muslin and one layer of nylon blutex. Keep the homogenate in a beaker and re-collect the grinded leaves remaining on the muslin. 3. Repeat step 2 with the grinded leaves by homogenization in 1 L of Leaf Grinding Medium. Add this homogenate to the first one. 4. Distribute equally the filtered suspension into two bottles for centrifugation and centrifuge them at 3500 rpm (2200 g) for 2 min (JLA 10.500 rotor, see Note 3). 5. Aspirate the supernatant with a pipet fitted to a water pump. 6. Carefully add 2 mL of Chloroplast Washing Medium to each crude chloroplast fraction pellet. Use a paintbrush to gently resuspend the organelles.

3.1.2 Percoll Purification of Intact Chloroplasts

1. Distribute the crude chloroplast fraction carefully on the top of the two preformed Percoll gradients. Centrifuge the gradients

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at 9000 rpm (12,600 g; see Note 3) for 10 min (swingingbucket rotor JS-13.1). 2. Remove the upper part of the gradient (including the upper green band representing broken chloroplasts) and then recover with a pipette the intact chloroplasts present in the broad, darkgreen band in the lower part of the gradient. The intact chloroplast fraction is collected in two 50 mL Falcon tubes. 3. Dilute the intact chloroplast fraction by filling up to 50 mL with Chloroplast Washing Medium. Centrifuge the suspension at 3000 rpm (1080 g) for 2 min (Eppendorf Centrifuge 5804R). 4. Add 150 μL of hypotonic medium to each pellet containing washed and purified chloroplasts. 3.2 Purification of Mitochondria from Arabidopsis Inflorescence

This protocol is based on the previously described protocol [24]. Finish preparing all the working solutions and keep them at 4 ° C or do everything in the cold room. All centrifugation steps are done at 4 °C, and always use the low break for centrifugation. Mitochondria purification should take half a day.

3.2.1 Preparation of a Crude Mitochondria Suspension

1. Collect the inflorescence of plants and put them on an aluminum foil placed on an ice bucket during the collection process. Weigh the material and transfer them to the cold room. Collect around 30 g of material per gradient. Keep 500 mg of inflorescence for the co-immunoprecipitation Subheading 3.3, which will serve as a control. 2. Place the 30 g of inflorescence in the bucket blender and cut them into small pieces using scissors. 3. Add 50 mL of extraction buffer and grid: 5 s high-speed – 30 s of pause – 5 s of low speed – 30 s of pause – 5 s of low speed. 4. Filter the solution through two layers of disposable Miracloth on a funnel and collect on a 500 mL centrifuge bottle the flowthrough. Squeeze the filter to extract all the liquid. 5. Recover the material that did not go through the Miracloth and put it back in the bucket blender with 50 mL of extraction buffer. 6. Grind the material a second time, as indicated before. 7. Filter the solution through the same two layers of disposable Miracloth. Rinse the bucket blender with 50 mL of extraction buffer, and collect the flow-through in the same 500 mL centrifuge bottle (squeeze until the last drop). 8. Centrifuge the filtrated solution for 5 min at 1500 g (JLA10.500 rotor) and transfer the supernatant into a new 500 mL centrifuge bottle and

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9. Centrifuge it for 10 min at 17,700 g (JLA10.500 rotor) and discard the supernatant. 10. Resuspend the pellet in 5 mL of washing buffer (using the paintbrush) and homogenize well with the 15 mL Tissue Grinder Dounce (3–4 maximum strokes to eliminate all aggregates). 11. Transfer the 5 mL to a 50 mL round polycarbonate centrifuge tube and add 30 mL of washing buffer. 12. Centrifuge for 5 min at 2000 g (JA25.5 rotor) and recover the supernatant in a new centrifuge tube. 13. Centrifuge for 10 min at 20,000 g and discard the supernatant. 14. Resuspend the pellet in 500 μL of washing buffer (using the paintbrush) and homogenize well with the 2 mL Tissue Grinder Dounce (3–4 maximum strokes to eliminate all aggregates). 15. Transfer the solution to a 2 mL tube. Rinse the Tissue Grinder Dounce with washing buffer. Do not exceed 2 mL for total volume. 3.2.2 Percoll Purification of Intact Mitochondria

1. Prepare two gradients during the differential centrifugations. Each gradient consists of, from bottom to top, a 5 mL solution gradient of 50% Percoll, a 25 mL solution gradient of 25% Percoll, and a 5 mL solution gradient of 18% Percoll. Put first the solution gradient of 50% Percoll on the bottom of the 50 mL round Polycarbonate centrifuge. Then, add the other solutions sequentially carefully and slowly. In the end, the different layers of the gradient are visible. 2. The crude mitochondria solution is gently layered onto the gradient. Do not overload the gradients (maximum loading: 2 mL of crude mitochondria extracted from 30 g of starting material). 3. Centrifuge for 45 min at 40,000 g with the break-off (JA25.5 rotor). 4. Mitochondria will form a white/yellow band at the top of the solution gradient 50% layer (Fig. 1). Remove with a vacuum pump the green band in the top part of the gradient. Collect the mitochondria by plunging the pipette tip into the mitochondria band using a 1 mL pipette with the 1 mL tip cut to have a larger opening. A 5 mL pipette can also be used. 5. Pool all mitochondria together and dilute at least 5 times the volume with wash buffer. 6. Centrifuge for 15 min at 18,000 g (JA25.5 rotor). 7. Discard with a vacuum pump most of the supernatant. Be careful; the pellet is loose at this step. Keep around 3–5 mL per tube and resuspend the pellet by swirling the tube. Add

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Fig. 1 Isolation of cytosolic ribosomes associated with mitochondria. Crude and purified mitochondria are first prepared from Arabidopsis thaliana inflorescences of FLAG-RPL18 line or wild type (WT). Co-immunoprecipitation (Co-IP) with anti-FLAG beads is then performed. Eluates are then analyzed, for example, by western blot (CB, Coomassie blue staining of the membrane; FLAG, FLAG antibody) or mass spectrometry. M, Mitochondria, T, Total extract

25 mL of washing buffer. The goal of this step is to remove the majority of Percoll from the mitochondria suspension. 8. Centrifuge for 15 min at 18,000 g (JA25.5 rotor). 9. Discard, as previously, the supernatant and resuspend the pellet in 1 mL of washing buffer by swirling the tube or pipetting (be careful). At this step, the mitochondria pellet should be stable. If not, do additional washing steps until obtaining a tight pellet. 10. The quantity of mitochondria is evaluated by performing a Bradford test. To get an accurate measure, dilute mitochondria with water at least ten times. At least 1–2 mg of mitochondria is obtained from 30 mg of inflorescence. 11. Do aliquots of 1 mg of purified mitochondria into 1.5 mL tubes. Centrifuge for 5 min at 16,000 g. Discard the entire supernatant, flash freeze the aliquots in liquid nitrogen, and store them at -80 °C. 3.3 Purification of Cytosolic Ribosomes from Organelles 3.3.1 Preparation of Samples for the Coimmunoprecipitation

Two samples are used for the co-immunoprecipitation. The sample named “Total” corresponds to the cytosolic ribosomes present in the whole plant cell, while the sample named “mito” or “chloro” corresponds to the cytosolic ribosomes associated with the organelles (Fig. 1). 1. For chloroplasts: Grind some fresh leaves in liquid nitrogen for the “total” sample. 100 mg of frozen powdered leaves is

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necessary for co-immunoprecipitation. For the “chloro” sample, use purified chloroplasts volume equivalent to 1 mg proteins. 2. For mitochondria: Grind the inflorescence in liquid nitrogen for the “total” sample. 0.1 mg of frozen powdered inflorescence is necessary for co-immunoprecipitation. For the “mito” sample, use pelleted purified mitochondria equivalent to 1 mg proteins. 3.3.2 Coimmunoprecipitation

Steps 1 through 5 should be conducted at 0–5 °C on ice or in a cold room. The lysis buffer should be pre-cooled, and the necessary volume of elution buffer pre-heated at 95 °C before use. 1. Resuspend samples with 1 mL of lysis buffer. 2. Incubate the tubes for 30 min at 4 °C on a rotating mixer (5–10 rpm). 3. Centrifuge for 10 min at 10,000 g at 4 °C and collect the supernatants. 4. Add 50 μL of beads to the lysates. Be careful to mix the beads before use. First, homogenize the tubes by gentle inversions and incubate them like in step 2. 5. Concurrently, insert M columns (see Note 4) in the magnetic rack and equilibrate them with 200 μL lysis buffer. 6. Add the lysates with beads on the M columns 200 μL by 200 μL and leave the liquid running through. Be careful to add in the center of the column without touching the edges and not putting any air bubbles. Discard what remains in the tip and change the tip each time. 7. Wash the M columns four times with 200 μL of washing buffer. 8. Add 40 μL of pre-heated elution buffer on the M columns to eliminate the void volume. 9. Detach the column from the magnetic rack and place it in a 2 mL tube. Add additional volumes of pre-heated elution buffer (30–35 μL, three times) and collect the eluted cytosolic ribosomes on fresh tubes (see Note 5).

3.3.3 Quality Control of the Purified Cytosolic Ribosomes

1. Load the eluates of the co-immunoprecipitation experiments (including negative controls from wild type) on a SDS-PAGE (Fig. 1). 2. Perform a western blot using the ANTI-FLAG® antibody to validate the presence of the FLAG RPL-18 protein in the eluates (total extract, purified mitochondria, or chloroplasts) derived from the FLAG-RPL-18 line samples (Fig. 1). 3. Analyze the protein composition of eluates (including negative controls) using mass spectrometry. If performed accordingly,

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Fig. 2 MS-based quantitative proteomic analyses of the cytosolic ribosomes purified from total cell extracts (left panel) and Percoll-purified chloroplast fractions (right panel). The Volcano plot displays the differential abundance of proteins in Flag-RPL18 and wild type (WT = negative control) co-immunoprecipitation eluates analyzed by MS-based label-free quantitative proteomics. The -log10 (limma p-value) on y axis is plotted against the log2(Fold Change Flag-RPL18/WT) on x axis. Blue dots represent proteins found enriched with Flag-RPL18 compared to WT eluates (Fold change ≥ 2 and p-value ≤ 0.01). Yellow dots represent proteins found enriched with WT compared to Flag-RPL18 eluates (Fold change ≥ 2 and p-value ≤ 0.01). Grey dots: no significant enrichment

this method allows the detection of proteins from the cytosolic ribosome that are enriched (Fig. 2) in eluates from mitochondrial or chloroplast derived fractions of FLAG-RPL18 line compared to the corresponding eluates from wild type plants (see Notes 6 and 7 for typical results obtained for cytosolic ribosomes issues from Percoll-purified mitochondria and chloroplasts, respectively).

4 Notes 1. Crude cell extracts (Total) and purified mitochondria or chloroplasts from Arabidopsis must be prepared from both FLAGRPL18 line and wild type (Col0) grown in the same conditions. The FLAG-RPL18 line expresses the RPL18 subunit of the cytosolic ribosome fused to a FLAG epitope [22]. After co-immunoprecipitation (Co-IP) of cytosolic ribosomes with anti-FLAG beads, the samples (crude cell extracts, mitochondria, and chloroplasts) purified from the wild type line will be used as negative controls to measure the enrichment of specific proteins in the samples derived from the FLAG-RPL18 line (see Figs. 1 and 2). 2. A 0.325-gallon (1.230 L) blender is recommended for lower amounts of leaf material.

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3. g values are given for the maximum radius of the given rotors. 4. μ columns can also be used instead of M columns. When the M column is used, it should not run dry. 5. A pipetting pear must be used to push out the eluate from the M column completely. 6. Using this method, 185 proteins were found enriched in the eluate from mitochondrial fraction of FLAG-RPL18 line compared to the eluate of Col0, and 111 out of the 185 proteins were cytosolic ribosome subunits [25]. Among these 185 proteins, 80% were also detected in eluates from FLAG-RPL18 total (inflorescences) extracts. However, a few proteins appeared specifically enriched in the mitochondrial fraction. Among them, one is the FMT protein, a member of the CLUSTERED MITOCHONDRIA (CLU) family, which is required for the correct distribution of mitochondria within the cell [20, 26]. 7. Following this procedure, it was also possible to purify cytosolic ribosomes present in the total (leaves) and the chloroplast fractions derived from the FLAG-RPL18 line (Fig. 2). As expected, most (82 proteins out of 92) proteins enriched in eluates derived from the chloroplast fraction of the FLAGRPL18 line compared to chloroplast fraction of Col0 were isoforms of subunits from the cytosolic ribosome. As observed for mitochondria, a few other proteins specifically enriched in the chloroplast fraction derived from the FLAG-RPL18 line were yet uncharacterized proteins.

Acknowledgments This work was supported by the CNRS, INRAE, the French Agence Nationale de la Recherche (ANR-18-CE12-0021-01 “Polyglot” and ANR-22-CE12-0012-01 “C-Trap”) and the Labex GRAL, funded within the University Grenoble Alpes graduate school (Ecoles Universitaires de Recherche) CBH-EUR-GS (ANR-17-EURE-0003). This work of the Interdisciplinary Thematic Institute IMCBio, conducted as part of the ITI 2021–2028 program of the University of Strasbourg, CNRS and Inserm, was supported by IdEx Unistra (ANR-10-IDEX-0002), STRAT’US (ANR 20-SFRI-0012), and EUR IMCBio (ANR-17-EURE0023) under the framework of the French Investments for the Future Program. L.D. got a fellowship from “Polyglot” and S.P. got a fellowship from “C-Trap.”

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References. 1. Rolland N, Curien G, Finazzi G et al (2012) The biosynthetic capacities of the plastids and integration between cytoplasmic and chloroplast processes. Annu Rev Genet 46:233–264 2. Møller IM, Rasmusson AG, Van Aken O (2021) Plant mitochondria – past, present and future. Plant J 108:912–959 3. Jarvis P, Lo´pez-Juez E (2013) Biogenesis and homeostasis of chloroplasts and other plastids. Nat Rev Mol Cell Biol 14:787–802 4. Sun Y, Jarvis RP (2023) Chloroplast proteostasis: import, sorting, ubiquitination, and proteolysis. Annu Rev Plant Biol 74:259–283 5. Murcha MW, Kmiec B, Kubiszewski-Jakubiak S et al (2014) Protein import into plant mitochondria: signals, machinery, processing, and regulation. J Exp Bot 65:6301–6335 6. Miras S, Salvi D, Ferro M et al (2002) Non canonical transit peptide for import into the chloroplast. J Biol Chem 277:47770–47778 7. Miras S, Salvi D, Piette L, Seigneurin-Berny D et al (2007) TOC159- and TOC75independent import of a transit sequence less precursor into the inner envelope of chloroplasts. J Biol Chem 282:29482–29492 8. Villarejo A, Bure´n S, Larsson S et al (2005) Evidence for a protein transported through the secretory pathway en route to the higher plant chloroplast. Nat Cell Biol 7:1224–1231 9. Moyet L, Salvi D, Bouchnak I et al (2019) Calmodulin is involved in the dual subcellular location of two chloroplast proteins. J Biol Chem 294:17543–17554 10. Ducheˆne AM, Drouard L (2021) Last news on plant mitochondria—a follow-up of ABR volume 63: mitochondrial genome evolution. Chapter 9. Adv Bot Res 100:275–229 11. Ferro M, Brugie`re S, Salvi D et al (2010) AT_CHLORO, a comprehensive chloroplast proteome database with subplastidial localization and curated information on envelope proteins. Mol Cell Proteomics 9:1063–1084 12. Bouchnak I, Brugie`re S, Moyet L et al (2019) Unravelling hidden components of the chloroplast envelope proteome: opportunities and limits of better MS sensitivity. Mol Cell Proteomics 18:1285–1306 13. Weis BL, Schleiff E, Zerges W (2013) Protein targeting to subcellular organelles via mRNA localization. Biochim Biophys Acta 1833:260– 273 14. Sun Y, Valente-Paterno M, Bakhtiari S et al (2019) Photosystem biogenesis is localized to

the translation zone in the chloroplast of Chlamydomonas. Plant Cell 31:3057–3072 15. Lesnik C, Golani-Armon A, Arava Y (2015) Localized translation near the mitochondrial outer membrane: an update. RNA Biol 12: 801–809 16. Lashkevich KA, Dmitriev SE (2021) mRNA targeting, transport and local translation in eukaryotic cells: from the classical view to a diversity of new concepts. Mol Biol 55:507– 537 17. Gold VA, Chroscicki P, Bragoszewski P et al (2017) Visualization of cytosolic ribosomes on the surface of mitochondria by electron cryotomography. EMBO Rep 18:1786–1800 18. Michaud M, Mare´chal-Drouard L, Ducheˆne AM (2010) RNA trafficking in plant cells: targeting of cytosolic mRNAs to the mitochondrial surface. Plant Mol Biol 73:697–704 19. Vincent T, Vingadassalon A, Ubrig E et al (2017) A genome-scale analysis of mRNAs targeting to plant mitochondria: upstream AUGs in 5′ untranslated regions reduce mitochondrial association. Plant J 92:1132–1142 20. Hemono M, Salinas-Giege´ T, Roignant J et al (2022) FRIENDLY (FMT) is an RNA binding protein associated with cytosolic ribosomes at the mitochondrial surface. Plant J 112:309– 321 21. Be´thune J, Jansen RP, Feldbru¨gge M et al (2019) Membrane-associated RNA-binding proteins orchestrate organelle-coupled translation. Trends Cell Biol 29:178–188 22. Zanetti MA, Chang IF, Gong F et al (2005) Immunopurification of polyribosomal complexes of Arabidopsis for global analysis of gene expression. Plant Physiol 138:624–635 23. Moyet L, Salvi D, Tomizioli M et al (2018) Preparation of membrane fractions (envelope, thylakoids, grana, and stroma lamellae) from Arabidopsis chloroplasts for quantitative proteomic investigations and other studies. Methods Mol Biol 1696:117–136 24. Waltz F, Nguyen TT, Arrive´ M et al (2019) Small is big in Arabidopsis mitochondrial ribosome. Nat Plants 5:106–117 25. Salih KJ, Duncan O, Li L et al (2020) The composition and turnover of the Arabidopsis thaliana 80S cytosolic ribosome. Biochem J 477:3019–3032 26. El Zawily AM, Schwarzlander M, Finkemeier I et al (2014) FRIENDLY regulates mitochondrial distribution, fusion, and quality control in Arabidopsis. Plant Physiol 166:808–828

Part III In Silico Tools

Chapter 19 A Practical Guide to the Representation of Protein Regulation in the Web Application ChloroKB Gilles Curien Abstract ChloroKB (http://chlorokb.fr) is a knowledge base providing synoptic representations of the metabolism of the model plant Arabidopsis thaliana and its regulation. Initially focused on plastid metabolism, ChloroKB now accounts for the metabolism throughout the cell. ChloroKB is based on the CellDesigner formalism. CellDesigner supports graphical notation and listing of the corresponding symbols based on the Systems Biology Graphical Notation. Thus, this formalism allows biologists to represent detailed biochemical processes in a way that can be easily understood and shared, facilitating communication between researchers. In this chapter, we will focus on a specificity of ChloroKB, the representation of multilayered regulation of protein activity. Information on regulation of protein activity is indeed central to understanding the plant response to fluctuating environmental conditions. However, the intrinsic diversity of the regulatory modes and the abundance of detail may hamper comprehension of the regulatory processes described in ChloroKB. With this chapter, ChloroKB users will be guided through the representation of these sophisticated biological processes of prime importance to understanding metabolism or for applied purposes. The descriptions provided, which summarize years of work and a broad bibliography in a few pages, can help speed up the integration of regulatory processes in kinetic models of plant metabolism. Key words Allostery, Arabidopsis, CellDesigner, Modeling, Plant metabolism, Protein regulation

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Introduction When implementing new pathways (for example with the current attempts to optimize photosynthesis [1–4]), researchers need to appreciate how the plant functions as a whole. Mathematical modeling is expected to contribute to the discovery of optimal solutions, and steady-state modeling of metabolic systems (Flux balance analysis) is now widely used [5, 6]. However, the kinetics of metabolic systems and their regulation must also be taken into account [7, 8]. Indeed, the time it takes for a living system to recover after a perturbation determines, in fine, the amount of biological material the organism can accumulate (i.e., the yield in an agronomical context). The speed of this recovery depends on the kinetic

Eric Mare´chal (ed.), Plastids: Methods and Protocols, Methods in Molecular Biology, vol. 2776, https://doi.org/10.1007/978-1-0716-3726-5_19, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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structure of the underlying metabolic systems [7]. This structure includes enzymatically catalyzed reactions, transport steps, and all the biochemical processes generally described as “regulation,” (see Note 1) specifically targeting individual enzymes and transporters in the metabolic network. As regulatory processes are diverse in nature, and some are exquisitely complicated, they may be difficult to implement in mathematical models without the aid of facilitating tools. The initial motivation behind the development of ChloroKB (http://chlorokb.fr) was to graphically represent metabolic systems and the regulation of the proteins involved as a means to facilitating mathematical modeling. The CellDesigner graphical notation [9, 10] allows precise description of the molecular interactions involved in the regulation of protein activity (protein oligomerization, protein assembly, binding of allosteric effectors, posttranslational modification). Protein modifications can be represented as interconversions of protein-protein and protein-ligand(s) complexes. These representations gather together information derived from both enzyme kinetics and three-dimensional protein analyses. When activation/inhibition of a given enzyme or transporter can be illustrated in such a way, the construction of mathematical equations is facilitated. This also paves way for higher-level integration and understanding. However, due to the complexity of regulatory processes, the representations used in ChloroKB might not be immediately comprehensible to users. The aim of this chapter is thus to help ChloroKB users to familiarize themselves with ChloroKB codes and the biochemical processes represented. Despite their apparent complexity, the metabolic maps in ChloroKB can be seen as a summary of years of research in a single picture or a limited set of pictures. As such, they should facilitate interpretation, communication, knowledge acquisition, and – in fine – mathematical modeling. ChloroKB could also be a useful tool outside the field of plant metabolic modeling, for instance for teaching or for the acquisition of knowledge by students to better understand and rationalize complicated biochemical processes such as lipid synthesis, molecular machine assembly, or enzyme activity regulation. ChloroKB is a regularly updated community resource.

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Material 1. ChloroKB (http://chlorokb.fr) has been extensively tested using Google Chrome and Safari. Tests on IE10 and Firefox39 indicate that browsing is possible but that some functionalities will not work. We therefore recommend the use of Chrome. 2. Users are reminded to clear browsing data to ensure any modifications on maps following updates are visible and to refresh the page(s) regularly.

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Methods 1. ChloroKB was designed to facilitate the storage, organization, and understanding of the metabolic network in Arabidopsis thaliana and its regulation [11, 12]. Figure 1 provides a schematic description of the organization of ChloroKB. It shows the different biological levels represented along with the three main types of information available (maps, textual information, tools for modeling).

3.1 ChloroKB Overview

2. Figure 2 provides an overview of ChloroKB content as displayed on the home page (http://chlorokb.fr). On this page, the user will find different interconnected processes grouped based on function (e.g., nucleotide metabolism, detoxification, pigments, and lipid metabolism). The clickable purple hexagons are links to maps.

Biological level adressed

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Fig. 1 Schematic representation of the organization of ChloroKB. Top: the different levels of detail displayed in ChloroKB and available as graphical representations: left, large overviews of available reconstructed processes (accessible from the home page); middle, representation of integrated processes at the cellular level; right, synoptic representation of integrated biochemical processes (e.g., photorespiration, phosphate transport, and metabolite hubs) and their interconnections. The view is comprehensive: all the proteins and metabolites involved in a specific process are represented. The cell represented in ChloroKB with all its subcelllar compartments is an “ideal” or “generic” cell: the proteins displayed in the maps are not necessarily all expressed in the same tissue or simultaneously. aChloroKB visualization requires Google Chrome. It also works with Firefox but with some degradation. bUnknown proteins are proteins for which the genes are as yet unknown but the presence of which has either been deduced from network structures or from biochemical analyses (e.g., transport using isolated organelles). In contrast with other databases, the presence of unknown proteins in ChloroKB allows literature information on protein functions and properties to be stored. cAutomatic building of a stoichiometry matrix is an option not described here. dQuantitative data related to metabolite abundance: a summary document is available on the website

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Welcome page

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Maps connexions illustrated with « dynamic » links in red (mouse-over tool)

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157 different processes reconstructed

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Fig. 2 Overview of the 200 maps currently available in ChloroKB. From the home page (a), the user can explore the initial network reconstructed with its interconnected maps (red), and connections appear dynamically using the mouse-over tool (b). A sub-part of the network dedicated to lipid synthesis can be accessed from the home page (c). With time, the size of the network has increased, and newly reconstructed processes no longer fit in the home page map: the content of the entire network can now be revealed by clicking the “All maps” purple hexagon (See a and d). The “All maps” map (d) lists the 200 biochemical processes reconstructed in ChloroKB, sorted in alphabetical order, including 42 maps dedicated to the representation of regulatory processes (right). Zooming on map functions can be achieved either with two fingers on the computer touch-pad/cell phone screen or with the navigation tool located in the upper left of the maps

3. Maps present the molecular entities involved in a given cellular function (e.g., photorespiration and fatty acid synthesis) and their interactions. The arrows between the hexagons represent the flow of carbon from one process to another for the major biosynthetic/biodegradative “pathways.” To fit within the space available on this home page, only a partial representation of the Arabidopsis metabolic network is provided. 4. By clicking on the “All maps” purple hexagon (see bottom left of Fig. 2), users can view the whole of the reconstructed network available in ChloroKB, with maps listed in alphabetical order (based on pathway). Biochemical processes involving complex enzymes and their regulation, which is the focus of the present chapter, are grouped on the right of this map, once

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again in alphabetical order (see Fig. 2, bottom right). On the home page, the mouse-over tool can be used to highlight connections between processes. For example, when sliding the cursor over “pyruvate HUB chloroplast,” the black lines change to red (Fig. 2), to indicate, e.g., the connections between pyruvate metabolism and acetyl-coA metabolism and the different downstream products of pyruvate in the plastid (arginine, L-branched chain amino acids, L-cysteine, and lipid synthesis). 5. Specific terms (e.g., vitamin and Calvin cycle) can be located using the Ctrl+F keyboard shortcut, and then clicked. The corresponding map will be displayed in a new webpage. Alternatively, key words can be typed into the search field (See [11]). In this chapter, three different enzymatic steps occurring partly in the plastid will be used to illustrate and explain the representational codes used in ChloroKB: (1) pyruvate kinases (representation of allosteric regulation), (2) pyruvate phosphate dikinase (representation of protein activity regulation by phosphorylation), and (3) ADP-glucose pyrophosphorylases (multilayered control of protein activity). Once the representations of these three examples are understood, ChloroKB users should be able to confidently navigate the other maps. We welcome all comments or reports of errors to continue to improve this community resource. 3.2 Representation of Allosteric Regulation

1. Graphical exploration of ChloroKB maps can proceed from a low to a high level of complexity, ending up with detailed representations of how protein activity is regulated. Protein activity regulation involves an equilibrium between protein complexes, ligand binding, and posttranslational modifications. 2. As a first example, we will consider pyruvate metabolism and pyruvate kinases (Fig. 3). Starting from the home page, using Ctrl+F on the keyboard and entering “pyruvate HUB” will highlight the maps containing these words. Clicking on the “pyruvate HUB chloroplast” purple hexagon opens a new page, showing the corresponding map, a zoom of which is displayed in Fig. 3a. The user can then either navigate in the map to look for “pyruvate kinases” or use the keyboard shortcut Ctrl+F and enter “pyruvate kinases” to highlight the occurrence of these terms in the map (see Fig. 3b). 3. Clicking on the “pyruvate kinases” icon opens a third page with the map displayed in the background of Fig. 3c. This map shows the 14 enzymes coded by the Arabidopsis genome (see Note 2) and how they are distributed in the cell. Four pyruvate kinases are represented in a compartment surrounded by two yellow “membranes” (the plastid), three pyruvate kinases are

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a Zoom on Pyruvate HUBs on the Welcome page

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b Plastidial Pyruvate kinases in the context of pyruvate metabolism (Pyruvate HUB chloroplast map)

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cPK2-5: unknown oligomeric state

Uncharacterized Pyruvate kinases

Fig. 3 Representation of pyruvate kinases and their allosteric regulation in ChloroKB. (a) connection of the pyruvate HUB maps with the other reconstructed processes. The mouse-over tool highlights the connections (carbon flow) in red. Pyruvate kinases are present in three different maps: (b) a zoom on the “pyruvate HUB chloroplast” map showing pyruvate kinase in the context of pyruvate metabolism in the plastid; (c) the 14 pyruvate kinases predicted from the Arabidopsis cellular genome (see text for more details) and their cellular localizations; (d) screen capture of the “pyruvate kinases regulation” map showing their regulation

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positioned in the cytosol (in white), two pyruvate kinases are associated with the outer membrane of the mitochondrion (pink), and the final four pyruvate kinases have yet to be localized (red rectangles). Pyruvate kinase 4 is positioned in the plastid but is represented in red as its localization remains putative (more information is available on the website by clicking on the pyruvate kinase 4 rectangle). Pyruvate kinases that have not yet been biochemically characterized (i.e., with putative activity) can easily be identified: there are no links between the protein symbols (rectangles) and the pyruvate kinase reaction (the arrow with a square in the middle) connecting phosphoenolpyruvate and ADP to pyruvate and ATP products. The user can thus immediately grasp the current state of knowledge for a given biochemical process. 4. If more detail on pyruvate kinase regulation is required, the user may click on the “pyruvate kinases regulation” purple hexagon (see zoom in Fig. 3c). The corresponding map (background image in Fig. 3d) shows an up-to-date representation of the biochemical knowledge and the current unknowns surrounding regulation of pyruvate kinase activity in Arabidopsis. We will describe this map in detail. Active pyruvate kinases are complexes consisting of a polypeptide associated with potassium and magnesium ions. These complexes are represented by the fine line surrounding the graphical symbols for proteins (rectangles) and ions (disks). The name of the complex pops up if the user positions and immobilizes the cursor over the complex. 5. Complexes can be embedded inside one another, as illustrated for the heteromeric plastidial pyruvate kinase αβ (α, β1, and β2 subunits are coded by three different genes, the ID of which pops up when using the mouse-over tool). In Fig. 3d, plastid enzymes (left) are represented as hetero-octamers. These proteins are colored in orange (see Note 3) because they were detected in the plastid (the localization is explained in a comment on the webpage dedicated to the protein, opened by ä Fig. 3 (continued) (dashed-outline boxes were added here to explain content).The map shows inhibition of octameric plastid pyruvate kinase [(4α)(4β1)] by L-glutamate (top left), as well as activation and inhibition of pyruvate kinase [(4α)(4β2)] by 6-phosphogluconate and L-glutamate, respectively (bottom left) in the plastid. Control of cytosolic PK1 tetramer is represented in the middle (white), with activation by fructose 1,6 bisphosphate (see inset with the “plus” sign highlighted by the green circle) and inhibition by L-aspartate. Modulation of the enzyme activity is symbolized by the thickness of the link joining the protein and the reaction catalyzed. The thicker the link, the higher the activity. The inset on the right shows that two pyruvate kinases (cPK3 and CPK4) are associated with the mitochondrion, as symbolized by their pink color and contact with the mitochondrial outer membrane. The oligomeric state of cPK2-5 is unknown, and proteins are thus represented as monomers. The metabolic control of their activity has not yet been described

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clicking on the protein rectangle). The [(4α)(4β1)] heteromer pyruvate kinase (top left) is inhibited by interaction with the allosteric effector L-glutamate. The [(4α)(4β2)] heteromer pyruvate kinase (bottom left) has several effectors: the activator 6-phosphogluconate or the inhibitor L-aspartate. Modulation of the enzymatic activity by allosteric effectors is also symbolized by the thickness of the “catalysis” link (a black line ending with a small disk). 6. The catalysis link is the symbol connecting the protein complexes to the reaction symbol. Low activity is represented by a very thin line, moderate activity by a thin line, and high activity by a thick line. The stoichiometry of the allosteric effectors in the plastid pyruvate kinase heterooctamers is not known yet: a single molecule of effector per complex is thus represented (L-glutamate, L-aspartate, or 6-phosphogluconate). One cytosolic pyruvate kinase has been extensively described (cPK1, in white, center of the map, Fig. 3). cPK1 is a homotetramer that is regulated by fructose 1,6 bisphosphate (activator) and L-aspartate (inhibitor). 7. The CellDesigner code is implemented with additional codes in ChloroKB allowing for the addition of a “+” (plus) in green or a “-” (minus) in red, to indicate that the ligand is an activator or an inhibitor (see the zoom on the top right part of Fig. 3d). Information for cPK2-5 is scarce (see zoom on the right of Fig. 3d): The enzyme’s oligomerization states are unknown (enzymes are therefore represented as monomers), and no regulators have yet been identified. The five red rectangles (Fig. 3d, bottom) represent putative pyruvate kinases awaiting characterization (they are the same ones displayed in the overall pyruvate kinase map in Fig. 3c). The red dot at the bottom right of some complexes is of interest. Clicking on these dots will display additional information (complex components, reaction catalyzed, references, comments, and explanations). With the three maps displayed in Fig. 3, ChloroKB users can readily access up-to-date information on pyruvate kinases in Arabidopsis, assess the knowledge accumulated, and evaluate the missing information. 8. More complicated representations of allosteric regulation are available for the following example enzyme families: “aspartate kinases” (where the bipartite and tripartite domain structures of the enzymes are accounted for) and “mitochondrial L-cysteine synthase,” where allosteric controls are combined with protein association-dissociation.

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1. Modifications to protein activity by phosphorylation and its representation in ChloroKB will be illustrated through the example of pyruvate phosphate dikinase (PPDK) activity (Fig. 4). The PPDK reaction and its regulation by phosphorylation are good examples of complex processes and how they can be represented and thus communicated. PPDK synthesizes phosphoenolpyruvate (PEP) from pyruvate (i.e., it operates in the reverse direction compared to pyruvate kinase, described in Fig. 3). 2. The PPDK maps (entitled “pyruvate phosphate dikinase PPDK” and “pyruvate phosphate dikinase PPDK regulation”) can be searched for on the ChloroKB home page by typing “PPDK” in the search field. The user can then click on the thumbnail images of the maps on the left or on the name of the maps on the right. The “pyruvate phosphate dikinase PPDK” map can also be selected from the “pyruvate HUB” maps (Fig. 2a). The pyruvate phosphate dikinase PPDK map (Fig. 4a) shows two different cellular compartments, the plastid at the top and the cytosol at the bottom. The enzyme is dually localized; therefore, its name (PPDK1) is identical in both compartments. (Users can access the localization information contained in a comment on the dedicated protein page by clicking on the protein symbol in the website.) 3. Another way to see that the two proteins are products of the same gene is to hover the cursor over the plastid stroma protein (orange) and then on the cytosolic protein (white): after one second, the gene ID appears (AT4G15530 in both cases). The PPDK reaction (Fig. 4a) and its regulation by phosphorylation (Fig. 4b) are quite complicated, as both processes involve amino acid phosphorylation. 4. The CellDesigner graphical code allows biochemical details to be represented. We will start with the catalytic reaction (Fig. 4a, upper right for the plastid enzyme): Reaction of the protein with ATP is followed by the formation of a covalent bond between a histidine (His545) and pyrophosphate (PPi, see the black line between the protein and PPi in the complex, symbolizing a covalent link (see Note 4), top right inset in Fig. 4a). AMP is still bound to the protein as shown on top of the complex. Once the reaction with phosphate is complete, AMP is set free, and phosphate exchange occurs, PPi is set free and the protein now displays a single phosphate on His 545. This is symbolized by a “P” inside the small white disk close to “His545” (see inset on the top left). This phosphate group can then be transferred to pyruvate to form phosphoenolpyruvate and regenerate an unphosphorylated protein (inset on the right in Fig. 4a). This reaction is itself regulated by phosphorylation on another residue (Thr543, see Fig. 4b).

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zoom

a zoom

zoom

PPDK1 reaction in the stroma(Pyruvate=>PEP)

zoom

PPDK1 reaction in the cytosol (Pyruvate=>PEP)

PPDK1 regulatory protein 1

Phosphorylated, inactive stromal PPDK1

b

* PPDK1 reaction (plastid)

zoom

PPDK1 regulatory protein 2

zoom

Phosphorylated, inactive cytosolic PPDK1

PPDK1 reaction (cytosol)

Fig. 4 Pyruvate phosphate dikinase (PPDK) reaction and regulation by phosphorylation in ChloroKB. (a) Screen capture of the “pyruvate phosphate dikinase (PPDK)” map: the map shows that PPDK1 is targeted both to the plastid stroma (orange, top) and to the cytosol (white, bottom). In this illustration, boxes are superimposed on the chemical reactions. The PPDK reaction is rather complicated: After reaction with ATP, a transitory covalent link is formed between PPi and PPDK His545. Subsequent binding of a phosphate and phosphate exchange

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Phosphorylation of Thr543 inactivates the enzyme (the zoom function can be used to see this level of detail on the website). The proteins catalyzing this regulation are PPDK regulatory protein 1 in the stroma and PPDK regulatory protein 2 in the cytosol. As shown in the map (Fig. 4B left part), the PPDK regulatory proteins 1 and 2 are bifunctional, catalyzing both the phosphorylation (upper reaction) and the dephosphorylation (reverse direction below) of PPDK. 5. The map also shows that plastid stromal PPDK1 is modulated by light/dark cycles, being activated in the light and inactivated in the dark as a consequence of an increase in ADP concentration (ADP activates PPDK regulatory protein 1, as indicated by the green arrow). The inactivity of PPDK phosphorylated on Thr 543 is represented in two ways: Firstly, a comment in the gray oval below the protein gives this information. Secondly, the reaction catalyzed (pyruvate -> PEP) is not connected to the Thr543-phosphorylated PPDK protein. Other examples of control by phosphorylation can be found in ChloroKB (see for example the following maps: “mitochondrial pyruvate dehydrogenase regulation,” “choline-phosphate cytidylyltransferase 1 regulation,” “sucrose-phosphate synthase regulation”). 3.4 Representation of Multilayered Regulation

1. Several examples of multilayered regulation can be found in ChloroKB: A relatively simple one concerns ADP-glucose pyrophosphorylases (Fig. 5), catalyzing the first step committing sugar to starch synthesis. 2. The representation principle is the same as described above for the pyruvate kinases: a first map presents the ADP-glucose pyrophosphorylase enzymes in the cellular context of the plant (in this case, the plastid, Fig. 5a), and a second map details the regulatory mechanisms (Fig. 5b). 3. To access the map describing ADP-glucose pyrophosphorylase enzymes from the home page, users can either write “ADPglucose pyrophosphorylases” in the search field or use the CTRL+F shortcut: in that case a map icon is highlighted in yellow on the home page, and clicking on it will show the map

ä Fig. 4 (continued) reaction release AMP, leaving a phosphate group on His545. The cycle is completed with the transfer of this phosphate group onto pyruvate to form PEP. (b) PPDK regulation by phosphorylation: plastidial and cytosolic PPDKs are regulated by two different PPDK bifunctional regulatory proteins (kinases and phosphatases), PPDK regulatory protein 1 and 2. Phosphorylation of PPDK on Thr543 is unusual as it involves ADP and not ATP. Phosphorylation leads to complete inhibition of PPDK1 (red dotted box, no link between this protein and the reaction catalyzed). Dephosphorylation reactivates the enzyme. Regulatory phosphorylation is stimulated by high ADP levels in the plastid (i.e., at night, see the green arrow on the website, highlighted here by a star)

a

zoom

Click

Oxidizing Condition

Reducing Conditions

Light Reaction catalyzed

Reaction catalyzed

Oxidized, Very low activity zoom

Oxidized Cys81-Cy81 bridge

b 3PGA bound, Highly active enzyme

Dark

Free enzyme moderately active

Pi-bound enzyme Not competent for 3PGA binding

Fig. 5 Multilayered representation of regulatory mechanisms in ChloroKB (ADP-pyrophosphorylase enzyme example). (a) The background picture is a screen capture of the “ADP-glucose pyrophosphorylases” ChloroKB map. It shows the four different complexes found in A. thaliana: within the leaf plastids on the left (proteins are colored orange as they are localized in the stroma) and in the plastids of sink tissue on the right (pale green color means their precise localization inside sink tissue plastids is unknown). ADP-pyrophosphorylase enzymes are heterotetramers formed of a dimer of small subunits (APS1) and a dimer of large subunits (APL1-4). In the APS1-APL3 and APS1-APL4 heteromers, only the APS1 dimer is catalytically active: this is represented by the link joining the complex to the reaction starting from the APS1 dimer and not from the entire APS-APL complex, in contrast to what is shown for the APS1-APL1 and APS1-APL2 heteromers (highlighted by red circles). (b) ADP-pyrophosphorylase regulation map (leaf plastid enzyme). Boxes superimposed on the chloroKB screen capture highlight the allosteric and redox control of the enzyme: Enzyme activity is activated by 3-phosphoglycerate (3PGA, green box, top right) and inhibited by phosphate (phosphate actually prevents 3PGA from binding and activating the enzyme, but does not affect enzyme activity in the absence of 3PGA). The differences in activity of the distinct complexes are symbolized by the thickness of the link between the complex and the reaction catalyzed, as in Fig. 4. Left part of the map represents the redox regulation of the enzymatic activity (inhibition, red box, left) with the formation of an intermolecular Cys bridge between two APS1 subunits (see zoom on the bottom left). Oxidation/reduction of the Cys81 pair is catalyzed by thioredoxins and NADPH thioredoxin reductase (central part). Clicking on the red dots (black arrows) provides specific functional information on complexes

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displayed in the background in Fig. 5a. In A. thaliana, four different complexes catalyze the activation of glucose 1 phosphate to produce PPi and ADP-α-D-glucose, the unique precursor of starch. The ADP-glucose pyrophosphorylase complexes are heteromers formed of homodimers of two small (APS) and two large (APL) subunits. These polypeptides are catalytically active when complexed with a magnesium ion. The complex on the left ([(APS1)2-APL1)2)], Fig. 4a) is the protein complex identified by proteomics in the leaf chloroplast. 4. The proteins are shown in orange because they were detected in the stroma fraction of isolated chloroplasts [13]. (Click on “legend” on the right of the page for more information on the color code used.) The protein complexes on the right in Fig. 5a contain the same small subunit (APS1) as the complex represented on the left, but the large subunit is different. The large subunits are coded by different genes (APL2, APL3, APL4). Their color is pale green because they have never been detected in leaf (the cDNA of these proteins are produced in sink tissues, as indicated in the gray ovals close to the complexes). Whether APL2-5 localizes inside non-photosynthetic plastids is currently unknown (hence the pale green color). Although these proteins are most probably present in the stroma, this coding allows the available experimental data or lack of data to be fully traced. 5. The usefulness of the CellDesigner graphical code for the precise economical representation of biochemical information is demonstrated by the following details: note that the connector between the complexes and the reaction differs for the APS1-APL1 and APS1-APL2 heterotetramers compared to the APS1-APL3 and APS2-APL4 heterotetramers (this difference is highlighted by the red circles in Fig. 5a). The reason for the difference is that APS1 and not the APL subunit is active in the latter two complexes (the connector starts from APS1 and not from the entire complex). The large subunits APL3 and APL4 only have regulatory properties. 6. From the map in Fig. 5a, more detailed information on the regulation of the plastidic enzyme can be accessed by clicking on the purple hexagon “ADP-glucose pyrophosphorylase regulation” (Fig. 5b). Regulation of the [(APL1)2-(APS1)2] leaf enzyme is represented, because more data are available for this complex in Arabidopsis (see Note 5). Figure 5b shows that the activation of the APS1-APL1 hetero-tetramer is regulated mutually by exclusive binding of 3-phosphoglycerate (3-PGA, activator) and inorganic phosphate (Pi, inhibitor). This is

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represented vertically on the right of the map. The free enzyme is moderately active; however, when bound to the Calvin cycle intermediate 3-phosphoglycerate, the APS1-APL1 heterotetramer activity is strongly stimulated. The enzyme complexed with phosphate displays the same activity as the free enzyme (same thickness of the catalysis link joining the enzyme-Pi complex (bottom right in Fig. 5b) and the free enzyme (middle)). Indeed, phosphate only inhibits enzymatic activity in the presence of 3-phosphoglycerate (both bind to the same site in the protein). In molecular terms, the simplest interpretation is that phosphate populates a moderately active Pi-enzyme complex that cannot bind 3-phosphoglycerate. Thus, at a given concentration of 3-phosphoglycerate, addition of phosphate will inhibit enzymatic activity by depleting the most active population of the enzyme complexed with 3PGA. 7. In addition to allosteric regulation, APS1-APL1 heteromer activity is also altered by reversible oxidation/reduction of the APS1 subunit of the protein complex. This is represented in the map on the horizontal axis (Fig. 5b): the oxidized, low-activity form is displayed on the left, while the active form—competent to bind the allosteric effectors—is displayed on the right. The reversible oxidation/reduction of ADP-glucose pyrophosphorylase APS1 subunit is catalyzed by thioredoxins f1 and m1 and the NADPH-dependent protein NTRC (central part of the map). Under oxidizing conditions (dark), two cysteines (Cys81) in the APS1 dimer form a covalent intermolecular disulfide bridge. This crosslinking reduces the enzyme’s activity. The representation of the chemical crosslink between the two APS1 proteins is not entirely satisfactory in ChloroKB (see zoom in Fig. 5b). A better representation would show the two cysteine 81 residues facing each other, with a link in between. Unfortunately, cysteine crosslinking modification is not (yet) available in the CellDesigner formalism. In addition, for technical reasons, the conversion of CellDesigner files into svg files for display on the website prevents the positioning of the two proteins with the oxidized cysteines facing each other. Future developments may solve this issue. Other examples are available in ChloroKB: see for example the redox control of the protein cp12 in the “GAPDH PRK redox” or “fructose 1 6 bisphosphatase 1” maps. See Note 6.

4

Notes 1. In the present context, the term “control” (of protein activity), used as a synonym of “modification,” would be preferable to “regulation.” Indeed, “regulation” is an integrated function,

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involving different control processes, the function of which is to stabilize a system’s variability. However, the term “regulation” is now widely used, especially in the protein structure community, and we will therefore comply with this usage. 2. Please note that the proteins displayed in the maps are not necessarily produced simultaneously in the cell: Maps in ChloroKB are representations of a “generic” cell with all the proteins known to contribute to the process in the plant. Further developments will deconvolute these representations to account for expression patterns in different cell types. 3. See color coding legend in the upper corner of the website and reference [11]. 4. No symbol for covalent links between two proteins or between a protein and a metabolite is available in CellDesigner, but it has been added in ChloroKB (CellDesigner pictures are redrawn from the CellDesigner xml code to display them on the website as svg pictures). 5. The regulatory mechanism is the same for the “sink-specific” enzymes (i.e., [(APL2)2-(APS1)2], [(APL3)2-(APS1)2], and [(APL3)2-(APS1)2]) with quantitative differences in the apparent affinities. These will be represented when more detailed information is published. 6. With this introduction to the representation of complicated (but not necessarily complex) biomolecular interactions in ChloroKB, we would like to convey the idea that “a picture is worth a thousand words” ([10]), especially when using standard graphical notations. Graphical representations are very helpful as they provide synoptic views of integrated biological information, allowing rapid comprehension and inter-human communication. We would like to point out that the reconstructions of the molecular interactions in ChloroKB require interpretation: each molecular model represents the synthesis of heterogeneous data from different publications ranging over many years. These representations are not necessarily stabilized and may be improved as new data is published, or following discussions with experts. In this context, feedback from the scientific community is very welcome to improve the resource that is ChloroKB. From the collection of regulatory processes described in ChloroKB, it will be easier to derive phenomenological or mechanistic mathematical equations. Indeed, to improve our understanding of the dynamics of metabolic systems, regulatory features must be included in mathematical models.

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References 1. Kebeish R, Niessen M, Thiruveedhi K et al (2007) Chloroplastic photorespiratory bypass increases photosynthesis and biomass production in Arabidopsis thaliana. Nat Biotechnol 25:593–599 2. Maier A, Fahnenstich H, von Caemmerer S et al (2012) Transgenic introduction of a glycolate oxidative cycle into A-thaliana chloroplasts leads to growth improvement. Front Plant Sci 3:38 3. South PF, Cavanagh AP, Liu HW et al (2019) Synthetic glycolate metabolism pathways stimulate crop growth and productivity in the field. Science 363(6422):eaat9077 4. Roell MS, von Borzyskowski LS, Westhoff P et al (2021) A synthetic C4 shuttle via the betahydroxyaspartate cycle in C3 plants. Proc Natl Acad Sci U S A 118(21):e2022307118 5. Blatke MA, Brautigam A (2019) Evolution of C4 photosynthesis predicted by constraintbased modelling. Elife 4(8):e49305 6. Shameer S, Ratcliffe RG, Sweetlove LJ (2019) Leaf energy balance requires mitochondrial respiration and export of chloroplast NADPH in the light. Plant Physiol 180:1947–1961 7. Kacser H (1963) The kinetic structure of organisms in biological organization at the

cellular and supercellular level (Harris RJC, ed). Academic, New York/London, pp 25–41 8. Farre G, Twyman RM, Christou P et al (2015) Knowledge-driven approaches for engineering complex metabolic pathways in plants. Curr Opin Biotechnol 32:54–60 9. Funahashi A, Matsuoka Y, Jouraku A et al (2008) CellDesigner 3.5: a versatile modeling tool for biochemical networks. Proc IEEE 96: 1254–1265 10. Le Novere N, Hucka M, Mi HY et al (2009) The systems biology graphical notation. Nat Biotechnol 27:735–741 11. Gloaguen P, Bournais S, Alban C et al (2017) ChloroKB: a web application for the integration of knowledge related to chloroplast metabolic network. Plant Physiol 174:922–934 12. Gloaguen P, Vandenbrouck Y, Joyard J et al (2021) ChloroKB, a cell metabolism reconstruction of the model plant Arabidopsis thaliana. C R Biol 344:157–163 13. Ferro M, Brugiere S, Salvi D et al (2010) AT_CHLORO, a comprehensive chloroplast proteome database with subplastidial localization and curated information on envelope proteins. Mol Cell Proteomics 9:1063–1084

INDEX A

D

Alpha-proteo-bacteria ...............................................31, 89 Amyloplast ................................................... 66, 72, 74, 91 Antimicrobial peptide ..................................................... 13 Apicomplexa .................................................10, 31, 43–60 Apicoplast ................................... 34–36, 43–60, 198, 201 Arabidopsis.................................................. 14, 75, 77, 91, 92, 95, 101, 137–139, 143, 145, 152, 162–164, 166, 174, 213, 214, 229, 237, 259, 263, 264, 291–298, 300, 307, 308, 310–312, 317 Archaea .............................................................7, 9, 11, 13

Database ................................................92, 275, 285, 307 Diatom..................................................22, 27, 28, 33, 36, 122, 177–183, 185–195, 270, 273 DNA .................................................. 7, 9, 13, 17, 30, 44, 48, 64, 65, 67, 110, 188–194, 243, 245, 247, 250, 252, 254–256, 263, 270–273, 275, 278, 281, 283, 285, 286

B Bacteria ....................................................... 3, 7, 9, 13–15, 17, 44, 52, 174, 260, 272, 274, 277, 281, 282, 286, 290 Biogenesis ................................................. 4–7, 10, 17, 54, 59, 81, 107, 110, 121, 137, 151, 161, 197, 198, 201, 290 Bioinformatics .................................................44, 47, 244, 245, 259–264, 276

C Carbon metabolism......................................................... 35 Cell compartments..................................... 93, 95, 98, 99, 102, 107–126, 163 Cell fractionation .............................................................. 4 Chlamydia........................................................... 14, 15, 17 ChloroKB ............................................................. 305–319 Chlorophyll ................................................ 14, 15, 26, 28, 33, 52, 65–67, 69, 71–73, 78, 79, 96, 97, 109, 146, 148, 158, 240 Chlorophyta .................................................................... 14 Chloroplast ................................................... 3, 29, 44, 63, 90, 108, 137, 152, 177, 185, 206, 231, 243, 259, 289, 317 Chromatophore........................................................... 3–17 Chromoplast......................................... 63, 69, 70, 72–76, 81, 90, 95, 97 CRISPR/Cas9...................................................... 269–286 Cyanobacteria ................................................3–17, 22, 29, 31, 93, 97, 231

E Endomembrane system.....................................6, 7, 9, 10, 26, 29, 30, 36 Envelope membranes ............................. 9, 13, 30, 65–67, 69, 91, 94, 96–98, 102, 138, 145, 148, 151, 152, 155–157, 185, 187, 290 Etioplast........................................................66–73, 76, 90 Eukarya ...................................................... 7, 9, 11, 13, 14

G Gas chromatography (GC) ................................. 198, 206, 207, 232, 233 Genome editing ............................................................ 270 Genome size reduction ................................................... 64 Green algae ................................................. 14, 16, 22, 24, 25, 34, 63, 64, 122, 290

H Horizontal gene transfer (HGT) ................ 9, 11, 14, 15, 17, 31, 32

L Lateral gene transfer ....................................................... 14 LC-MS/MS.......................................................... 206, 227 Leaf ............................................................. 68, 70, 72, 73, 76–79, 90, 93, 94, 110, 123, 137–148, 152, 153, 155, 157, 158, 164, 245, 247, 291, 292, 295, 298, 300, 301, 316, 317 Lipidome .................................... 118, 206, 212, 231–241 Lipidomics ....................................................173, 198–203 Lipids galactolipids ................................................. 10, 65, 97, 117–119, 231, 232

Eric Mare´chal (ed.), Plastids: Methods and Protocols, Methods in Molecular Biology, vol. 2776, https://doi.org/10.1007/978-1-0716-3726-5, © The Editor(s) (if applicable) and The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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Lipids (cont.) phosphatidylglycerol ...............................5, 67, 96, 97, 113, 114, 229, 231, 238, 239 sulfolipid ...................................................65, 113, 231

M Mitochondria................................................ 9, 30, 51, 64, 89, 109, 161, 182, 193, 264, 289, 311 Mutant ...................................................... 34, 56, 60, 110, 112, 116, 118, 119, 121, 125, 164, 244, 269, 270, 283–285

P Pea................................................................. 72, 138, 141, 145, 146, 152, 156, 157 Percoll gradient ..................................139, 141–143, 146, 153, 155, 162, 181, 182, 295 Phaeodactylum ...................................... 27, 122, 177–183, 186, 229, 269–286 Phenotype..................................................... 48, 111, 118, 119, 125, 244, 283–285 Photosynthesis................................... 3, 5, 21, 24, 25, 28, 30, 31, 33–36, 64–66, 68, 69, 73, 74, 80, 91, 93–94, 111, 118, 121–123, 137, 162, 185, 186, 289, 305 Phytoplankton ...................................................... 178, 205 Pigments ........................................ 14, 51, 65–67, 71, 73, 75, 78, 93, 95–96, 289, 307 Plants ....................................................4, 56, 64, 89, 107, 137, 151, 162, 185, 205, 231, 290, 305 Plasmodium .......................................................28, 34, 36, 43–46, 48, 51–54, 56, 58, 59 Primary endosymbiosis ......................... 11, 14–17, 22, 64 Primary plastid ..................................................14, 17, 22, 26, 29, 47, 64, 185 Proplastid..................................65, 70, 72–75, 78, 79, 90 Protein import..........................29, 36, 53, 123, 125, 187 Proteome .................................................... 28, 29, 31, 47, 90–93, 100, 101, 152 Proteomics..........................................56, 91–93, 96, 101, 163, 171, 290, 300, 317 Protist ...............................................................43, 49, 197

R Red algae ..................................14, 16, 22, 25, 29, 43, 64 Rhizaria ......................................................................15–17 Rhodophyta ..................................................................... 14 RNA ........................................................7, 9, 44, 48, 189, 190, 194, 244, 245, 247–250, 259–264, 270, 290 Root ................................................ 70, 72–75, 78, 79, 90

S Secondary endosymbiosis ................................... 4, 22, 32, 36, 43, 46, 64 Secondary plastid ......................................... 7, 10, 36, 49, 57, 177, 185–195 Spinach....................................... 114, 137–139, 141–143, 146, 148, 152, 155, 158 Starch ............................................. 66, 69, 72, 74, 81, 91, 93, 94, 100, 101, 138, 146, 157, 315, 317 Stramenopile.................................................................... 24 Stroma................................................... 13, 49, 53, 65–70, 73, 77, 78, 91, 92, 94, 96, 109, 137–148, 195, 206, 313–317 Sucrose gradient ................................. 138–141, 143–148, 152, 154, 156–158

T Thylakoids .............................................. 3–17, 65, 67–69, 72–79, 91, 92, 96, 114, 118, 137–148, 231 Toxoplasma............................................... 28, 44, 197, 198 Transcriptome ...................................................... 243–256 Transit peptide (TP) ..................23, 29, 30, 49, 186, 290 Translocon of the inner chloroplast membrane (TIC)................................................ 13, 23, 30, 49 Translocon of the outer chloroplast membrane (TOC) .............................................. 13, 23, 30, 49 Transporters ............................................... 30, 31, 36, 45, 52, 54–59, 74, 93–95, 98, 99, 101, 102, 116–118, 306 Triose phosphate ...............................................30, 35, 44, 54, 55, 58, 59, 122