Biology of Algae, Lichens and Bryophytes 9783662657119, 9783662657126

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Burkhard Büdel · Thomas Friedl · Wolfram Beyschlag Editors

Biology of Algae, Lichens and Bryophytes

Biology of Algae, Lichens and Bryophytes

Burkhard Büdel • Thomas Friedl Wolfram Beyschlag

•

Editors

Biology of Algae, Lichens and Bryophytes

Editors Burkhard Büdel Department of Biology RPTU Kaiserslautern Kaiserslautern, Rheinland-Pfalz Germany

Thomas Friedl Department of Experimental Phycology and Culture Collection of Algae (EPSAG) Georg August University Göttingen Göttingen, Germany

Wolfram Beyschlag Department of Experimental and Systems Ecology University of Bielefeld Bielefeld, Nordrhein-Westfalen Germany

ISBN 978-3-662-65711-9 ISBN 978-3-662-65712-6 https://doi.org/10.1007/978-3-662-65712-6

(eBook)

© Der/die Herausgeber bzw. der/die Autor(en), exklusiv lizenziert durch Springer-Verlag GmbH, DE, ein Teil von 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. Grafiken: Dr. Frederik Spindler This Springer Spektrum imprint is published by the registered company Springer-Verlag GmbH, DE, part of Springer Nature. The registered company address is: Heidelberger Platz 3, 14197 Berlin, Germany Paper in this product is recyclable.

Prolog

Writing a new textbook about algae, lichens, and bryophytes is a challenge. We would not have been able to write that book without our coauthors Tatyana Darienko (University of Göttingen, Germany), Olivier De Clerck (University of Gent, Belgium), Jan de Vries (University of Göttingen, Germany), Louis Graf (Sungkyunkwan University, Suwon, Korea), Kerstin Hoef-Emden (University of Cologne, Germany), Mona Hoppenrath (Senckenberg Institute, Wilhelmshaven, Germany), Frederik Leliaert (Meise Botanic Garden, Belgium), and Angelika Preisfeld (University of Wuppertal, Germany). Special thanks go to the excellent scientific artist Frederick Spindler. He understood perfectly how to adapt to our individual working style, and the resulting Illustrations are wonderful. The English of many, but not all, chapters was considerably improved by Laura Briegel-Williams, many thanks for that Laura. Without the critical comments from our colleagues, that were highly appreciated and valuable, the book might have missed some important topics. These colleagues are Wolf-Rüdiger Arendholz (University of Kaiserslautern, Germany), T. G. Allan Green (University of Waikato, Hamilton, New Zealand), Martin Hagemann (University of Rostock, Germany), Thorsten Lumbsch (Field Museum Chicago, USA), and Patrick Jung (University of Applied Sciences, Kaiserslautern, Germany). We are deeply grateful to our friends, colleagues, and many more people for providing us with their often-magnificent pictures and micrographs of the organisms and their environments included in this book. We can only mention a few who provided us with either a number of pictures or pictures from difficult environments. So thanks go to Robert A. Andersen, Siegmar-Walter Breckle, Laura Briegel-Williams, Scott Camazine, Claudia Colesie, Jan Eckstein, Lars and Christina Frank, Michael Geyer, Janice Glime, Stjepko Golubic, Gerd Günther, Antje Gutowski, Alan S. Heilmann, Patrick Jung, Heike Hofmann, Lothar Krienitz, Christian Kleinert, Michael Lüth, Tony Markham, Mark Milburn, Manuel Müller, Walter Obermayer, Joachim Reitner, Karen Renzaglia, Klaus Rudlof, Nataliya Rybalka, Hermann Schachner, Michael Schagerl, Norbert Schnyder, Rhena Schumann, Sergei Shalygin, Norbert J. Staper, Ralf Wagner, Brian Whitton, Heike Wägele, David H. Wagner, Solvin Zankl, Jonas Zimmermann, and all the others whose names are given in the figure legends. The manuscript for this book was prepared over a period of four years. When we started our project, it was commonly thought that pandemia and war in Europe were threats that human society has already overcome since many decades. We believe that in preparing this textbook, we have dealt with providing convincing evidence that success in evolution leading to diversity and adaptations to rather diverse habitats is only possible by close cooperation and integration leading to mutual benefits. The close interaction of organisms of rather different ancestries has led to the origin of new evolutionary lineages with broad offspring and complex, well-adapted symbiotic communities, which would not have been possible when being separated from each other or attacked by aggression.

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Prolog

Working on the book manuscript was often a challenging task for our families. Despite many occasions of absences, either physically or mentally, they took it with patience and sympathy. We are deeply grateful to you. You are our stronghold. Finally, we would like to thank Stefanie Wolf and Martina Mechler from Springer-Nature for the careful preparation of this book. We dedicate this book to our deceased academic mentors and later friends Prof. Dr. Dr. h.c. mult. Otto Ludwig Lange (Julius Maximilians University, Würzburg) and Prof. Dr. Dieter Mollenhauer (Senckenberg Institute, Frankfurt, Germany). Both of them have served as an inspiration to our studies of pro- and eukaryotic algae, lichens, and bryophytes, their phylogeny, systematics, and ecology. Kaiserslautern, Germany Göttingen, Germany Bielefeld, Germany February 2023

Burkhard Büdel Thomas Friedl Wolfram Beyschlag

Contents

1 Introduction: The New Cryptogams . . . . . . . . . . . . . . . . . . . . . . . . . . . . Burkhard Büdel and Thomas Friedl 1.1 Oxy-Phototroph Life Forms and Traditional Botany . . . . . . . . . . . . . . 1.2 The New Cryptogams—a Modern Concept for the Non-Vascular Oxy-Phototroph Life Forms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.3 Functional Aspects of New Cryptogams: Hydro-Passivity . . . . . . . . . 1.4 The New Cryptogams Predated the Conquest of the Third Dimension on Land . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.5 The New Cryptogams and Chapter Selection of This Textbook . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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2 Endosymbioses: Origin and Diversity of Photosynthetic Eukaryotes and Their General Genetic Exchange Modes . . . . . . . . . . . . . . . . . . . . . . . Jan de Vries and Thomas Friedl 2.1 Introduction: The Role of Endosymbiosis in the Evolution of Eukaryotes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2 Primary Endosymbiosis: The Endosymbiotic Origin of Plastids . . . . . . . 2.3 Eukaryote-Eukaryote Endosymbioses . . . . . . . . . . . . . . . . . . . . . . . . . . 2.4 A Modern Overview of Eukaryotic Diversity—The Opuntia-Tree-of-Life 2.5 Life histories, Plastid Spread, and Sexual Reproduction in the New Cryptogams . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Cyanobacteria/Blue-Green Algae . . . . . . . . . Burkhard Büdel 3.1 Short History of Cyanobacterial Research 3.2 Origin of Cyanobacteria . . . . . . . . . . . . . 3.3 Structure and Function . . . . . . . . . . . . . . 3.4 Survival and Cell Specialization . . . . . . . 3.5 Reproduction, Life Cycle . . . . . . . . . . . . 3.6 Phylogeny and Diversity . . . . . . . . . . . . 3.7 Ecology . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . .

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4 Algae from Primary Endosymbioses . . . . . . . . . . . . . . . . . Burkhard Büdel and Thomas Friedl 4.1 Rhodophyta, Red Algae . . . . . . . . . . . . . . . . . . . . . . . . Burkhard Büdel 4.2 Chloroplastida—Green Algae . . . . . . . . . . . . . . . . . . . . Frederik Leliaert, Olivier De Clerck, Tatyana Darienko, Jan de Vries, and Thomas Friedl 4.3 Glaucophyta . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Burkhard Büdel

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Contents

4.4 Cercozoa—A Second Primary Endosymbiosis . . . . . . . . . . . . . . . . . . . . . . . 203 Burkhard Büdel References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 207 5 Algae from Secondary Endosymbiosis . . . . . . . . . . . . . . . . . . . . Thomas Friedl 5.1 Heterokontophyta—Photosynthetic Stramenopiles . . . . . . . . . Louis Graf 5.2 Dinoflagellates . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Thomas Friedl and Mona Hoppenrath 5.3 A Cercozoan Secondary Endosymbiosis: Chlorarachniophyta . Angelika Preisfeld and Burkhard Büdel 5.4 Euglenids—(Excavates, Discoba, Euglenozoa, and Euglenida) Angelika Preisfeld 5.5 Haptista . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Burkhard Büdel and Angelika Preisfeld 5.6 Cryptista . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Kerstin Hoef-Emden References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6 Symbioses . . . . . . . . . Burkhard Büdel 6.1 Algal Symbioses 6.2 Lichens . . . . . . . References . . . . . . . . .

. . . . . . . . . . . 219 . . . . . . . . . . . 220 . . . . . . . . . . . 280 . . . . . . . . . . . 297 . . . . . . . . . . . 302 . . . . . . . . . . . 323 . . . . . . . . . . . 333 . . . . . . . . . . . 353

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7 Bryophytes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Wolfram Beyschlag 7.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . 7.2 Division: Marchantiophyta—The Liverworts . 7.3 Division Bryophyta—The Mosses . . . . . . . . . 7.4 Division: Anthocerotophyta—The Hornworts . 7.5 Asexual Reproduction . . . . . . . . . . . . . . . . . 7.6 Physiology and Physiological Ecology . . . . . 7.7 Ecology . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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476 478 495 532 538 540 554 580

Glossary . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 605 Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 615

1

Introduction: The New Cryptogams Burkhard Büdel and Thomas Friedl

Contents

1.1

1.1 Oxy-Phototroph Life Forms and Traditional Botany .....................................................................

1

1.2 The New Cryptogams—a Modern Concept for the Non-Vascular Oxy-Phototroph Life Forms ....................................................................................................................................................

2

1.3 Functional Aspects of New Cryptogams: Hydro-Passivity ..............................................................

6

1.4 The New Cryptogams Predated the Conquest of the Third Dimension on Land

7

1.5 The New Cryptogams and Chapter Selection of This Textbook ....................................................

7

References .....................................................................................................................................................

8

Oxy-Phototroph Life Forms and Traditional Botany

This textbook concerns a large fraction of the tree of life, the non-vascular oxygenic photoautotrophs (oxy-phototrophs). They comprise algae, cyanobacteria, lichens, and bryophytes. They are at the borderline of micro- to macroorganisms and originated very early in evolution. Also, their number is a plethora: estimates suggest roughly 290,000 species of algae (incl. cyanobacteria), lichens, and bryophytes (Table 1.1). Considering that the estimated number of existing species of vascular plants, which comprise ferns, gymnosperms, and angiosperms, is about 435,000, there is a relation of 40% non-vascular to 60% vascular oxy-phototrophs (Table 1.1). However, the global impact of the non-vascular oxy-phototrophs on the net biological B. Büdel (&) RPTU Kaiserslautern, Department of Biology, Erwin-Schrödinger-Str. 13, 67663 Kaiserslautern, Germany e-mail: [email protected] T. Friedl Department of Experimental Phycology and Culture Collection of Algae (EPSAG), Georg August University Göttingen, Nikolausberger Weg 18, 37073 Göttingen, Germany e-mail: [email protected]

oxygen production is more than 50% of the Earth’s annual oxygen release, of which 50–80% comes from marine (https://oceanservice.noaa.gov/facts/ocean-oxygen.html) and 10 -15% from terrestrial habitats. In terrestrial habitats, algae in soils and biological soil crusts as ground and plant covers form the dominant oxy-phototrophs (Elbert et al. 2012; Jassey et al. 2022). Consequently, the non-vascular oxy-phototrophs are of enormous ecological importance as primary producers, occurring in almost all habitats where sufficient light is available to fix carbon by photosynthesis from atmospheric CO2. Their provision of organically bound carbon fuels terrestrial habitats with energy, an essential prerequisite to enable other forms of life. Traditional botany has provided a clear separation of the oxy-phototroph organisms into two artificial categories. The structurally simpler photoautotrophic life forms, i.e., those not reaching the complex structural design of seed-forming plants, have collectively been summarized as “cryptogams” as opposed to the phanerogams, which encompass only the seed-forming plants. The traditional cryptogams have even included prokaryotic life forms, i.e., the photoautotrophic cyanobacteria. However, also non-photosynthetic and structurally simpler forms, i.e., the fungi and slime molds, have traditionally been included in the cryptogams as well. For example, Karls Esser's popular textbook “Kryptogamen. Praktikum und Lehrbuch” (Cryptogams. Practical and

© Der/die Autor(en), exklusiv lizenziert durch Springer-Verlag GmbH, DE, ein Teil von Springer Nature 2024 B. Büdel et al. (eds.), Biology of Algae, Lichens and Bryophytes, https://doi.org/10.1007/978-3-662-65712-6_1

1

2 Table 1.1 Known and estimated numbers of species of oxy-phototroph life forms on Earth

B. Büdel and T. Friedl

New Cryptogams (non-vascular oxy-phototrophs)

Vascular oxy-phototrophs

No. described species

No. estimated species

Source

Algae (including cyanobacteria)

50,900

250,000

Kamiya et al. (2017), Hofbauer et al. (2015), Guiry 2024, Wang et al. (2022)

Lichens

17,600

18,600

Frey et al. (2009); this book, Sect. 6.2 Frey et al. (2009), http:// www.theplantlist.org/ browse/B/

Bryophytes

17,900

20,000

Total

86,400

288,600

19%

40%

Ferns

11,020

13,000

Frey et al. (2009), http:// www.theplantlist.org/ browse/P/

Gymnosperms

1,040

1,100

Fischer et al. (2015)

Angiosperms

352,000

422,100

Total

364,060

435,100

Fischer et al. (2015), http:// www.theplantlist.org/ browse/A/

81%

60%

textbook) discussed the prokaryotic cyanobacteria (or blue-green algae) as “Schizophyta” in the 1st and 2nd edition, the eukaryotic algae as “Phycophyta” in the 1st edition. The various groups of heterotrophic fungi, Oomycetes, slime molds, and symbiotic lichens were all treated under “Mycophyta”. However, those diverse groups are non-related to each other (see Figs. 1.1, 2.8). All those non-vascular plants which are reproducing by spores (see Sect. 2.5), i.e., mosses, liverworts, and hornworts (the bryophytes, i.e., Bryophyta, Marchantiophyta, and Anthocerophyta; Figs. 1.1, 2.8; see Chap. 7) were together with the vascular non-seeds producing lycophytes and ferns all collectively considered as “Cryptogams” as well (Esser 1992, 2000). Vascular tissue, a typical feature of seed plants that enables them to grow to large sizes and up to several meters above the ground, visually dominating most terrestrial habitats, is also shared by ferns and lycophytes. The traditional concept of cryptogams was developed before the enormous phylogenetic diversity of the oxy-phototroph life forms was recognized. From today’s view, i.e., with a synthesis of all information available from phylogenetics, phylogenomics, and cell biology, the traditional cryptogams comprise such an extremely heterogeneous assemblage that using the term makes no sense as a systematic category to describe the oxy-phototroph life forms. Having cryptogams as opposed to tracheophytes/ phanerogams now would mean an unprecedented diversity of life forms, including the fungi, being opposed to a single monophyletic autotrophic lineage, the Tracheophyta (sensu Adl et al. 2019; Figs. 1.1, 2.8). The Tracheophyta is a well-supported monophyletic lineage that comprises all

vascular plants, i.e., ferns, lycophytes, and the phanerogams, which consist of gymnosperms and angiosperms (One Thousand Plant Transcriptomes Initiative 2019).

1.2

The New Cryptogams—a Modern Concept for the Non-Vascular Oxy-Phototroph Life Forms

There is a need for a handsome term that describes all oxy-phototroph (photoautotroph) life forms that do not develop vascular tissues and considers their multiple origins in evolution. Firstly, there is a complex reticulate evolution that governed the origins of the oxy-phototroph life forms and unified them like a network, contrasting bifurcating tree-like evolution. This requires already a new graphical display of the current view of their origins which we show in the “Opuntiatree-of-life” in Figs. 1.1 and 2.8. The oxy-phototroph (photoautotroph) lifestyle originated in cyanobacteria. After that, it spread over many independent lineages of the eukaryotes, leading to the various types of plastids in the algae (see Chaps. 2, 4, and 5). Algae are defined here as all organisms permanently capable of oxygenic photosynthesis and not developing embryos, unlike bryophytes, ferns, lycophytes, and seed plants (Melkonian et al. 1995). In addition to plastid origins, the oxy-phototroph lifestyle has also been developed in various symbioses. Examples of intracellular (endosymbiotic) and extracellular (exosymbiotic) symbioses are in the fungi, such as the enigmatic Geosiphon (a unique endosymbiosis of a glomeromycete with the cyanobacterium

Introduction: The New Cryptogams

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Fig. 1.1 Opuntia-tree-of-life and the New Cryptogams within the diversity of eukaryotes. The New Cryptogams comprise cyanobacteria, eukaryotic algae, bryophytes, and lichens (see text). Clubbed presentation of groups (“leaves of the Opuntia”): green, photosynthetic; gray, primarily non-photosynthetic. The multiple endosymbiotic origins of plastids connect the single groups through primary or secondary endosymbiosis (curved arrows). Lichens are symbiotic associations of members of Ascomycota or Basidiomycota with Cyanobacteria, Xanthophyceae, or Chlorophyta that develop multicellular and structurally unique forms bound permanently to photosynthesis. Inset, Opuntia echios (by “Haplochromis,” from Wikipedia). Names of the various groups of eukaryotes follow Adl et al. (2019). Original drawing Spindler & Friedl

4

Nostoc, Figs. 1.1, 2.8; see Sect. 6.1) and the lichens which are diverse associations of members of Ascomycota and Basidiomycota with cyanobacteria and/or algae (Figs. 1.1, 2.8; see Sect. 6.2). The corals, being of enormous ecological importance (see Chap. 6.1), are further compelling examples of photosymbiosis, i.e., of marine invertebrate animals (Holozoa, Anthozoa) with several unrelated lineages of algae (Figs. 1.1, 2.8). Secondly, many lineages of diverse oxy-phototroph (photoautotroph) life forms dominated the terrestrial vegetation for a very long time in Earth's history until the advent of vascular plants about 300 million years ago (=Mya; Fig. 1.2). This is, thirdly, intimately connected with functional aspects of exceptional importance for the ecology and evolutionary success of plant life on the Earth's surface. The mode of water uptake and loss unifies all those diverse oxy-phototroph life forms of cyanobacteria, algae, bryophytes, and photosymbioses such as lichens. They all lack vascular tissue in their vegetative life forms; hence, there is no active spread of photosynthetic products and no controlled uptake or loss of water, but environmental conditions govern both. While being so diverse, those oxy-phototroph life forms are unified by featuring a hydro-passive lifestyle. Consequently, considering the evolutionary, historical, and functional aspects, we propose to renew the term cryptogams in a different sense and introduce the term “New Cryptogams”. The New Cryptogams comprise all oxy-phototroph (photoautotrophic) life forms whose capability of performing oxygenic photosynthesis is permanent and who, at the same time, feature a hydro-passive lifestyle. Hydro-passivity includes dependence on water to keep the shape of bodies or cells while exhibiting the full range from desiccation sensitivity to desiccation tolerance. The members of the New Cryptogams originated through a preceding reticulate evolution pattern triggered by several endosymbiotic events (see Sects. 2.1–2.4). The New Cryptogams also include those symbiotic associations which develop unique forms specific to the symbiosis. The New Cryptogams include the lichens (Ascomycota, Basidiomycota; Figs. 1.1, 2.8) because their multicellular and structurally unique forms are hydro-passive and bound permanently to photosynthesis. The lichens’ specific morphology is being formed in symbiosis exclusively; neither the dikaryotic fungi alone nor the cyanobacteria or eukaryotic algae can develop such specific unique forms (Figs. 1.1, 2.8; see Sect. 6.2). In contrast, no unique forms and structures are formed neither by the cyanobacterial endosymbiosis Geosiphon (Mucoromycota, Glomeromycetes; see Sect. 6.1) nor the phenomena of photosymbioses and kleptoplasty as found in the Holozoa, Radiolaria, and Foraminifera (Figs. 1.1, 2.8; see Sect. 6.1). In the latter, the host

B. Büdel and T. Friedl

(fungus, animal, or heterotroph cercozoan) and the oxy-phototroph symbiont (alga or cyanobacterium) can live at least for some or a longer time separately without being functionally associated with each other. Defining the diverse algae as united by being permanently capable of oxygenic photosynthesis and not embryophytes (Melkonian et al. 1995) already lifted, at least partly, the discussion of what an alga is to a functional level. Here we complement that definition, following our preceding discussion, and define algae as those uni- and multicellular functional units that are hydro-passive and perform permanent oxygenic photosynthesis. Thus, the algae are extremely heterogeneous (Figs. 1.1, 2.8). Whether or not the definition of algae should include the prokaryotic cyanobacteria has often been a debate. Because they represent the single origin to which all algal plastids can be traced back (see our extensive discussion on plastid origins in Chap. 2) and because their visual appearance is mostly like eukaryotic algae (e.g., in size thus, they have traditionally been referred to as “blue-green algae”), they may also be regarded as algae. Therefore, there would be no conflict in including the prokaryotic cyanobacteria in the definition of algae, and we suggest including them in the definition of algae. Following present-day phylogenetic systems (e.g., Adl et al. 2019), algae are situated in various “supergroups” of the eukaryotes (Figs. 1.1, 2.8): the Archaeplastida, which encompass the three major lineages, green algae (Chloroplastida), red algae (Rhodophyta), and Glaucophyta (see Sects. 4.1, 4.2, and 4.3), the Stramenopiles with the heterokont algae (Heterokontophyta, see Sect. 5.1), the Alveolata with the Dinoflagellata (see Sect. 5.2) and Colpodellida, the Rhizaria with the cercozoans Chlorarchniophyta (see Sect. 5.3) and the enigmatic Paulinella (see Sect. 4.4), the two supergroups by their own, Haptista (see Sect. 5.5) and Cryptista (see Sect. 5.6), and finally the Euglenida (Excavata; see Sect. 5.4). Consequently, algae can be considered a complex network that spans almost all lineages of eukaryotes. Likely, only a single eukaryotic supergroup, the Amorphea, has been left without algae (Figs. 1.1, 2.8). However, there are many animals, e.g., the corals and other Anthozoa in the Holozoa (see Sect. 6.1), Geosiphon, and the lichens (Dikarya), that take advantage of photosynthesis by symbioses (Figs. 1.1, 2.8). In the context of algae, another popular term is often mentioned, the protists. Unlike New Cryptogams and algae, the term protist is defined by the exclusion of large groups of eukaryotes, i.e., the animals (Holozoa), plants (meaning bryophytes, lycophytes, ferns, and phanerogams of Chloroplastida) and fungi (meaning the “true fungi,” i.e., fungi sensu Adl et al. 2019). There is no reason other than convenience for combining the photoautotrophic algae, already

~1.600 mya: eukaryotic algae

~500 mya: fossil record of lichens (?)

filamentous green alga

morphological similarity only (!) Riccia-like liverwort

~34 mya: first ~125 mya: expansion of Cretaceous terrestrial grasslands revolution, insect pollination

~3.550 mya: first bacterial life

~600 mya: terrestrial algae

unicellular green alga

~2.600 mya: cyanobacteria with oxygenic photosynthesis

~2.400 mya: terrestrial biocrusts

~3 mya: shift to Pleistocene grassland bioms

molecular phylogenetic record

~200-250 mya: origin of ascomycete lichens

~450 mya: liverworts hornworts, mosses

Polytrichum-like moss

~320 mya: origin of seed plants

ascomycete lichens with cyanobionts

ascomycete lichens with chlorobionts

Fig. 1.2 From bacteria to lawn: evolution of algae, lichens, bryophytes, vascular plants, and landscapes. Original drawing Spindler & Büdel

Calothrix-like filamentous cyanobacterium with tapering trichomes

Nostoc-like filamentous cyanobacterium forming gelatinous colonies

Microcoleus-like filamentous cyanobacterium

(mya = million years ago)

From Bacteria to lawn...

1 Introduction: The New Cryptogams 5

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B. Büdel and T. Friedl

heterogeneous by themselves, with various assemblages of heterotrophic life forms from at least three supergroups of the eukaryotic tree of life (Amorphea, Excavata, and SAR). The diverse lineages of the New Cryptogams may not only be united by their hydro-passivity (see below) but also by analogous forms of their sexual reproduction, apart from the cyanobacteria, which lack sexuality. Historically, the traditional cryptogams have been viewed as plant-like organisms that reproduce in a hidden fashion because their sexual reproduction cannot be observed by the naked-eyed. In the algae, the diplontic life cycle has independently developed twice, i.e., in the photoautotrophic Stramenopiles (Diatomeae and Phaeophyceae in the Heterokontophyta; see Sect. 5.1; Figs. 1.1, 2.8), thus reaching the same level as multicellular animals (Holozoa). In the phanerogams, the reproductive organs are integrated into a reduced gametophyte enclosed within specific sporophyte structures for protection from desiccation. Phanerogams never left the level of haplo-diplontic change of generations, albeit with highly reduced male gametophytes. Today, the modern phanerogamic plants include gymnosperms, angiosperms, and form, together with the lycophyte ferns, a monophyletic unit (Tracheophyta; e.g., Ran et al. 2018). Further general aspects of the New Cryptogams are discussed in Sect. 2.5.

1.3

Functional Aspects of New Cryptogams: Hydro-Passivity

Water and life are inseparably bound together (e.g., Billi 2009). Despite their numerous origins, the great majority of oxy-phototroph (photoautotrophic) life forms are characterized by being hydro-passive. However, they are not necessarily poikilohydric regarding water uptake and loss. The New Cryptogams display a range of hydro-passivity, from strict desiccation intolerance in the aquatic lifestyle to the desiccation-tolerant, poikilohydric, and terrestrial lifestyle, in numerous examples. The marine and freshwater phytoplankton are entirely dependent on water, are desiccationsensitive, and cannot withstand drought at all. Other New Cryptogams exhibit some desiccation tolerance and are thus poikilohydric, as many cyanobacteria, some lineages of green algae, and lichens. There is evidence that early proand eukaryotic organisms capable of oxygenic photosynthesis were entirely dependent on water and could not stand desiccation at all. However, some early cyanobacteria, e.g., the genus Chroococcidiopsis, developed the ability to survive almost complete desiccation, a feature that reappeared

then later in evolution in the eukaryotes. When dry, such organisms change into a state of inactivity called anhydrobiosis, where no biological activity can be measured (Alpert 2006). Rehydration results in a more or less rapid resurrection of metabolic and photosynthetic activity. The transition from the wet to the dry stage and then back again into the wet stage also depends on the organism's structure. It ranges from seconds to minutes in unicells and may take hours in morphologically more complex forms. For an overview of desiccation tolerance, see, for example, Lüttge et al. (2011). Lichens are normally poikilohydric (see Sect. 6.2). Though some bryophytes might also be desiccation-sensitive, most are poikilohydric (see Sect. 7.6). The term “bryophytes” may be an artificial term of convenience. The common monophyletic origin of the hornworts, liverworts, and mosses (Anthocerophyta, Bryophyta, and Marchantiophyta still has not unambiguously been resolved (see Figs. 1.1, 2.8), whereas the shared monophyletic origin of liverworts and mosses (Marchantiophyta and Bryophyta) is not in doubt (for discussion, see Cox et al. (2014), and One Thousand Plant Transcriptomes Initiative (2019). The hydro-passive New Cryptogams never developed active transport of water through their bodies or formed multicellular structures for protection against drought. The vascular tissues developed by members of the Tracheophyta are responsible for the controlled uptake and distribution of water, as well as the circulation of photosynthetic products, thus, providing them extensive independence from the presence of liquid water. Consequently, they are hydro-active or homoiohydric in contrast to the New Cryptogams, which are hydro-passive. With only a few exceptions, the vascular plant sporophytes are hydro-active and control water uptake, normally exclusively by roots, and water loss, primarily via their leaves. Together with the ability to include the hydro-passive gametophyte phase into the structural entity of the sporophyte, these features unify the vascular plants (Tracheophyta). Tracheophytes that somewhat bridge the hydro-passive New Cryptogams with the hydro-active vascular plants are the ferns. Although the latter develop tiny hydro-passive gametophytes with a thalloid organization level, the main presence of ferns in vegetation is their sporophytes, which have a vascular plant organization and are hydro-active (therefore, they do not belong to the New Cryptogams). Ferns and other vascular plant groups are monophyletic with Gymnosperms and Angiosperms; they are regularly discussed in special publications and books (e.g., Frey et al. 2009).

1

Introduction: The New Cryptogams

1.4

The New Cryptogams Predated the Conquest of the Third Dimension on Land

Cyanobacteria, and probably photoautotrophic bacterial predecessors, dominated the aquatic (*3300 million years ago = Mya) and terrestrial habitats on Earth (2400 Mya). Eukaryotic algae followed in aquatic (1.5 Mya) and later in terrestrial habitats (roughly 600–700 Mya), whereas fungi (600 Mya?), bryophytes (470 Mya), and lichens (*200 Mya) followed in terrestrial habitats (Fig. 1.2; Sect. 6.2). The advent of vascular plants changed the Earth’s land surface dramatically. Their hydro-active nature allowed water transportation from soil to heights of more than 100 m. This enabled vascular plant vegetation to expand into the third dimension, up to about 100 m above the ground. Tall growing forest stands developed (300 Mya). Later, grasslands accrued as a vegetation type (about 34 Mya) and largely replaced the dominance of cryptogams wherever climatic conditions allowed (Figs. 1.2). Due to their hydro-active (homoiohydric) nature, the phanerogams outcompete poikilohydric organisms wherever a regular rainfall regime allows the formation of subterranean, root-accessible water. In general, grasses occur where annual precipitation rates roughly reach 300 mm and trees from 500 mm or more. Below this critical precipitation rate, the realm of the non-vascular oxygenic photoautotrophs, the New Cryptogams, spreads. Cyanobacteria, algae, lichens, and bryophytes are still omnipresent. Even in dense tropical forests, all the New Cryptogams form an important part of the vegetation and biodiversity, such as epiphytes on vascular plants or as crucial components of biological soil crusts. New Cryptogams, especially after disturbance events, are the initial re-vegetation kick-off which starts plant succession. The harsher the climatic conditions are, the fewer vascular plants are present, and members of the New Cryptogams dominate the vegetation. New Cryptogams have their realm as essential elements of vegetation succession on rocks and soils of deserts, semi-deserts, savannas, polar and high alpine regions, and on surfaces of long-living woody plants. They may not be as eye-catching as vascular plants, but New Cryptogams are everywhere. In the marine environment, vascular plants are rare and restricted to very few species and to coastal or shallow waters. The tidal zone of cold ocean regions and the open oceans are the domain of algae and cyanobacteria. Freshwater habitats are somehow different. The shallower regions of lakes and rivers are colonized by quite a number of vascular plants, liverworts, mosses, and lichens adapted to an, at least temporarily, submerged lifestyle. In these

7

habitats, planktonic and benthic algae are a plethora, with a high number of epiphytic algae growing on submerged vascular plants or submerged parts of plants.

1.5

The New Cryptogams and Chapter Selection of This Textbook

Our textbook emphasizes all lineages of the New Cryptogams, i.e., the algae, including the cyanobacteria, the lichens, and the bryophytes (Figs. 1.1, 2.8). A focus is on the many origins of plastids and algae. A more general overview of plastid origins that link the various algal lineages is provided in Chap. 2, including general aspects of the sexual reproduction of the New Cryptogams. We then present a broader discussion of the various structural and ecological features of cyanobacteria (Chap. 3) which, despite their enormous ecological and evolutionary importance, have previously often been underrepresented in textbooks. Guided by primary endosymbiosis, which initiated the spread of photosynthesis in the eukaryotes by using cyanobacteria as plastid-progenitors (see Sect. 2.2.1), the lineages of Archaeplastida and the enigmatic unrelated Paulinella are discussed in Chap. 4. Those algae lineages that originated from eukaryote-eukaryote (secondary) endosymbioses, i.e., the heterokont algae (Stramenopiles), Dinoflagellates (Alveolata), Haptista, Cryptista, and Euglenida (Excavata), are discussed in another single chapter (Chap. 5). Because the algae are involved in numerous symbiotic associations other than lichens, for which a decent account is presented in Sect. 6.2, more algal symbioses (though not included in the New Cryptogams) are discussed in Sect. 6.1. Finally, we discuss the more complex hydro-passive but desiccationtolerant (poikilohydric) liverworts, mosses, and hornworts in Chap. 7. To the best of our knowledge, no other work so far has brought such diverse groups of oxy-phototroph (photoautotrophic) organisms closely together in one textbook. We feel that this is important. In terrestrial habitats, other than in marine and freshwater habitats, cyanobacteria and algae are often overlooked due to their inconspicuously small size (mostly comprising of microscopic forms). We aim to pronounce the aspect of “evolution by cooperation” as an important and omnipresent evolutionary tool for life on Earth, demonstrating how important it is to gain knowledge about even the most inconspicuous organisms. We hope that with this textbook, we will reach a large audience and help to explore this fascinating and ecologically important group that was also crucial for the evolution of modern plants. We are aware that the presentation of such a diverse group of organisms in a textbook will always suffer from never being able to meet the latest outcomes of

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research. However, our focus is to present an overview of the stunning variety of oxy-phototroph (photoautotrophic) species. Species are the only natural unit existing in nature, independent of systematic arrangements that are often imperfect or artificial. This book is meant as an invitation to become aware of the fascinating diversity developed in amazing evolutionary processes over the history of life.

References Adl SM, Bass D, Lane CE, Lukeš J, Schoch CL, Smirnov A, Agatha S, Berney C, Brown MW, Burki F, Cárdenas P, Čepička I, Chistyakova L, del Campo J, Dunthorn M, Edvardsen B, Eglit Y, Guillou L, Hampl V, Heiss AA, Hoppenrath M, James TY, Karnkowska A, Karpov S, Kim E, Kolisko M, Kudryavtsev A, Lahr DJG, Lara E, Le Gall L, Lynn DH, Mann DG, Massana R, Mitchell EAD, Morrow C, Park JS, Pawlowski JW, Powell MJ, Richter DJ, Rueckert S, Shadwick L, Shimano S, Spiegel FW, Torruella G, Youssef N, Zlatogursky V, Zhang Q (2019) Revisions to the classification, Nomenclature, and Diversity of Eukaryotes. J Eukaryot Microbiol. 66: 4–119. https://doi.org/10.1111/jeu.12691 Alpert P (2006) Constraints of tolerance: why are desiccation-tolerant organisms so small or rare? J Exp Biol 209:1575–1584 Billi D (2009) Subcellular integrities in Chroococcidiopsis sp. CCMEE 029 survivors after prolonged desiccation revealed by molecular probes and genome stability essays. Extremophiles 13:49–57 Cox CJ, Li B, Foster PG, Embley TM, Civan P (2014) Conflicting phylogenies for early land plants are caused by composition biases among synonymous substitutions. Syst Biol 63:272–279 Elbert W, Weber B, Burrows S, Steinkamp J, Büdel B, Andreae MO, Pöschl U (2012) Contribution of cryptogamic covers to the global cycles of carbon and nitrogen. Nat Geosci 5:459–462 Esser K (1992) Kryptogamen 2: Moose, Farne. Springer-Verlag Berlin Heidelberg; p 220 Esser K (2000) Kryptogamen 1: Cyanobakterien, Algen, Pilze, Flechten 3rd ed. Springer-Verlag Berlin Heidelberg; p 585 Frey W, Stech M, Fischer E (eds) (2009) Part 3: Bryophytes and seedless vascular plants. Frey W (ed). Syllabus of plant Families—

B. Büdel and T. Friedl A Engler's syllabus der pflanzenfamilien. Stuttgart, Germany: Schweizerbart Science Publishers; p 419 Fischer E, Frey W, Theisen I (eds) (2015) Part 4: Pinopsida (Gymnosperms) Magnoliopsida (Angiosperms). Frey W (ed). Syllabus of plant families—A Engler's syllabus der pflanzenfamilien. Stuttgart, Germany: Schweizerbart Science Publishers. p 495 Guiry MD (2024) How many species of algae are there? A reprise. Four kingdoms, 14 phyla, 63 classes and still growing. J Phycol 60. https://doi.org/10.1111/jpy.13431 Hofbauer W, Kawai H, Nakayama T, Cox EJ, de Reviers B, Rousseau F, Silberfeld T, Neustupa J, Leliaert F, Lopez-Bautista J, De Clerck O, Leliaert F, Blindow I, Schudack M (eds) (2015) Part 2/1: Photoautotrophic eukaryotic Algae. Frey W (ed). Syllabus of plant Families—A engler's syllabus der Pflanzenfamilien. Stuttgart, Germany: Schweizerbart Science Publishers; p 324 Jassey VEJ, Walcker R, Kardol P, Geisen S, Heger T, Lamentowicz M, Hamard S, Lara E (2022) Contribution of soil algae to the global carbon cycle. New Phytologist. https://doi.org/10.1111/nph.17950. Kamiya M, Lindstrom SC, Nakayama T, Yokoyama A, Lin S-M, Guiry MD, Gurgel CFD, Huisman JM, Kitayama T, Suzuki M, Cho TO, Frey W (eds) (2017) Part 2/2: Photoautotrophic eukaryotic Algae—Rhodophyta. Frey W (ed). Syllabus of plant Families—A engler's syllabus der Pflanzenfamilien. Stuttgart, Germany: Schweizerbart Science Publishers, p 171 Lüttge U, Beck E, Bartels D (2011) Plant desiccation Tolerance. Ecological Studies 215: p 386, Springer-Verlag Berlin Heidelberg Melkonian M, Marin B, Surek B (1995) Phylogeny and evolution of the algae. In Biodiversity and Evolution Proceedings of the 10th international Symposium on Biology. Arai R. KM, Doi Y. (ed). Tokyo: The National Science Museum Foundation, pp 153–176. One Thousand Plant Transcriptomes Initiative (2019) One thousand plant transcriptomes and the phylogenomics of green plants. Nature 574:679–685. https://doi.org/10.1038/s41586-019-1693-2 Ran J-H, Shen T-T, Wang M-M, Whang X-Q (2018) Phylogenomics resolves the deep phylogeny of seed plants and indicates partial convergent or homoplastic evolution between Gnetales and angiosperms. Proceedings of the Royal Society Series B 285:20181012. https://doi.org/10.1098/rspb.2018.1012 Wang Y, Liu S, Wang J, Yao Y, Chen Y, Xu Q, Zhao Z, Chen N (2022) Diatom biodiversity and speciation revealed by comparative analysis of mitochondrial genomes. Front Plant Sci 13:749982. https://doi.org/10.3389/fpls.2022.749982

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Endosymbioses: Origin and Diversity of Photosynthetic Eukaryotes and Their General Genetic Exchange Modes Jan de Vries and Thomas Friedl

Contents 2.1 Introduction: The Role of Endosymbiosis in the Evolution of Eukaryotes.................................. 9 2.1.1 Endosymbiotic Gene Transfer and Reductive Evolution .......................................................... 9 2.1.2 Protein Import by Organelles ..................................................................................................... 11 2.2 Primary Endosymbiosis: The Endosymbiotic Origin of Plastids................................................... 12 2.2.1 An Ancient Endosymbiotic Event and the Cyanobacterial Progenitor of Plastids ................................................................................................................................... 12 2.2.2 The Origin of Archaeplastida..................................................................................................... 13 2.3 Eukaryote-Eukaryote Endosymbioses............................................................................................... 16 2.3.1 A Brief Overview of the Endosymbiotic Potpourri................................................................... 16 2.4 A Modern Overview of Eukaryotic Diversity—The Opuntia-Tree-of-Life .................................. 18 2.5 Life histories, Plastid Spread, and Sexual Reproduction in the New Cryptogams

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References ..................................................................................................................................................... 22

2.1

Introduction: The Role of Endosymbiosis in the Evolution of Eukaryotes

All eukaryotes are chimeras of the two known types of prokaryotes: archaea and bacteria (Fig. 2.1). The biological reason for this chimerism is endosymbiosis. It was through endosymbiosis that an archaeal host incorporated a free-living proteobacterium. How this came about is the subject of an elaborate debate (for a scholarly overview, see Martin et al. 2015). In brief, most of the theoretical framework entails complementary biochemistry between the J. de Vries (&) Institut für Mikrobiologie und Genetik, Abt. Angewandte Bioinformatik, Georg-August-Universität Göttingen, Goldschmidtstr. 1, 37077 Göttingen, Germany e-mail: [email protected] T. Friedl Department of Experimental Phycology and Culture Collection of Algae (EPSAG), Georg August University Göttingen, Nikolausberger Weg 18, 37073 Göttingen, Germany e-mail: [email protected]

archaeal and bacterial partner, resulting in a mutually beneficial cross-feeding of metabolic products (“syntrophy”). The critical outcome from this interaction is, however, clear: the archaeal host served as a chassis for what was to become the first eukaryotic cell, whereas the proteobacterial partner was the progenitor of today’s mitochondria. Thus, the evolution of eukaryotes is intertwined with endosymbiosis. And mitochondria are the mother of all endosymbiotically derived organelles. In the evolution of eukaryotic organelles, two of the main evolutionary processes determined how their molecular biology works are: (1) endosymbiotic gene transfer (EGT) and (2) establishment of protein import.

2.1.1 Endosymbiotic Gene Transfer and Reductive Evolution How did the engulfed proteobacterium turn into a mitochondrion? The answer is reductive evolution; the foremost mechanism behind this reductive force was the so-called

© Der/die Autor(en), exklusiv lizenziert durch Springer-Verlag GmbH, DE, ein Teil von Springer Nature 2024 B. Büdel et al. (eds.), Biology of Algae, Lichens and Bryophytes, https://doi.org/10.1007/978-3-662-65712-6_2

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10 Fig. 2.1 Endosymbioses shaped the evolution of eukaryotes. An endosymbiotic event marks the origin of eukaryotes. Likely more than 2 billion years ago, incorporating a proteobacterial cell into an archaeal cell gave rise to the first eukaryotes. Some of the fully fledged eukaryotes that emerged from this event engaged in another fateful endosymbiosis: at least 1.5 billion years ago, a cyanobacterial cell was incorporated by a heterotrophic unicellular eukaryote, giving rise to the first representatives of the archaeplastidal cell. Archaeplastida is the lineage of eukaryotes with primary plastids, which radiated into Glaucophyta, Rhodophyta, and Chloroplastida (see Figs. 1.1 and 2.8). Original drawing F. Spindler & T. Friedl

J. de Vries and T. Friedl

prokaryotic cell uptake of a proteobacterial cell

eukaryotic cell endosymbiotic origin of mitochondria

endosymbiotic gene transfer (EGT). Before we delve into this matter, it is prudent to note that the endosymbiotic gene transfer is a general pattern that can be observed following various independent endosymbiotic events—from the origin of mitochondria. So how does endosymbiotic gene transfer work? Endosymbiotic gene transfer describes the relocation of genes from endosymbionts (and nascent organelles) to the nucleus (Fig. 2.2). The process is thought to hinge on a series of random events that start with the lysis of the endosymbionts. Through natural and random lysis of the endosymbiont cells (as part of their life and death while swimming in the host’s cytosol), their DNA is released. Sometimes, the DNA of the endosymbionts ends up in the nucleus of the host. Mediated by the action of DNA repair enzymes (e.g., shown for yeast mitochondrial DNA; Ricchetti et al. 1999), random parts of the endosymbiont DNA become incorporated into the host genome. This can result in the incorporation of large pieces of endosymbiont DNA. Indeed, in several flowering plant genomes, including the genome of the model system for flowering plants, Arabidopsis thaliana, there are several thousands of bases long insertions of organellar DNA (Huang et al. 2005; Michalovova et al. 2013). How do we know that such endosymbiotic gene transfer has occurred? There are multiple lines of evidence. The first is simply the amount of the genetic material (i.e., the size of the

endosymbiotic origin of plastids

plastid genome) found in extant plastids. Based on the similarity of the genes found in plastid genomes to those of cyanobacteria, we can tell that this material clearly is of cyanobacterial origin. Yet, at the same time, plastid genomes have a mere fraction of the number of genes found in a free-living cyanobacterium (see more on plastid genomes in the sub-chapter below). Here, the second line of evidence comes into play. Many of exactly those genes that one would expect to find in a cyanobacterial genome that are missing from the plastid genome are indeed found in the nucleus. Thus, the genes got not merely lost but, before the loss, found their way to the nucleus (or rather the reason for their loss being that they ended up in the nucleus). Overall, the photosynthetic organism thus still has the cyanobacterial genes needed to maintain the functionality of the plastid. These genes are merely spread out over different genetic compartments. As just described, the process of endosymbiotic gene transfer hinges on a genetic redundancy between the organellar genome and the nuclear genome that becomes increasingly enriched in genes through EGT. But it is more complicated. In order to create such redundancy having the genes is not enough. The critical point is that the gene products (i.e., in most cases, the proteins) end up in the place where they are needed—that is, these proteins had to find their way back to the organelle. This process is carried out by post-translational protein import machinery, and this is what the next sub-chapter is all about.

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Endosymbioses: Origin and Diversity of Photosynthetic Eukaryotes and Their General Genetic Exchange Modes

cyanobacterium with 2.000 to 12.000 genes

11

primary host

primary endosymbiosis

lysis

massive EGT

endosymbiont

plastid with 300 genes

ancestral archaeplastid

Fig. 2.2 Endosymbiotic gene transfer and the last common ancestor of Archaeplastida. The cyanobacterial progenitor of plastids likely had a couple of thousand up to even more than 10,000 genes. After being incorporated by the primary host, a heterotrophic single-celled eukaryote, the cyanobacterium lived as an independent endosymbiont population. Within the cytosol of the host, this population propagated via fission and died, resulting in lysis and the release of all its DNA. Perchance, some of the thusly released DNA was integrated into the nucleus of the host. If the products (proteins) of these now nuclear genes found their way back to the endosymbiont/plastid through protein import, a genetic redundancy between the nuclear genome and endosymbiont genome was created. Under these conditions, constraints on retaining genes were lifted, eventually resulting in the shrinkage of the endosymbiotic genome. This process circumscribes the massive endosymbiotic gene transfer (EGT) that resulted in the coding capacity of the last common ancestor of Archaeplastida and, eventually, extant plastids. Original drawing F. Spindler & J. de Vries

2.1.2 Protein Import by Organelles One of the hallmarks of the eukaryotic cell is its compartmentalization. The question of where in a eukaryotic cell proteins reside is a critical determinant of the functions these proteins can carry out. Thus, in eukaryotes, cellular logistics is essential. This is also true when it comes to the organelles of eukaryotic origin. As described above, most genes of the bacterial progenitors of the mitochondrion and plastid were transferred to the nucleus. In extant plants and algae, the products of these genes (proteins) are spread out over any locality that we find in a eukaryotic cell: in the cytosol, in the endoplasmic reticulum, being secreted to the outside of the cell, and so on.

Mitochondria and plastids have two membranes. Both membranes (the outer and inner membrane) are most likely homologous to the two membranes of their gram-negative bacterial progenitors. For a free-living gram-negative bacterium, the outer membrane is the interface for interacting with the environment; for an organelle, the outer membrane is the interface for interaction with the host cytosol. It is thus at the outer envelope where the first committed steps in protein import into organelles occur: the recognition by receptor proteins that establish specificity in protein import (Fig. 2.3). Both mitochondria- and plastid-targeted proteins encoded in the nucleus have an n-terminal extension plugged onto the native protein sequence. Together, the n-terminal extension

12 Fig. 2.3 Protein import into mitochondria. Mitochondria are organelles that are surrounded by a double membrane. Proteins imported from the cytosol can be translocated to the outer envelope, inner envelope, intermembrane space, and matrix. Protein import is carried out by large protein complexes, most notably the complexes TIM (translocon at the inner mitochondrial membrane) and TOM (translocon at the outer mitochondrial membrane). Targeting the mitochondrion hinges on a transit sequence removed via proteases after import. Original drawing F. Spindler & J. de Vries

J. de Vries and T. Friedl incoming protein TOM complex

outer membrane

SAM complex

Tiny Tims

MIA

OXA1

TIM23 complex

TIM22 complex

inner membrane that is plugged onto the native protein sequence makes up the precursor proteins that are targeted to the organelles. These precursors are biosynthesized by the cytosolic ribosomes. At the outer organellar envelope, receptors recognize the n-terminal extension and initiate the process of protein import (Figs. 2.3 and 2.4). The import across the first outer membrane is carried out by the complexes TOM (translocon at the outer mitochondrial membrane) in mitochondria and TOC (translocon at the outer chloroplast membrane) in plastids. Import across the second inner membrane is then carried out by TIM (translocon at the inner mitochondrial membrane) and TIC (translocon at the inner chloroplast membrane), respectively. After the precursor protein has thusly reached the mitochondrial matrix or plastidial stroma, the n-terminal sequence extension is cleaved off, and the proteins are folded into their native confirmation through the action of chaperones. The critical reader will notice that the just described mechanism does not explain how proteins end up, for example, in the organellar membranes—this is a more involved process that includes additional non-cleavable sequence stretches for sorting and shall not be elaborated upon here. The evolutionary origin of the protein import machinery has allowed the integration of the bacteria-derived organelles into the cell biology of the host. Protein import has cemented

the shift from a (relatively) self-sufficient prokaryotic entity to a subcellular structure that was dependent on the cell biology of its host. Further, the need for protein import has also created a need for tight communication between organelle and host, ultimately creating the tight links that nowadays integrate the organelles into almost any process in a eukaryotic cell. After this general introduction, the following parts of this chapter will delve into the specific biology of plastids and the roles endosymbiotic events have played in their evolutionary history. However, it is important to reiterate that the endosymbiotic gene transfer and the question of how protein import was realized are recurrent themes.

2.2

Primary Endosymbiosis: The Endosymbiotic Origin of Plastids

2.2.1 An Ancient Endosymbiotic Event and the Cyanobacterial Progenitor of Plastids Photosynthetic eukaryotes are everywhere. Important for now is that all of this diversity (see Figs. 1.1 and 2.8) can

2

Endosymbioses: Origin and Diversity of Photosynthetic Eukaryotes and Their General Genetic Exchange Modes

Fig. 2.4 Protein import into plastids. Like mitochondria, plastids are organelles that are surrounded by a double membrane. Proteins imported from the cytosol can be translocated to the outer envelope, inner envelope, intermembrane space, and matrix. Protein import is carried out by large protein complexes, most notably the complexes TIC (translocon at the inner chloroplast membrane) and TOC (translocon at the outer chloroplast membrane). Targeting the plastid hinges on a transit sequence removed via proteases after import. Original drawing F. Spindler & J. de Vries

TOC complex

13

incoming protein

TOC 75-V

outer membrane

inner membrane

TIC complex

ultimately be traced back to a fateful endosymbiotic event. In this case, an ancient heterotrophic unicellular eukaryote with a nucleus, mitochondria, and all other typical eukaryotic features incorporated a free-living cyanobacterium (Figs. 2.2 and 2.5). The nature of that particular cyanobacterial progenitor to plastids is still being debated. Indeed, different analyses recover that it might have been a filamentous, nitrogen-fixing cyanobacterium or that it was a unicellular one. The interested reader is referred to, e.g., Dagan et al. (2012), de Vries and Archibald (2017), and Ponce-Toledo et al. (2017). This notion warrants attention. In extant photosynthetic eukaryotes, there are a couple of symbiotic interactions that are built on cyanobacterial nitrogen fixation —such as the diatom Rhopalodia bearing intracellular nitrogen-fixing cyanobacteria, the fern Azolla that houses cyanobionts in its leaves, and more. Hence, fixed nitrogen is a powerful currency of symbioses. Another interesting train of thought emerging from the most recent analyses is that the cyanobacterial plastid progenitor likely lived in a freshwater setting (Delwiche and Cooper 2015; Ponce-Toledo et al. 2017; Sánchez-Baracaldo et al. 2017; Lewis et al. 2017).

This places the origin of photosynthetic eukaryotes in a freshwater environment—and not a marine setting. What this also means is that the conquest of marine environments by algae is a secondary adaptation to living in salt water.

2.2.2 The Origin of Archaeplastida No matter what the exact nature of the cyanobacterial plastid progenitor was, the main outcome is the same: through endosymbiosis, a free-living cyanobacterium that had the ability to photosynthesize was incorporated by an ancient but fully fledged eukaryote. Estimates place this event at least 1.5 billion years before the present (Eme et al. 2014)— but it might have occurred in even more distant times. With the incorporation of the photosynthesizing cyanobacterium, the lineage of photosynthetic eukaryotes was borne: the Archaeplastida. Before the Archaeplastida split into any other surviving lineage of descendants, most of the core processes described before (Sect. 2.2.1) occurred. The last (most recent) common ancestor of Archaeplastida (Fig. 2.5)

14

J. de Vries and T. Friedl

Fig. 2.5 Primary endosymbiosis —from heterotrophy to photoautotrophy and three major groups of plastids which the Archaeplastida lineage comprises. The endosymbiotic incorporation of the cyanobacterial plastid progenitor (for details, see Fig. 2.2) marks the birth of the lineage of Archaeplastida, i.e., eukaryotes with primary plastids. Most of the transformative events that led to algal cell biology as we know it (such as the protein import and thus the division of labor between the genetic compartments) likely transpired before the last common ancestor (LCA) of all Archaeplastida came about. From that LCA, three major lineages emerged: the Rhodophyta (red algae), the Glaucophyta (a species-poor group), and the Chloroplastida (the green lineage of green algae and land plants). Original drawing Spindler & Friedl

incorporation of the cyanobacterial plastid progenitor

last common ancestor

Archaeplastida

Rhodophyta Glaucophyta

Chloroplastida had a fully established protein import machinery, as evident by the set of homologous components shared by all Archaeplastida. Furthermore, much of the reduction in the coding capacity of the nascent organelle also occurred early. Based on comparative genomics of the now available thousands of plastid genomes, we can estimate that the plastid genome of the last common ancestor of Archaeplastida likely had a few hundred protein-coding genes. This stands in stark contrast to the 3000 up to whooping 12,000 protein-coding genes found on the genomes of extant cyanobacteria. After these initial events have molded a eukaryotic organelle out of the once free-living cyanobacterium, the plastid-bearing lineage eventually split into the three primary plastid-bearing lineages of eukaryotes: the Glaucophyta, the Rhodophyta (red algae), and the Chloroplastida (the green lineage). Together, these lineages constitute the Archaeplastida (Fig. 2.5). The Glaucophyta is probably the most enigmatic of the three lineages of Archaeplastida. There are at the time only 15 extant species described that fall into the

glaucophyte lineage (see Sect. 4.3). One of the most astounding features of the Glaucophyta is that their plastid is surrounded by a thick peptidoglycan layer—just as in free-living cyanobacteria. All other plastids do not have a thick peptidoglycan layer. It is particularly due to this layer (and additional features such as carboxysomes) that, for a long time, it was thought that the glaucophytes had retained many ancestral features—going hand in hand with a frequently retrieved topology for the tree of Archaeplastida in which glaucophytes branched sister to all other Archaeplastida (Figs. 2.5, 2.6 and 2.8). However, currently, the position of glaucophytes being a “missing link” must be revisited. In the most recent phylogenomic analyses, the Rhodophyta branch is sister to all other Archaeplastida. Furthermore, while it still holds that only glaucophytes have a thick peptidoglycan layer around their plastids, Herrero et al. (2016) recently showed that moss chloroplasts have a thin peptidoglycan layer (Hirano et al. 2016). More details on the three major lineages of Archaeplastida can be found in Chap. 4 of this book.

2

Endosymbioses: Origin and Diversity of Photosynthetic Eukaryotes and Their General Genetic Exchange Modes

Fig. 2.6 Secondary endosymbioses—the lateral spread of plastids across the tree of eukaryotes. Through eukaryote-eukaryote endosymbioses (secondary or higher-order symbioses), plastids spread to other lineages, oftentimes distantly related to the Archaeplastida. This occurred at least two times, involving a primary chloroplastidial (green) alga and a heterotrophic host, giving rise to Euglenida and Chlorarachniophyta. Additionally, likely several secondary (and maybe even higher-order) endosymbioses resulted in the spread of rhodophyte (read algal)-derived plastids to algal groups such as the Stramenopiles, Alveolata, Haptista, and Cryptista (see Fig. 2.8). At least in the Stramenopiles the secondary endosymbiosis with a red algal cell likely was preceded by an earlier secondary endosymbiosis with a green algal cell (red-green mosaicism or chimerism; see Sects. 5.1 and 5.2). Original drawing F. Spindler & T. Friedl

15

last common ancestor primary endosymbiosis

Archaeplastida

glaucophyte

green alga red alga secondary endosymbioses

Euglenida

2.2.2.1 Plastid Genomes As outlined above, much of the EGT-driven reduction of the cyanobacterial genome toward an organellar genome occurred before the Archaeplastida diversified. Therefore, plastid genomes have many shared features. Therefore, plastid DNA is a good biomolecular marker for tracing endosymbiosis and evolution. So, what exactly are plastid genomes? Plastid genomes (or short, plastomes) are miniature cyanobacterial genomes. As such, they map as a circular genome that contains between about 80–250 protein-coding genes. In most organisms, most of the plastid genome comprises these protein-coding genes; thus, there is little “intergenic space” between the genes and generally few non-coding regions. This genomic efficiency notwithstanding, there are cases of (a) extremely bloated and bizarre

Chlorarachniophyta

Stramenopiles Alveolata Haptista Cryptista

plastid genomes and (b) a peculiar phenomenon called the inverted repeats. Many plastid genomes (but not all) contain two identical stretches of sequence information. These regions face each other; that is, one is a flipped version of the other—hence the name “inverted repeats.” Due to the inverted repeats, plastid genomes can be separated into four regions: the large single-copy region, the small single-copy region, and the two inverted repeats. Another feature shared by all plastid genomes is that they are usually low in GC content—notably lower than the genome of the nucleus with which they share a cytosol. All of this said—and as is usually the case in biology—there are many exceptions to these features. There are plastid genomes without inverted repeats, large intergenic regions, high GC content, etc. What tends to be more conserved is what proteins are encoded by these genomes.

16

While the plastid clearly depends on the nucleus, the genome of the plastid necessitates that certain systems required for basal molecular “cell” biology are in place. A plastid genome usually codes for quintessential proteins involved in information processing, such as RNA polymerase and ribosomal proteins; furthermore, also non-protein-coding genes are on a plastid genome, including tRNAs and the ribosomal RNAs. It goes without saying that all these factors involved in information processing are clearly of bacterial origin. Machineries for information processing are only advantageous when there is relevant information to be processed. Hence (as the critical reader might have already guessed), most of the other proteins encoded on plastid genomes can be tied to functioning in photosynthesis. This includes genes for photosystem II and I proteins, other components of the electron transport chain of the light reaction, and the ATP synthase complex. Interestingly, while plastid genomes of all photosynthesizing algae and plants code for proteins of the light reaction, genes whose protein products act in the process of carbon fixation are mostly limited to red algae. Overall, it can be said that red algae (Rhodophyta) have roughly 1.5 to two times the number of protein-coding genes on their plastid genomes as compared to green organisms (Chloroplastida or Chlorobionta). Plastid genomes have genes that code for hallmark proteins of, for example, the photosystems. That said plastid genomes do not code for all components of a given photosystem complex. Each of the protein complexes involved in the light reaction of the photosynthesis chain is a chimera of proteins encoded by plastid genes and nuclear genes. The same holds even for information processing—for example, the sigma factors for the plastid RNA polymerase are encoded on the nuclear genome. Furthermore, many of these plastid basal processes have been modified by recruiting nuclear (host) proteins. Most prominent among these modifications likely is the addition of the light-harvesting complexes around photosystems II and I. These are an evolutionary innovation of the green lineage, i.e., the Chlorobionta (and a defining feature thereof)—which is therefore solely encoded in the nucleus. The Chlorobionta also gained nuclear-encoded proteins targeted to the plastid, where they exercise regulatory control over the prokaryotic RNA polymerase. And many more such examples exist. This touches upon an important aspect that shall not be discussed further here: the need for communication and coordination of the two genetic compartments in which genes for plastid function reside—the plastid and the nucleus. The interested reader can find more on these topics in the scientific literature under the terms anterograde and retrograde signaling.

J. de Vries and T. Friedl

2.3

Eukaryote-Eukaryote Endosymbioses

2.3.1 A Brief Overview of the Endosymbiotic Potpourri In the previous sub-chapters, we discussed the general processes that revolve around the endosymbiotic incorporation of a prokaryote (the cyanobacterial plastid progenitor) into a eukaryotic cell. The diversity of endosymbioses features, however, an even more astounding phenomenon: secondary endosymbiosis. This process involves the endosymbiosis between two eukaryotes—one being the (in most cases) heterotrophic host, the other being the photosynthetic eukaryotic alga (Fig. 2.6). Just as it happened after the incorporation of a prokaryote, the photosynthesizing endosymbiont was ultimately reduced through the process of EGT-driven reductive evolution and establishment of protein import. However, in this case, the reductive forces acted on an entire eukaryotic cell—not only carrying a plastid but with its own nucleus, endomembrane system, mitochondria, etc. This was likely as messy as it sounds—and we will later in this sub-chapter (and other chapters in this book) give some examples of the cell biological peculiarities that have spawned from these events. The number of secondary (and possibly even tertiary; see Fig. 5.47, Sect. 5.2) endosymbiotic events that have occurred is still being debated. However, even the most conservative interpretations agree that it happened at least three times (Fig. 2.6)—and most researchers would argue more often. Twice, secondary endosymbiosis involved distinct hosts that took up a chlorophyte (green) alga; these events gave rise to the chlorarachniophytes ((Chlorarachniophyta, Fig. 2.8; see Sect. 5.3) and the euglenophytes (Euglenida, Fig. 2.8; see Sect. 5.4), both of which are thus being called “secondary green” (due to the incorporation of a green plastid via secondary endosymbiosis; Fig. 2.6). Phylogenetic analyses indicate that the primary green algae that were incorporated through secondary endosymbiotic events are phylogenetically quite distinct; Jackson et al. (2018) pinpointed that euglenophyte plastid to have likely originated from the incorporation of a prasinophyte green alga while the chlorarachniophyte plastid likely had an ulvophyceaen progenitor (for details on these two green algae groups, see Chap. 4). When it comes to the secondary red endosymbios(is/es), things get complicated (Figs. 2.6 and 2.7). Various lineages have acquired red plastids through secondary endosymbiosis. While the exact nature of the red algae acquired through secondary endosymbiosis is not clear (and probably lost in time), it is clear that secondary red algae all have plastids that ultimately stem from rhodophytes. When it comes to the

2

Endosymbioses: Origin and Diversity of Photosynthetic Eukaryotes and Their General Genetic Exchange Modes

green alga

a

b

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c Stramenopiles Alveolata* Haptista Euglenida*

Cryptista Chlorarachniophyta

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d

17

e

Fig. 2.7 Remnants of cell-cell fusions during secondary endosymbioses: nucleomorphs. a–c From heterotrophic Stramenopiles-like motile cells (a) to the uptake of a green algal cell (b) or a red algal cell (c) by independent secondary endosymbioses; d Both the secondary green chlorarachniophytes (Chlorarachniophyta), as well as the secondary red cryptophytes (Cryptista) bear a remnant nucleus, called nucleomorph, which is the small nucleus-like structure in the periplastidial compartment. The latter corresponds to the cytoplasm of the primary (eukaryotic) alga that was incorporated. The schematic cell shown in (d) represents a member of Cryptista. e Not all the algae that have emerged from secondary (or higher-order) endosymbioses bear a nucleomorph, i.e., Stramenopiles, Alveolata, Haptista, and Euglenida. The schematic cell in (e) shows a Stramenopiles cell. An arrow indicates the chloroplast endoplasmatic reticulum (CER; arrow), which arose from the fusion of the outer pair of plastid membranes with the nuclear envelope, and is a characteristic feature of complex plastids. Such complex plastids are lacking in the Alveolata and Euglenida (marked by an asterisk). Original drawing F. Spindler & T. Friedl

host cells of the secondary red algae, things are less clear. The host cells are spread out over several “supergroups” of eukaryotes (although the term “supergroups” has recently fallen into disgrace). They include (1) Stramenopiles, which are best-known for including the plastid-bearing brown algae and diatoms (the photosynthetic lineages of Stramenopiles form the Heterokontophyta Fig. 2.8; Sect. 5.1). However, Stramenopiles also encompass, for example, oomycetes, which are mostly pathogenic/parasitic organisms that resemble fungi but are, of course, completely unrelated to the “true” fungi which belong to the Amorphea supergroup (Fig. 2.8); (2) Alveolata (Fig. 2.8), which include some of

the most bizarre plastid-bearing organisms, the dinoflagellates (Fig. 5.47; Sect. 5.2). The latter also include non-photosynthesizing but plastid-bearing organisms like the causal agent of malaria (Plasmodium), but also, for example, the ciliates; (3) Haptista, which include photosynthesizing coccolithophorids like Emiliania but also non-plastid-bearing centrohelids (Fig. 2.8; Sect. 5.5); (4) Cryptista, which are best-known for the photosynthesizing cryptophytes, but also include little-known heterotrophic protists such as Palpitomonas (Fig. 2.8; Sect. 5.6). Through secondary endosymbioses, many features and genetic material of primary plastid-bearing organisms (the

18

Archaeplastida) have jumped into more distantly related eukaryotes. The cell biological consequences of secondary endosymbiosis were manifold. In the following, we will discuss three main aspects: EGT, protein targeting, and the peculiar case of nucleomorphs (Fig. 2.7).

2.3.1.1 Secondary Endosymbiosis: EGT Mayhem While in the case of the primary endosymbiotic origin of plastids, a prokaryote (cyanobacterium) was incorporated into a eukaryotic cell, secondary (and higher-order) endosymbiosis is the merger of only eukaryotic cells. The primary algae incorporated by the host cells during secondary endosymbiotic events bring along their nuclear genome, plastid genome, and mitochondrial genome. All this genetic material ends up in a cell with its own nucleus and mitochondrion (and, in some cases, it is even suspected already their own plastid—but we shall not dwell on this issue here). This means that during the early events that transpired after secondary endosymbioses, at least 5 genetic compartments were included in a single cell, i.e., the symbiont plastid, symbiont nucleus, symbiont mitochondrion, host nucleus, and host mitochondrion. In extant organisms that harbor plastids of secondary origin, the organization of the genetic compartments resembles the situation observed in primary algae (the exception of nucleomorphs will be discussed in a separate sub-chapter below): they have a nuclear genome, plastid genome, and mitochondrial genome. The plastid genome contains a similar amount of—or even less—protein-coding genes as those plastid genomes of primary algae. Furthermore, the genetic material of the mitochondrion and nucleus of the primary alga no longer form their own chromosome (but see below). This means that the primary algae’s genetic material has been seamlessly merged with the host. Therefore, not only genes that ultimately came from the cyanobacterium—in the form of the genetic material that has been transferred to the nucleus during the early evolution of the plastid—but also additional eukaryotic genes from the archaeplastidal nucleus ended up in the nucleus of the secondary algae. In sum, in algae with plastids of secondary origin, genetic material with three very different evolutionary histories is merged: the host nucleus, the nucleus of the primary alga, and the genetic material that ultimately came from the cyanobacterial plastid progenitor.

J. de Vries and T. Friedl

—not in the plastid. Further, EGT is usually unidirectional. Hence, plastids usually do not get enriched with protein-coding genes again; as outlined above, the numbers also speak to the fact that genetically, a cell with a secondary plastid works the same way as a cell with a primary plastid. But the genetic consequences are just one aspect to consider —the other aspects are the cell biological implications of secondary endosymbiosis. Plastids of secondary (or potentially even higher-order) origin are surrounded by three or four membranes (Figs. 2.6, 2.7 and 5.47). How this situation came about is still being debated. What is, however, undebatable is the consequences this had for protein translocation: a nuclear-encoded protein that is bound to end up in the plastid has to find its way from the cytosol across additional membranes of the host endoplasmic reticulum.

2.3.1.3 Nucleomorphs Secondary endosymbiosis is a baffling process. That said, as outlined above, there is overwhelming evidence from both genetic data and cell biological explanations. One of the most tangible lines of evidence for incorporating an entire eukaryotic cell (i.e., the alga) by another eukaryote is that some secondary plastids still have a remnant nucleus—the nucleomorph (Fig. 2.7). Nucleomorphs are miniature nuclei. They came about through drastic reduction of the fully fledged eukaryotic nucleus of the primary algae that have been incorporated during secondary endosymbioses (Fig. 2.7). Nucleomorphs as remnant primary algal nuclei came about on at least two occasions: in the secondary red cryptophytes (Cryptista) and the secondary green chlorarachniophytes (Chlorarachniophyta). Interestingly, though both endosymbiotic events that have resulted in plastid acquisition are completely independent of one another—and hence have chartered a completely independent evolutionary course after secondary endosymbiosis—they have convergently resulted in a similar situation: nucleomorphs have three chromosomes, in total coding for about 300–500 proteins. Like most organellar genomes (and prokaryotic genomes), the nucleomorph chromosomes are extremely gene-dense—i.e., the amount of non-coding regions is minuscule.

2.4 2.3.1.2 Complexities of Protein Targeting and Membrane Since secondary endosymbiosis entailed the incorporation of a fully fledged photosynthetic eukaryote (which acted as the donor of photosynthesis), the division of labor between the genetic compartments was pre-defined: in the primary alga that was incorporated, most protein-coding genes required for the maintenance of plastid function reside in the nucleus

A Modern Overview of Eukaryotic Diversity—The Opuntia-Tree-of-Life

An up-to-date overview of the diversity of eukaryotes clearly highlights the multiple origins of algae and New Cryptogams, which is the particular theme of this book, through endosymbioses (Fig. 2.8). The scheme also highlights numerous successful cases of cooperation on which the evolution of eukaryotes is based on. Chimerism, symbioses,

Endosymbioses: Origin and Diversity of Photosynthetic Eukaryotes and Their General Genetic Exchange Modes

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Fig. 2.8 The Opuntia-tree-of-life, a modern view on the phylogenetic origins of the algae by endosymbioses and the New Cryptogams, the particular theme of this book. Endosymbiosis is seen as evolution through cooperation, which is central to the evolution of eukaryotes and New Cryptogams. Clubbed presentation of groups (“leaves of the Opuntia”): green, photosynthetic; gray, primarily non-photosynthetic. Curved arrows, origins of plastids by endosymbioses. Arrows encircling the Opuntia-tree-of-life, symbiotic associations (photosymbioses) or kleptoplasty. The Eukarya are seen as the chimera following the fusion of Bacteria and Archaea, which developed from the last universal common ancestor (LUCA). Through primary endosymbiosis, the Cyanobacteria gave rise to the Archaeplastida, which led to the diversification of the main algal lineages Chloroplastida, Rhodophyta, and Glaucophyta. Chloroplastida finally led to the land plants comprising the bryophytes and tracheophytes. Secondary endosymbioses gave rise to additional algal lineages within other main groups of eukaryotes, i.e., the Stramenopiles (Heterokontophyta), Alveolata (Dinoflagellata, Colpodellida), Rhizaria (Chlorarachniophyta, Paulinella), Haptista, Cryptista, and finally the Euglenida (Excavata). Tertiary endosymbioses with members of Cryptista, Haptista, and Chloroplastida gave rise to plastids in some Dinoflagellata. Arrows encircling the Opuntia-tree-of-life mark symbiotic associations (photosymbiosis or kleptoplasty) independent of plastid origins in which the New Cryptogams are involved. Many primarily non-photosynthetic lineages are symbiotically associated with Cyanobacteria, Dinoflagellata, Xanthophyceae, and Chlorophyta to take advantage of the photosynthetic lifestyle (photosymbioses). Temporary chloroplasts, found in the Dinoflagellata and some Holozoa, originated from Cryptista, Haptista, Xanthophyceae, and Chlorophyta through the phenomenon of kleptoplasty. Names of the various groups of eukaryotes follow Adl et al. (2019). Original drawing F. Spindler & T. Friedl

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and cooperation between different unrelated lineages (taxonomic groups) are keys to describing the evolution of eukaryotes. Endosymbioses, photosymbiosis, and kleptoplasty are more detailed terms for the numerous acquisitions of the photosynthetic lifestyle, which was made possible through the cooperation of unrelated taxonomic groups of which the New Cryptogams consist (Fig. 2.8). Their success in evolution would not have been possible without the cooperation among them—no survival without cooperation. We are quite convinced that there are still even more essential associations and interactions between unrelated groups of organisms waiting to be discovered. If we look at the dynamic field of microbiome research, where it is more and more realized that likely every larger organism has its own characteristic microbiome, sometimes even on an individual level, there remains still a lot to learn about the “evolution via cooperation.” A leaf-like structure of every single element allows for the most satisfying presentation we could think about (Fig. 2.8). Thus, the whole scheme ends up as a clubbed presentation of the different lineages and clades of organisms (see also Bessey 1915), and the resulting tree of life resembles an Opuntia-cactus (which, albeit is not a member of New Cryptogams). We use the term “Opuntia-Tree-of-Life” (Fig. 2.8) to name it. The many names shown by the leaves and the numerous organism/organism interactions indicated by the arrows surveyed in that scheme may be quite puzzling at first glance. To disentangle those is not an easy task—that it is so complicated is another essential feature of eukaryotes and the New Cryptogams. Therefore, it is just moderate to abstain from attempting to summarize them here. However, endosymbiosis may form the central theme that guides through the evolution of eukaryotes and New Cryptogams. First, the incorporation of a cyanobacterium that remained photosynthetically active (plastid progenitor) led to the primary endosymbiotic origin of the plastids and the major eukaryotic lineage of the Archaeplastida (Fig. 2.5). A second independent primary endosymbiosis between a cyanobacterial prokaryote and a eukaryotic cell happened later in evolution, i.e., within another major eukaryotic lineage, i.e., in Paulinella (Paulinellidae, Cercozoa, Rhizaria) (Fig. 2.8; see Sect. 4.4). Secondary endosymbiotic events with cells of descendants of a primary endosymbiosis serving as a source for plastids, i.e., the red and green algae, led to several additional lineages of algae spread on a diverse array of lineages of unrelated eukaryotes, i.e., Stramenopiles, Alveolata, Haptista, Cryptista, Chlorarachniophyta (Cercozoa, Rhizaria), and the Euglenida (Fig. 2.8; see Chap. 5). A rainbow of plastids of diverse origins from the Cryptista, Haptista, and the Diatomeae (Stramenopiles) is exhibited in the Dinoflagellates (Dinoflagellata, Alveolata) by independent tertiary endosymbioses and an additional secondary

J. de Vries and T. Friedl

endosymbiosis with a green algal cell (Figs. 2.8 and 5.47; see Sect. 5.2). Numerous other lineages of eukaryotes acquired the advantages of photosynthesis through (additional) symbiotic associations with algal and/or cyanobacterial cells, often with the incorporation of the photosynthetic cells in their bodies or cells (photosymbioses). Cyanobacteria are partners in symbiotic associations with (other) photoautotroph algae (Haptista, Diatomeae, Dinoflagellata), the Radiolaria (Rhizaria), Embryophyta (Streptophyta), and the fungi (lichens, and Geosiphon). Figure 2.8 depicts only a few examples from the broad variety of the many cyanobacterial symbioses described in Chap. 6. Dinoflagellates (Dinoflagellata, Alevolata) are known to form photoautotrophic partners in photosymbioses with the Holozoa (e.g., corals) and within the Rhizaria, with the Foraminifera, and Radiolaria (Fig. 2.8; see Sect. 6.1.9). Members from various lineages of the Chlorophyta serve as the phototroph partners in symbioses with fungi (lichens), Holozoa (e.g., sea anemones), Radiolaria, and Foraminifera (Fig. 2.8; see Chap. 6). A few species of Xanthophyceae also act as photobionts in lichen symbioses (Fig. 2.8; see Sect. 6.2). Members of the Cryptista are also involved in symbioses with the Radiolaria and Foraminifera (Fig. 2.8). The phenomenon of kleptoplasty is found in the Dinoflagellata where members of Haptista and Cryptista are temporarily exploited for photosynthesis (Figs. 2.8 and 5.47; see Sect. 5.2), and in sacoglossan sea slugs (Holozoa) where chloroplasts taken from members of Xanthophyceae (Vaucheria) or Ulvophyceae (Chlorophyta) serve as temporary chloroplasts (Fig. 2.8; see Sect. 6.1.9). In summary, there are five hotspots known where an otherwise non-photosynthetic lineage receives members of three or more algal lineages as photoautotroph symbiotic partners, i.e., the Holozoa (5 lineages), the fungi by forming lichens (4 lineages; see Sect. 6.2), the Radiolaria (4 lineages), Foraminifera (3 lineages), and Dinoflagellata (3 lineages, through kleptoplasty; see Sect. 5.2) (Fig. 2.8).

2.5

Life histories, Plastid Spread, and Sexual Reproduction in the New Cryptogams

Plastid evolution is an excellent example of understanding the enormous outreach of a single evolutionary event. A single primary endosymbiosis gave rise to the plastids of all the New Cryptogams and tracheophytes (from Cyanobacteria to Archaeplastida, Fig. 2.8). Once the plastids were established in eukaryotes, this primary event led to the further diversification of the oxy-phototroph organisms with eukaryote-eukaryote endosymbioses spreading plastids throughout almost all major phylogenetic lineages of life. These were truly landmarking evolutionary events as the resulting oxy-phototroph species formed the base of the food

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chain for many ecosystems on our planet (Reyes-Prieto and Bhattacharya 2007). All the plastids are incapable of developing de novo. Curiously, no matter if primary or eukaryote-eukaryote endosymbiotic origins, plastids proliferate by their own organelle division, still left somehow independent of nuclear division (mitosis). This can be seen as an obvious mark of ancient evolution in the present-day oxy-phototroph eukaryotes tracing back to that early single primary event. Also, in sexual reproduction, the plastids are typically inherited maternally, i.e., as plastids (or not fully differentiated proplastids) in female gametes and, after fertilization, subsequently undergo division. This makes the endosymbiotic origins of plastids intimately connected with sexual reproduction. Sexual reproduction is a typical feature of eukaryotes. It connects all lineages of the New Cryptogams apart from the Cyanobacteria, where sexuality is lacking. It is the biological pathway for any species to achieve genetic diversity through recombination. The word “sex” is derived, at least partially, from Latin seco, meaning to cleave or to divide. It involves the fusion of gametes (from Greek gamete “a wife” or gametes “a husband”), which leads to the production of a zygote (from Proto-Indo-European Greek, meaning “to join”) and zygote development into a body. Gametes can only develop after fusion with another gamete; they are produced within a dedicated cell or organ, the gametangium. Sexual reproduction leading to genetically different offspring with new genotypes requires gametes of different mating types from genetically different individuals; it is summarized as hologamy. Opposed to it is autogamy, where both gametes are of the same genotype, produced by the same individuum. Their fusion will not result in genetic diversity; it means a proliferation of the same genotype. The fusion of the nuclei of gametes of different mating types results in the diploid zygote. The haploid condition is restored directly in the zygote via meiosis, or the zygote first divides mitotically, producing a generation of diploid cells. There are three principal types of sexual life cycle or life history (Fig. 2.9). Most widespread in eukaryotic algae is the haplontic life cycle (Fig. 2.9a). Here, the zygote serves as a resting stage and is the only diploid stage. No mitotic divisions occur with the zygote. The only mode of division of the zygote is meiosis (reduction division, indicated by “R!”), after which the haploid condition is restored. Thus, the haplontic life cycle is also called “zygotic meiosis.” The result of the first meiotic division is two haploid cells. Completing the second meiotic division (which resembles mitosis) leads to four haploid cells of different mating types. Proliferation by subsequent mitotic divisions of these cells leads to either a population of unicells or a multicellular haploid organism (thallus). Upon sexual reproduction, the haploid cells function as gametes, or the multicellular organism forms gametes by mitosis. After the fusion of the

21

gametes (fertilization, indicated by “K!” for karyogamy, the fusion of nuclei), the zygote is formed. There are numerous examples of haplontic life cycles in the algae, e.g., the green algae Chlamydomonas and Ulothrix (Fig. 4.39). As opposed to the haplontic life cycle, in the diplontic life cycle, all the vegetative cells are diploid, and the gametes are the only haploid cells (Fig. 2.9b). In algae, the diplontic life cycle is restricted to the brown algal order Fucales (e.g., the genus Fucus; Phaeophyceae, Stramenopiles) and the diatoms (Diatomeae, Stramenopiles). The zygote undergoes mitosis, leading to a population of unicellular diatom cells of different mating types or the multicellular thallus of Fucales. Meiotic divisions of a diatom cell or specialized cells on the diploid thallus of Fucales finally result in haploid gametes. Therefore, the diplontic life cycle is also called “gametic meiosis.” Within a short time, the gametes fuse to form the zygote, which immediately starts proliferation by mitosis. An alternation of generation occurs when the zygote does not undergo meiosis but is capable of proliferation by repeated mitosis (Fig 2.9c). This results in a generation of diploid cells forming a diploid thallus. This diploid phase of the life cycle is called the sporophyte. Following meiotic divisions, four haploid cells named spores or meiospores arise, which are not gametes but proliferate by mitosis, forming a haploid thallus. Finally, in that haploid phase, the gametes are formed by mitotic divisions, and, therefore, the haploid phase is called the gametophyte. If both generations, the diploid sporophyte and the haploid gametophyte, look alike, the alternation of generations or phase change is called isomorphic alternation (with the green alga Ulva as an example; Fig. 4.40). Most widespread, however, is the heteromorphic alternation of generations or phase change where the sporophyte is dominant in the life cycle and much larger than the smaller (microscopic) gametophyte. A typical heteromorphic alternation of generations is seen in the brown alga Laminaria (Figs. 2.9c and 5.7) and the green alga Derbesia (Fig. 4.40). The bryophytes also display a heteromorphic alternation of generations. Still, the gametophyte is the dominant phase (Figs. 2.9c, 7.3 and 7.21). In sexual reproduction, the gametes may be more or less differentiated, describing different types of gametes’ mating. Isogamy describes that the gametes of both mating types (+ and −) are structurally indistinguishable. Examples are the motile flagellated isogametes of many flagellated green algae (see Sect. 4.5) and the non-flagellated isogametes of pennate diatoms (Sect. 5.1, Fig. 5.18). Anisogamy involves gametes of both mating types of different sizes and/or behavior. Examples are found in the colonial flagellated green algae (see Sect. 4.5) and the green alga Derbesia (Fig. 4.40). In oogamy, there is one large non-motile gamete which, by definition, is regarded as the egg (female), and one smaller but motile. Among the various lineages of the New

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h haplontic llife cycle

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Fig. 2.9 Principal types of sexual life cycles of the New Cryptogams; a with either the main living phase formed by the haploid organism, or b the diploid organism, or c where both phases alternate

Cryptogams with oogamy, the motile gamete is viewed as male; it may be called sperm (if motile) or spermatium (if immotile) in the algae (see Chaps. 4 and 5), spermatium in lichens (see Sect. 6.2), or spermatozoid in bryophytes (see Chap. 7). Oogamy is a shared feature among various unrelated lineages of the New Cryptogams, e.g., the red algae (Figs. 4.8, 4.9 and 4.10), heterokont algae (Phaeophyceae, Figs. 5.6 and 5.7 Vaucheria, Fig. 5.15), green algae (e.g., Oedogonium Fig. 4.60, and Chara, Figs. 4.80 and 4.82), the bryophytes (Figs. 7.3 and 7.21), and the lichens (Fig. 6.44). The lichens, in turn, developed some types of the mating of gametes of their own. There, genetically different vegetative fungal cells can fuse, called somatogamy, or whole gametangia, then called gametangiogamy (see Sect. 4.2; Box 4.2). In addition, so-called basidiolichens (see Sect. 6.2) pertain to a dikaryotic phase with two unfused nuclei of different mating types within the same cell (functional diploidy), which lasts extensively long after the fusion of cells from a short-lived unicellular stage. In contrast, in so-called ascolichens, the fusion of haploid cells occurs only shortly before meiosis, followed by a rather short dikaryotic stage during the ascus formation (see Sect. 6.2, Fig. 6.44).

References Adl SM, Bass D, Lane CE, Lukeš J, Schoch CL, Smirnov A, Agatha S, Berney C, Brown MW, Burki F, Cárdenas P, Čepička I, Chistyakova L, del Campo J, Dunthorn M, Edvardsen B, Eglit Y, Guillou L, Hampl V, Heiss AA, Hoppenrath M, James TY, Karnkowska A, Karpov S, Kim E, Kolisko M, Kudryavtsev A, Lahr DJG, Lara E, Le Gall L, Lynn DH, Mann DG, Massana R, Mitchell EAD, Morrow C, Park JS, Pawlowski JW, Powell MJ, Richter DJ, Rueckert S, Shadwick L, Shimano S, Spiegel FW, Torruella G, Youssef N, Zlatogursky V, Zhang Q (2019) Revisions to the classification, nomenclature, and diversity of eukaryotes. J Eukaryotic Microbiol 66:4–119. https://doi.org/10.1111/jeu.12691 Bessey CE (1915) The phylogenetic taxonomy of flowering plants. Ann Mo Bot Gard 2(1/2):109–164 Dagan T, Roettger M, Stucken K, Landan G, Koch R, Major P, Gould SB, Goremykin VV, Rippka R, Tandeau de Marsac N, Gugger M, Lockhart PJ, Allen JF, Brune I, Maus I, Pühler A, Martin WF (2012) Genomes of stigonematalean cyanobacteria (Subsection V) and the evolution of oxygenic photosynthesis from prokaryotes to plastids. Genome Biol Evol 5:31–44. https://doi.org/ 10.1093/gbe/evs117 Delwiche CF, Cooper ED (2015) The evolutionary origin of a terrestrial flora. Curr Biol 25:R899–R910. https://doi.org/10.1016/j.cub.2015. 08.029

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de Vries J, Archibald JM (2017) Endosymbiosis: Did plastids evolve from a freshwater cyanobacterium? Curr Biol 27:R103–R105. https://doi.org/10.1016/j.cub.2016.12.006 Eme L, Sharpe SC, Brown MW, Roger AJ (2014) On the age of eukaryotes: evaluating evidence from fossils and molecular clocks. Cold Spring Harbor Perspect Biol 6:a016139–a016139. https://doi. org/10.1101/cshperspect.a016139 Herrero A, Stavans J, Flores E (2016) The multicellular nature of filamentous heterocyst-forming cyanobacteria. FEMS Microbiol Rev 40:831–854. https://doi.org/10.1093/femsre/fuw029 Hirano T, Tanidokoro K, Shimizu Y, Kawarabayasi Y, Ohshima T, Sato M, Tadano S, Ishikawa H, Takio S, Takechi K, Takano H (2016) Moss chloroplasts are surrounded by a peptidoglycan wall containing D-amino acids. The Plant Cell 28:1521–1532. https:// doi.org/10.1105/tpc.16.00104 Huang CY, Grünheit N, Ahmadinejad N, Timmis JN, Martin W (2005) Mutational decay and age of chloroplast and mitochondrial genomes transferred recently to angiosperm nuclear chromosomes. Plant Physiol 138:1723–1733. https://doi.org/10.1104/pp.105. 060327 Jackson C, Knoll AH, Chan CX, Verbruggen H (2018) Plastid phylogenomics with broad taxon sampling further elucidates the distinct evolutionary origins and timing of secondary green plastids. Sci Rep 8:1523. https://doi.org/10.1038/s41598-017-18805-w Lewis LR, Ickert-Bond SM, Biersma EM, Convey P, Goffinet B, Hassel K, Kruijer HJD, La Farge C, Metzgar J, Stech M,

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Villarreal JC, McDaniel SF (2017) Future directions and priorities for arctic bryophyte research. Arctic Sci 3:475–497. https://doi.org/ 10.1146/annurev.genet.41.110306.130134 Martin WF, Garg S, Zimorski V (2015) Endosymbiotic theories for eukaryote origin. Philos Trans R Soc Lond, B, Biol Sci 370:20140330–18. https://doi.org/10.1098/rstb.2014.0330 Michalovova M, Vyskot B, Kejnovsky E (2013) Analysis of plastid and mitochondrial DNA insertions in the nucleus (NUPTs and NUMTs) of six plant species: size, relative age and chromosomal localization. Heredity 111:314–320. https://doi.org/10.1038/hdy.2013.51 Ponce-Toledo RI, Deschamps P, López-García P, Zivanovic Y, Benzerara K, Moreira D (2017) An early-branching freshwater cyanobacterium at the origin of plastids. Curr Biol. 27:386–391. https://doi.org/10.1016/j.cub.2016.11.056 Ricchetti M, Fairhead C, Dujon B (1999) Mitochondrial DNA repairs double-strand breaks in yeast chromosomes. Nature 402:96–100. https://doi.org/10.1038/47076 Reyes-Prieto A, Bhattacharya D (2007) Phylogeny of nuclear-encoded plastid-targeted proteins supports an early divergence of glaucophytes within plantae. Mol Biol Evol 24:2358–2361. https://doi.org/ 10.1146/annurev.genet.41.110306.130134 Sánchez-Baracaldo P, Raven JA, Pisani D, Knoll AH (2017) Early photosynthetic eukaryotes inhabited lowsalinity habitats. Proc Natl Acad Sci 114(37):E7737–E7745. https://doi.org/10.1073/pnas. 1620089114

3

Cyanobacteria/Blue-Green Algae Burkhard Büdel

Contents 3.1 Short History of Cyanobacterial Research....................................................................................... 26 3.2 Origin of Cyanobacteria ..................................................................................................................... 28 3.2.1 Cyanobacteria and the Origin of Plastids .................................................................................. 30 3.3 Structure and Function....................................................................................................................... 30 3.3.1 Cell Wall..................................................................................................................................... 31 3.3.2 Cytoplasm ................................................................................................................................... 33 3.4 Survival and Cell Specialization ........................................................................................................ 3.4.1 Heterocytes ................................................................................................................................. 3.4.2 Akinetes ...................................................................................................................................... 3.4.3 Necridic Cells ............................................................................................................................. 3.4.4 Hormogonia ................................................................................................................................

43 44 47 48 48

3.5 Reproduction, Life Cycle .................................................................................................................... 48 3.5.1 Unicellular Cyanobacteria .......................................................................................................... 48 3.5.2 Filamentous Cyanobacteria ........................................................................................................ 49 3.6 Phylogeny and Diversity ..................................................................................................................... 51 3.6.1 Taxonomy and Systematics........................................................................................................ 52 3.6.2 Biogeography.............................................................................................................................. 68 3.7 Ecology.................................................................................................................................................. 3.7.1 Rock and Soil ............................................................................................................................. 3.7.2 Epiphytic..................................................................................................................................... 3.7.3 Freshwater and Marine Habitats ................................................................................................ 3.7.4 Eco-Physiology ...........................................................................................................................

69 70 79 79 87

References ..................................................................................................................................................... 91

Dating back to the earliest period of geological time (Archean), cyanobacteria are the oldest known photoautotrophic organisms that release oxygen during their photosynthesis (= oxygenic photosynthesis). This ability, likely

B. Büdel (&) RPTU Kaiserslautern, Department of Biology, Erwin-Schrödinger-Str. 13, 67663 Kaiserslautern, Germany e-mail: [email protected]

evolved roughly 3000 million years ago (Mya), related to the “Great oxidation event” (GOE) and forever altered the Earth’s environment, both in water and on land (Schopf 2014; Schirrmeister et al. 2015, 2016). With the oxidation of water at photosystem II, protons (hydrogen) and electrons are gained and used to form energy-rich carbohydrates, the solar fuel of living cells. As a byproduct of water scission, the resulting oxygen combines with the molecule O2 that is discharged to the environment. This provided the highly

© Der/die Autor(en), exklusiv lizenziert durch Springer-Verlag GmbH, DE, ein Teil von Springer Nature 2024 B. Büdel et al. (eds.), Biology of Algae, Lichens and Bryophytes, https://doi.org/10.1007/978-3-662-65712-6_3

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3.1

Fig. 3.1 Aliterella sp, colony with baeocytes; courtesy of Patrick Jung

reactive and biologically useable molecular oxygen required for aerobic respiration, a decidedly more efficient energy-generating process than its anaerobic (fermentative) precursors (Schopf 1999, 2014). Having been the dominant photosynthesizers on the land surface, cyanobacteria were also the drivers of initial soil formation (e.g. Lalonde and Konhauser 2015; Mergelov et al. 2018). However, the evolutionary timing of photosynthetic oxygen release is poorly understood. There is an increasingly observed disconnection between the presence of O2 during weathering as early as 3000 Mya and its atmospheric accumulation 500 million years later. It is suggested that local O2 production and immediate consumption in surface-bound microbial ecosystems (e.g. early biocrusts on rock and soil; Fig. 3.2c– e) at profound disequilibrium conditions can be the explanation for this delay (Lalonde and Konhauser 2015) (Fig. 3.1).

Short History of Cyanobacterial Research

The early perception of cyanobacteria is perhaps best illustrated with the macroscopic terrestrial species of the widespread genus Nostoc. In Europe, the first reports of Nostoc by THOMAS DE CANTIPRATO (1186–1270) and KONRAD VON MEGENBERG (1309–1378) date back to the Middle Ages and refer unmistakably to the species N. commune, as also can be inferred from many colloquial names (German: Sternschnuppen, English: star snot) in several European countries (Mollenhauer 1985a, b). These colloquial names originate from the supposed celestial derivation, because the large thalli all of a sudden become visible when soaked in water after rainfall. The celestial derivation, as well as the belief that N. commune thalli can serve as the philosopher’s stone, led PARACELSUS (THEOPHRASTUS BOMBASTUS VON HOHENHEIM, 1493?–1541) to include it in his studies and he was also the first who named it “Nostoc” (Mollenhauer 1985b). An even earlier historic report was found by Hirose (1962), with the description of a macroscopic Nostoc species from Asia in a collection of poems from the year 730. According to modern taxonomy, this species can be identified as N. verrucosum. The first scientific studies were those of the Swiss preacher JEAN PIERRE VAUCHER (1763–1841), who carefully described the life history of the genus Nostoc, and thus prepared the field for further studies. The early scientists JEAN-HENRI FABRE (1823–1915), GUSTAVE ADOLPHE THURET (1817–1875), ÉDOUARDE BORNET (1828–1911), CHARLES FLAHAULT (1852– 1935), and ÉDOUARD DE GLINKA-JANCZEWSKI (1846–1918) all contributed to the unraveling of the developmental stages of the genus Nostoc (Mollenhauer 1986a). Even today, Nostoc species play an important ecological role in terrestrial habitats (see referring chapters in this book). After these early pioneers of cyanobacterial research, the number of scientists devoted to cyanobacteria is many and I will not be going into detail, but I will refer the interested reader to specific publications such as, for example, Mollenhauer (1985a), or the most recent summary of Donald E. Bryant (2014). In cyanobacterial research, Bryant (2014) distinguished between the Dark Ages (1800 to 1950), the Middle Ages (1950–1979), the Renaissance (1980–1995), the Age of Enlightenment (1996 to present), and the Post-Modern Era

Fig. 3.2 Fossil cyanobacteria and microbialite. a + b from the Bitter Springs chert of central Australia, dated roughly 850 million years old, c courtesy of Williams Schopf; a Chroococcalean form and b the filamentous fossil genus Palaeolyngbya, courtesy of UMCP Berkeley; c microbialite from the Laguna Negra, a hypersaline lake of the Laguna Verde complex in the Puna region of Argentina (4000 m above sea level); cross section showing the concentrically laminated structure of the microbialite, nucleus of origin in the center; from Buongiorno et al. (2019), with permission of Willey, Geobiology; d cross section of the surface of a modern aragonitic stromatolite, 1 = Calothrix sp., filamentous cyanobacterium with basal heterocytes, 2 = unicellular cyanobacteria at the base of the biofilm, 3 = newly formed aragonite, 4 = mature stromatolite with organic remains; Walker Lake, Nevada, courtesy of Joachim Reitner; e lacustrine stromatolite formation, Municipio Othón, Quintana Roo, Yucatán peninsula, Mexico, courtesy of Ulrike Büdel

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Cyanobacteria/Blue-Green Algae

27

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(the future). The “eras” were largely defined by the prevailing methodological approaches used to study cyanobacteria. The first 150 years (“Dark Ages”) were basically marked by field and light microscopic observations accompanied by enrichment methods and physiological studies. The “Middle Ages” are defined by the author as the period when the first cyanobacterial pure cultures became available, but most research still relied on physiological and biochemical approaches. The major innovation in research methods of this “era” was the use of transmission electron microscopy; it revealed, not only the prokaryotic nature of cyanobacteria, but also it was an important tool for studying their light harvesting complexes. The “Renaissance era” is characterized by the advent of molecular biological and genetic methods. Molecular methods were also applied to taxonomical and phylogenetical research and the use of 16S rRNA became commonplace. The “Age of Enlightenment” is certainly an important milestone, and a few things impacted global knowledge of microorganisms in the same manner as the sequencing of their genomes did. Today, the genome of roughly 200 cyanobacterial species is fully sequenced. Fluorescence-based microscopic methods allowed new studies into the diverse processes in cyanobacteria; the visualization of proteins and study into their localization have been developed in the last 20 years and markedly increased our knowledge of the cell biology of cyanobacteria (Bryant 2014). Very recently, a redefinition of the cyanobacteria has been proposed based on phylogenomic analysis of distantly related non-phototrophic lineages. In this work, cyanobacteria are defined as “Organisms in the domain bacteria able to carry out oxygenic photosynthesis with water as an electron donor and to reduce carbon dioxide as a source of carbon, or those secondarily evolved from such organisms” (Garcia-Pichel et al. 2019a).

3.2

Origin of Cyanobacteria

Inferred from morphological similarities, the scarce fossil record (Fig. 3.2a, b) suggests an age of about 3500 million years for the cyanobacterial lineage (Schopf 2014). In addition, there is strong evidence of cyanobacteria being the oldest microorganisms with oxygenic photosynthesis and are the cause for a sharp rise in atmospheric oxygen about 2450– 2320 Mya (Rasmussen et al. 2008; Schirrmeister et al. 2015, 2016). The rise of oxygen was enabled by the evolution of oxygenic photosynthesis in the ancestors of cyanobacteria (Fig. 3.3). However, there is a debate on the origin and phylogenetic relationship of cyanobacteria with other,

non-photosynthetic bacteria and related to that, the origin of oxygenic photosynthesis. Recent molecular studies, using genes related to photosynthesis and aerobic respiration, have revealed the existence of two bacterial clades, the Vampirovibrionia (former Melainabacteria) and Sericytochromatia. Both groups are non-photosynthetic with the Vampirovibrionia being the direct sister group to cyanobacteria and both are closely related to the Sericytochromatia. Both are proposed as new classes and, together with the class Oxyphotobacteria (the former cyanobacteria), forming the newly proposed phylum Cyanobacteria (Soo et al. 2014, 2017, 2019). Three main theories are dealt with in the debate about the timing of the origin of oxygenic photosynthesis related to the three classes (for a recent comprehensive discussion, see Soo et al. 2019): (1) The fusion model assumes that reaction center I (RCI) and reaction center II (RCII) were obtained via horizontal gene transfer from a non-photosynthetic cyanobacterial ancestor of two different anoxyphototrophic bacteria (Hohmann-Marriott and Blankenship 2011). (2) The selective loss model hypothesizes a single unknown photosynthetic ancestor, having both reaction centers. Subsequently, all segregated photosynthetic lineages lost either RCI or RCII except the Cyanobacteria (Olson and Pierson 1987). (3) The export model hypothesis is based on the comparison of 15 complete cyanobacterial genome sequences. The fact that the anoxygenic phototrophs had only a few components of the cyanobacterial photosynthetic machinery suggests that photosynthesis originated in the cyanobacterial lineage. According to the hypothesis, anoxygenic photosynthetic lineages acquired their reaction centers via horizontal gene transfer from anoxygenic ancestors (see also Chap. 2, Fig. 2.1). Molecular clock analyses of the three most ancient phototrophic groups, cyanobacteria (phylum Cyanobacteriota), green non-sulfur bacteria (phylum Chloroflexi), and green sulfur bacteria (phylum Chlorobi) let to the conclusion that the cyanobacteria most probably did not receive their RCI from green sulfur bacteria. However, they might have been the donor or recipient of RCII proteins from the green non-sulfur bacteria before the rise of atmospheric O2 (Magnabosco et al. 2018). On the other hand, a second molecular clock analysis came to the conclusion that in green non-sulfur bacteria, phototrophy was acquired markedly after the Great Oxygenation Event and therefore could not have donated photosynthetic genes to the ancestor of the cyanobacteria (Shih et al. 2017a).

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Fig. 3.3 Geological timescale and the great oxidation event as inferred from recent literature with and without Vampirovibrionia (former Melainabacteria) impact on timing of the molecular clock method. Modified from Schopf (2014) with additions taken from Yoon et al. (2004), Tomitani et al. (2006), Parfrey et al. (2011), McFadden (2014), Archibald (2015), Schirrmeister et al. (2011, 2015), Neustupa (2015a, b), Shih et al. (2017a, b), Garcia-Pichel et al. (2019b), and Cardona et al. (2019)

According to an expanded analysis, the crown group of the oxyphotoautotrophic bacteria evolved only 2000 Mya, well after the rise of atmospheric dioxygen (Shih et al. 2017b). The authors estimate the divergence between Cyanobacteria and Vampirovibrionia at ca. 2500–2600 Mya. Furthermore, if oxygenic photosynthesis would be a joint new feature (synapomorphy) of the oxyphotoautotrophic bacteria, this would then mark the upper limit for the origin of oxygenic photosynthesis (Fig. 3.3). However, a very recent combined study of fossil records and relaxed molecular clock models sets the minimum age for the evolutionary advent of scytonemin, an important indol-phenolic UV-A sunscreen of cyanobacteria, at 2100 ± 300 million years. In the same analysis, the advent of cyanobacteria and thus the appearance of oxygenic photosynthesis was set at 3600 ± 200 million before the present (Garcia-Pichel et al. 2019a, b). This view is supported by new results, suggesting a homodimeric photosystem with sufficient oxidizing power to split water. This apparently had already appeared in the early Archean about 1000 million years before (3220 Mya) in the most recent common ancestor of all described cyanobacteria capable of oxygenic photosynthesis, and well

before the diversification of some of the known groups of anoxygenic photosynthetic bacteria (Cardona et al. 2019). The oldest fossil evidence of cyanobacteria-like organisms is derived from stromatolites dating back to roughly 3500 Mya. Stromatolites are laminated microbialites, biogenic rocks with a concentrically laminated structure, caused by alternating sediment precipitation and growth periods of the cyanobacteria-like oxygenic bacteria. Each growth period is followed by a period of sediment trapping and/or active precipitation of calcium carbonate due to the photosynthetic activity of the microorganisms (see Box 3.2). Comparable processes can still be observed today, in actively growing microbialites formed under the participation of cyanobacteria (often Rivularia-like). Microbialites were not only very important biogenic limestone formations in the Precambrian and Phanerozoic, but still can be found actively growing in often extreme, marine, and freshwater habitats such as, for example, those of Laguna Negra in the high Andes of Argentina where they form oncolites (a special form of microbialites with concentric lamination) in the size of up to 30 cm (Fig. 3.2c). They are among the most sensitive paleoenvironmental indicators and are commonly

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used in reconstructing lake evolution as environmental fluctuations are recorded in sedimentary archives of lacustrine depositional systems (Buongiorno et al. 2019).

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plant-like organisms developed, the Chloroplastida, Rhodophyceae, and the minor group Glaucophyta, all are summarized under the group Archaeplastida (Adl et al. 2012, 2018).

3.2.1 Cyanobacteria and the Origin of Plastids

3.3 Photosynthetic eukaryotes are the product of an endosymbiotic event between a eukaryotic host and a cyanobacterium that became today’s plastid (see also Chap. 2, this book). A recent phylogenomic study suggests (Fig. 3.4) that the closest relative of plastids among extant cyanobacteria is the recently discovered freshwater-dwelling Gloeomargarita lithophora (Couradeau et al. 2012; Ponce-Toledo et al. 2017; de Vries and Archibald 2017). After discovering the universality of self-reproducing chromatophores (earlier term for plastids), ANDREAS FRANZ WILHELM SCHIMPER (1856–1901) published, in a footnote, the theory of the origin of plastids as probably being based on an association of a colorless cell with a chlorophyll-containing organism (Schimper 1883). In September 1905, a contribution by KONSTANTIN SERGEEVIČ MEREŽKOVSKY (1855–1921) appeared, where the author argued strictly against the increasingly popular opinion that the chromatophores are nothing but normal organelles of the cell (Merežkovsky 1905). MEREŽKOVSKY argued that the chromatophores could not be organelles as they are exclusively self-reproducing and independent from the cell. Based on that, he postulated (translated from German by the author): “If chromatophores are not organs and never have been, then the only possibility remaining is to consider these entities as organisms, as symbionts”. Today it is well-established knowledge that via primary endosymbiosis, the three major lineages of Fig. 3.4 Suggested scenario of the endosymbiogenetic origin of plastids (from de Vries and Archibald 2017, Current Biology, with permission from Elsevier), (see also Fig. 2.1)

Structure and Function

Cyanobacterial cells are extraordinarily large compared to bacteria or archaea. They range from 0.5 lm to 50–100 lm in width and multicellular filaments can form thalli several centimeters big. A number of species develop macroscopic colonies of up to 30 cm; for example, the heaviest macroscopic cyanobacterial colony reported is a ball-shaped Nostoc pruniforme colony with a fresh weight of 3 kg (Dodds and Castenholz 1988). Vegetative cells are not specialized and unicellular cyanobacteria usually have cells with a spherical, bacilloid, or fusiform shape (Fig. 3.5). In the bacterial world, cyanobacteria are among the few representatives exhibiting complex morphologies. Many are multicellular and grow as branched or non-branched filaments of cells. Within those filaments, some cells can differentiate to carry out specialized functions. This cell differentiation process of cyanobacteria leads to a compartmentalized function and demands intercellular communication. There is strong evidence that the supracellular structure of cyanobacterial filaments is related to basic principles governing the process of, for example, heterocyte differentiation (Flores and Herrero 2010). The cells of filamentous cyanobacteria range from discoid to barrel-shaped. A number of filamentous cyanobacterial species are pleomorphic, meaning that they can vary in size and shape. The well-known species Starria zimbabweensis

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Fig. 3.5 Cross fracture of a colony of the unicellular cyanobacterium Chroococcidiopsis sp., an isolated lichen photobiont. Prepared by freeze fracturing, covered in the frozen state by carbon, and then viewed in a transmissionelectron-microscope, artificially colored

31 Cell wall layers Glykogen granules

View from outside on top of cytoplasm

Thylakoids

EPS-layers (sheath envelope)

Nucleoplasm

Carboxysomes Chromatoplasm (thylakoid region) Polyphosphate vesicle Glykogen granules

(Fig. 3.36i, j) presents a u-shaped or sometimes even a triradiate symmetry in cross sections. Not all filamentous forms necessarily imply a functional integration of cells into a truly multicellular organism. However, in the heterocytous group, and some oscillatorians, truly multicellular organisms with all attributes required for such a distinction are realized (Garcia-Pichel 2009; Herrero et al. 2016). The cyanobacterial filament is composed of the “trichome” which refers to the cyanobacterial cell arrangement only plus the sheath envelope.

3.3.1 Cell Wall The cyanobacterial cell wall outside the cell membrane is composed of a peptidoglycan layer with a thickness ranging from 6 to 12 nm in unicellular taxa, reaching 12 to 36 nm in filamentous species and more than 700 nm in large Oscillatoria-species (Hoiczyk and Hansel 2000). The peptidoglycan layer is composed of glucosamine, muramic acid, diaminopimelic acid, glutamate, alanine, and glycine in different proportions (Hahn and Schleiff 2014). On top of the peptidoglycan layer follows an outer membrane of the lipopolysaccharide-type. In contrast to the peptidoglycan layer, the outer membrane does not enter the septum between neighboring cells but forms a continuity in filamentous cyanobacteria. Between the peptidoglycan and the cytoplasmic layers, and also the peptidoglycan and the outer membrane, a periplasmic space is inserted (Figs. 3.6, 3.18,

1 µm

3.20). In filamentous species, it is thought that the periplasmic space might connect heterocytes and vegetative cells and is the space where the transfer of reduced carbon from vegetative cells to heterocytes, and fixed nitrogen in the reverse direction, takes place (Wolk et al. 1974). Three decades later, the exchange of solutes via the periplasmic space was demonstrated for the filamentous genus Anabaena (Mariscal et al. 2007). It is suggested that in the periplasmic space, a regulatory peptide is present to repress heterocyte differentiation of adjacent cells and ensure a correct spacing of heterocytes of approximately 10–20 vegetative cells between heterocytes (Yoon and Golden 1998). Cell–cell connections of filamentous cyanobacteria via the so-called cyanobacterial septal junctions (formerly named microplasmodesmata; Giddings and Staehelin 1981) allow an intercellular molecular diffusion of substances such as reduced carbon or combined nitrogen between cells (Nieves-Morión et al. 2017). The cyanobacterial septal junctions, however, have so far only been detected in the filamentous heterocyte-forming Nostocales, not in the Oscillatoriales (Mullineaux et al. 2008). In a number of unicellular and some filamentous cyanobacterial species, the wall is completed by an S-layer on top of the outer membrane (Figs. 3.6, 3.7). This is a two-dimensional crystalline array formed by a single species of (glyco-)protein covering the entire cell surface (Hoiczyk and Hansel 2000; Šmarda et al. 2002; Zhu et al. 2017). A specific gliding motility mode of filamentous cyanobacteria was found in the cyanobacterium Phormidium

32 Fig. 3.6 Cyanobacterial cell wall (according to Sleytr et al. 1996; Hahn and Schleiff 2014). Please note that not all cyanobacteria have an S-layer

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S-layer

Lipopolysaccharide Outer membrane

Cell wall

Phospholipid Lipoprotein Peptidoglycan layer Cytoplasmic membrane

unciatum, where a specific pore complex, together with the S-layer, provides the necessary structure for motility of filamentous cyanobacteria (Fig. 3.6; Hoiczyk and Baumeister 1998). S-layers of multiple different structures have been found in several unicellular and filamentous cyanobacteria (Fig. 3.7) but are not so far known from members of the Nostocales (Hahn and Schleiff 2014). Many cyanobacteria produce extracellular sheaths that contain visible and UV-light-protecting pigments like scytonemin and mycosporine-amino acids (= MAAs) as a response to environmental conditions (Fig. 3.8; Garcia-Pichel and Castenholz 1991; Böhm et al. 1995). While scytonemin is restricted to the sheath, MAAs are also

found in the cytoplasm. In older literature, the sheath is also referred to as “slime”. It is composed of high-molecular-mass heteropolysaccharides, proteins, nucleic acids, and lipids and should therefore be referred to as extracellular polymeric substances (= EPS; Flemming and Wingender 2010; Fig. 3.5). The composition of the cyanobacterial EPS of the major taxonomic groups, their structural complexity, the biosynthetic pathways, and the genes involved are reviewed in Pereira et al. (2009). Cyanobacterial sheaths, especially those of terrestrial taxa, can be vividly dyed with colors ranging from yellow, red, and violet to blue (Fig. 3.8). In some species, the color varies with the pH of the substratum or might also vary

Fig. 3.7 a S-layer with a striated pattern of the unicellular cyanobacterium Chroococcidiopsis sp.; EPS = extracellular polymeric substances (sheath), cm = cytoplasmic membrane, om = outer membrane, arrow indicates direction of shadowing (courtesy of Archives of Microbiology); b model of the S-layer illustrating the striated pattern of the 2–3 nm thick ribbons that are interwoven to a patchwork-like leaflet (courtesy of Archives of Microbiology; c model of the supramolecular structure of the pore complex, the S-layer, and the gliding elements of Phormidium unciatum (recolored, from Hoiczyk and Baumeister 1998, Current Biology, with permission from the authors and Elsevier)

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Fig. 3.8 a Laminated sheath of the cyanobacterium Petalonema alata from a rock surface of the Swiss Alps. Note the funnel-shaped interlaced sheath layers with the inner layers having the strongest pigmentation by yellow scytonemin; b Stigonema ocellatum from biological soil crust, Queensland, Australia, with red sheath (courtesy of W. Williams); c red sheath of Gloeocapsa sanguinea, granite rock surface from Inselberg, Venezuela; d yellow sheath of G. kuetzingiana, calcareous rock surface, Poland; e blue sheath of G. sp., calcareous rock surface, France; f violet sheath of G. cf. sanguinea, granite rock surface, Venezuela

during the life cycle. When kept in culture under low light conditions, the sheath color often disappears, a feature that can also be observed in nature when cyanobacteria are not directly exposed to sunlight (e.g. in caves or in deeper layers of mats). It has been shown by various researchers that the sheath color is an effective sunscreen, and it reduces the amount of light reaching the cell content and is an effective and photo-stable ultraviolet shield (Garcia-Pichel and Castenholz 1991; Proteau et al. 1993). The yellow to yellow-brownish pigment scytonemin is an indol alkaloid strongly absorbent in the spectral region 325–425 nm (UV-A-violet-blue) but also has major absorption in the UV-C, kmax = 250 nm and UV-B, 280–320 nm region (Proteau et al. 1993). Until now, seven different forms of scytonemins have been reported. The UV sunscreen function of scytonemins is supported by intracellular substances known as mycosporine-like amino acids (MAAs). Due to their prominent UV-filtering properties, these cyanobacterial metabolites are presently under discussion as a source of UV-sunscreens and moisturizers in human care products (Derikvand et al. 2017). Multiple environmental signals influence the scytonemin and MAA synthesis, and the regulation of the induction of these UV screening compounds is a part of a complex stress response pathway. In addition to the photo-protective function, scytonemin and MAAs also play important roles as antioxidant molecules, compatible solutes, and nitrogen reservoirs and act in defense against temperature, desiccation, and other stress conditions (Pathak et al. 2019). The functional role of the cyanobacterial EPS is more than only acting as a light and UV-radiation screen. Additionally, it supports the colonization process of exposed

terrestrial surfaces on rock and soil but also plays an important role in resurrection after desiccation. The EPS of the filamentous Leptolyngbya ohadii was reported to stabilize bare sandy substrates occupied by them, thus preparing the ground for successful colonization (Mugnai et al. 2018). The resurrection process after desiccation of the green alga Chlorella sp., a species normally not able to revive in axenic cultures, was possible for the alga when provided with EPS or its main sugars constituents from the cyanobacterium Leptolyngbya sp. Nevertheless, besides this positive effect for Chlorella sp., it was simultaneously inhibited in growth (Kedem et al. 2020). In summary, there are clear indications on the eco-physiological importance of the EPS in the hydration-desiccation process of cyanobacteria (see also Sect. 3.7.4).

3.3.2 Cytoplasm Using a reasonably good quality light microscope, the cell content of cyanobacteria can often be separated into a darker region containing thylakoids in different arrangements, mostly in the cell periphery, and a somehow brighter region free of thylakoids named the nucleoplasm that hosts the cellular DNA (Fig. 3.5). The genome size of cyanobacteria ranges between 1.4 and 9.1 Mbp and has a G + C content of 31–63%; the number of protein-coding genes is between 1214 and 8446 and has a coding nucleotide proportion of 52–94%. The evolution of the genome size in cyanobacteria apparently involves a mix of gains and losses in the morphologically complex cyanobacteria, while in unicellular cyanobacteria a single event of reduction was evident. An

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ancestral character reconstruction suggests the most recent ancestor of cyanobacteria had a genome size of roughly 4.5 Mbp with 1678 to 3291 protein-coding genes, 4–6% of which are unique to cyanobacteria (Larsson et al. 2011).

3.3.2.1 Thylakoids Thylakoids are composed of a double-membrane and carry photosystem I and II with the accessory light harvesting pigments (Figs. 3.5, 3.9, 3.20), the ATP-synthase, the cytochrome complex, and the NDH1-complex. The only exception is the cyanobacterial genus Gloeobacter, which does not develop thylakoids. In this genus, the photosynthetic apparatus is located along the plasma membrane. Cyanobacterial photosynthesis, oxygenic photosynthesis, uses chlorophylla to convert visible light into chemical energy. The photochemical active pigments are the two photosystems I (PS I) and II (PS II), and the major light absorbing pigments are chlorophylla, more rarely chlorophyllb (Prochloron, Prochlorococcus, Prochlorothrix), or, in the marine cyanobacterium Acaryochloris marina chlorophylld, allowing this species to use far-red light. Modern high-resolution cryo-electron microscopy revealed the structure of PS I. Embedded in the thylakoid membrane of cyanobacteria, the PS I structure is either trimeric as in most cyanobacteria, tetrameric in heterocyte-forming cyanobacteria and their relatives (Fig. 3.8a), or, more rarely monoand dimeric, in contrast to the strictly monomeric form of PS I in plants and algae (e.g. Kato et al. 2019; Li et al. 2019a, b; Semchonok et al. 2021). One monomer is the minimal functional unit of cyanobacterial PSI and comprises 12 protein subunits (PsaA, PsaB, PsaC, PsaD, PsaE, PsaF, PsaI, PsaJ, PsaK, PsaL, PsaM, and PsaX), 96 chlorophylls, more than 20 carotenoids, four structural lipids, and three [Fe4S4] iron-sulfur clusters (Rögner et al. 1990). Recently, the presence of chlorophyllf was reported from a number of cyanobacteria. The first cyanobacterial chlorophyllf was detected when an extract of actively living stromatolites from Hamelin pool, Shark Bay, Western Australia, was made. It was assignable to a new filamentous cyanobacterium of the Oscillatoriales-group, Halomicronema hongdechloris (Chen et al. 2010, 2012). Later, another oscillatoralean cyanobacterium, Leptolyngbya

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sp. was found to produce chlorophyllf when grown in far-red light (Gan et al. 2014), as well as a unicellular cyanobacterium that could not be properly assigned, neither to “Aphanocapsa muscicola”, to whom it had a 97.5% gene sequence identity, nor to Acaryochloris sp. (Miyashita et al. 2014). When grown under 750 nm far-red light, the unicellular cyanobacterium Chroococcidiopsis thermalis contained *90% chlorophylla, *10% chlorophyllf, and 45 °C and those that are able to grow at even higher temperatures, >80 °C are referred to as hyperthermophile. Synechococcus sp. from the Lower Geyser Basin of the Yellowstone National Park, US, holds the upper temperature record for photosynthesis (  70 °C), whereas species of Calothrix, Nostoc, and Scytonema are restricted to the more moderate surrounding of hot springs and grow at  45–50 °C (Castenholz and Garcia-Pichel 2012). Generally, there are no hyperthermophile cyanobacteria, and all so far known hot spring cyanobacteria are considered as being thermophile organisms. Thermophilic cyanobacteria are present in two independent lineages of phylogenetic trees based on the small subunit of rDNA: the Synechococcales (with affinity to Gloeobacter) and the Nostocales (Bhattacharya et al. 1999).

3.7.3.3 Marine Plankton The plankton of the open oceans (71% of the surface of the Earth) generally has a high proportion of cyanobacteria in terms of biomass, oscillating in presence over the year. Unicellular single-celled cyanobacteria are the most abundant, with species of the genera Synechococcus, Prochlorococcus, Chroococcus, Synechocystis, and Crocosphaera belonging to the fraction picoplankton. In terms of primary

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production and biomass, they often account for more than 50% of the biomass. Common filamentous forms are Oscillatoria, Trichodesmium, Anabaena, and Nodularia. For a more complete overview on marine cyanobacterial plankton, please refer to the review of Pearl (2012). The picoplankton species in particular express very limited numbers of morphological features, making a sound species determination and often also species definition (!) extremely difficult and unsafe. The problematic situation, in periods of rapidly declining cyanobacteria taxonomy experts and due to the inherent morphology problems, might be illustrated by a very recent study. For more than 20 years, it was observed that in a shallow, eutrophic brackish lagoon of the Baltic Sea, the phytoplankton was dominated by an Aphanothece-like morphospecies. However, a study based on 16S rRNA gene phylogeny of clone sequences and isolates indicated the dominance of a Cyanobium species. This difference between morphologically and genetically based species identifications is in accordance with literature that shows the same pattern: morphologically Aphanothece-like species are abundant in eutrophic shallow lagoons, and genetically Cyanobium is found in similar habitats. This discrepancy is found worldwide in the literature on freshand brackish-water habitats and there is strong evidence that most Aphanothece-like morphospecies may be, genetically, members of Cyanobium. Other planktonic cyanobacteria found in the Baltic Sea lagoon were Aphanothece c.f. nidulans, Aphanocapsa incerta, Chroococcus vacuolatus, Cyanodictyon planctonicum, Gloeothece sp., Merismopedia sp., Planktolyngbya sp., Rhabdoderma cf. linearis, Snowella sp., Synechocystis cf. diploca, and Woronichinia sp. (Fig. 3.50; Albrecht et al. 2017). There is an urgent need for proper combined taxonomic revisions, including old-type species/specimens and modern methodology. Cyanobacterial harmful algal blooms have also been reported from the marine environment worldwide. A quite well-documented and regularly occurring marine bloom is that dominated by the filamentous, heterocyte, and akinete-forming cyanobacterium Nodularia spumigena in the Baltic Sea that can nicely be observed via satellite (Fig. 3.51a). The total area covered by this cyanobacterial bloom reached more than 62 000 km−2 in 1992 (Kahru et al. 1994). Recently, oscillations with a period of *3 years have been observed, a high concentration year is generally followed by one or two low concentration years of blooms (often Nodularia spumigena dominated) in the Baltic Sea (Fig. 3.51b). The reason for this seasonality remains unclear and the authors assume that the oscillations might be intrinsic to the marine system (Kahru et al. 2018).

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Fig. 3.50 Marine cyanobacterial plankton; a Gloeothece sp.; b Snowella sp.; c Cyanodictyon sp.; d Woronichinia sp.; e Merismopedia sp.; f Planktolyngbya contorta. Courtesy of Rhena Schumann, Rostock with permission of Frontiers in Microbiology

Fig. 3.51 Blooming cyanobacteria; a Satellite view of the Baltic Sea (July 8th 2005) showing the bright green bloom of cyanobacteria south and east of Gotland (white arrow); courtesy of Mati Kahru, produced using data provided by NASA; b Nodularia spumigena, coastal sediment, Schiermonikoog island, the Netherlands, (CCY Strain 1407), note the Akinetes (arrow) and heterocytes; courtesy of Michele Grego, NIOZ, the Netherlands.7.4 Eco-physiology

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Box 3.4: Toxicity of Cyanobacteria Among the cyanobacteria, especially the bloom-forming ones, a number of taxa exist that can be harmful to animal and human populations due to their production of secondary metabolites that are collectively known as cyanotoxins. Such toxic blooms have led to deaths of wild and domestic animals and are a health risk for human beings all over the world via recreational or drinking water (Kuiper-Goodman et al. 1999). Three major groups of toxins are distinguished: the hepatotoxins that imply chemical-driven liver damage, neurotoxins that are poisonous or destructive to the nerve system, and cytotoxins that are toxic to cells. The heptaotoxins microcystine and nodularin are cyclic peptides and are produced most commonly by cyanobacteria of the genera Microcystis, Dolichospermum, Planktothrix, Phormidium, and Nodularia spumigena; more rarely some Nostoc species can also produce toxins. Hepatotoxins contain uncommon amino acids like dehydroalanine, which covalently bond to and inhibit the protein-phosphatases (Mackintosh et al. 1990). Critical exposure may lead to liver failure or even death (Chorus and Bartram 1999). Contrastingly, the hepatotoxic cylindrospermopsin is an alkaloid and is also toxic to kidneys. It is produced by the cyanobacterium Cylindrospermopsis raciborski when bloom-forming. The neutrotoxin anatoxin-a is an alkaloid that acts by disrupting the signal transmission between neurons and muscles, which can lead to death by respiratory arrest. They are produced by species of the genera Dolichospermum, Oscillatoria, Phormidium, and Aphanizomenon (Carmichael et al. 1975; Metcalf and Codd 2009). Another potent neurotoxin is the alkaloid saxitoxin, produced by the cyanobacteria Anabaena spp. Aphanizomenon spp., Cylindrospermopsis spp., Lyngbya sp., and Planktothrix sp. Other chemical types are the non-proteinogenic amino acid neurotoxins, one of which is beta-Methylamino-Lalanine, produced by many cyanobacteria of terrestrial, freshwater, or brackish water origin. Human exposure to a cocktail of cyanobacterial toxins is likely; however, the implications of combined exposure to these toxins have not been fully explored. Increased understanding of the combined effects of cyanobacterial toxins is required to fully understand how these molecules impact on human health (Rodgers et al. 2018). Increasing temperatures as presently occur in the global change climatic complex might favor cyanobacterial growth over the phytoplankton (Paerl and Huisman 2008) and these altered conditions can have the potential to increase the toxin production.

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The occurrence of Nodularia spumigena, a typically cyanobacterium from brackish waters in Europe (Baltic Sea, Mediterranean), Canada, USA, South Africa New Zealand, and Australia, was recently reported forming a dense bloom in a shallow brackish water in south-east Queensland, sub-tropical Australia for the first time (McGregor et al. 2012). Satellite monitoring of cyanobacterial harmful algal blooms are a successful tool to indicate locations with high exposure to cyanobacterial blooms and to assist management actions for recreational and drinking water sources (Kahru et al. 1994; Clark et al. 2017). The function and the ecological role of hepatotoxins and neurotoxins are currently unresolved. It has been suggested that they have evolved in response to grazing pressure by zooplankton (e.g. DeMott and Moxter 1991). However, fossils of akinete-forming cyanobacteria and thus the common ancestors of mycrocystin-producing cyanobacteria suggest that they are at least 1500–2000 million years old, so that microcystin production predates the metazoan lineage (Rantala et al. 2004).

3.7.4 Eco-Physiology 3.7.4.1 Desiccation Tolerance and Anhydrobiosis. In terrestrial habitats, cyanobacteria are frequently exposed to desiccation and thus must be poised for rapid resuscitation upon wetting. One of the desiccation-related problems is that, like all hydro-passive organisms, cyanobacteria cannot regulate their cell water content and thus completely depend on environmental conditions. Water loss from cyanobacteria and other poikilohydric organisms, like, for example, many eukaryotic algae, most lichens, and many bryophytes, can go as far down as 1–2% of cell/thallus water content (Billi and Pots 2000). Further desiccation might kill the cells. This phenomenon to revive after desiccation is termed “anhydrobiosis” and is also known as “desiccation tolerance” (Alpert 2005). In contrast to eukaryotic algae, cyanobacteria have some peculiarities that are unique to this group. They have no organelles, and thus the chains of electron transport carriers of respiration and photosynthesis are localized in the same membrane. The second and unique difference is the presence of a special super antenna for light harvesting: the phycobilisomes, proteinaceous structures that allow PPFD capture from low flux densities and also from the green part of the light spectrum (for an overview, see also Lüttge 2011). Desiccation-tolerant cyanobacteria must either protect cellular structures from damage during desiccation or repair

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them upon rewetting (e.g. review Büdel 2011). The strategies of desiccation tolerance were investigated on colonies of Chroococcidiopsis sp. and it became clear that the surviving cells were avoiding and/or limiting genome fragmentation, preserving intact plasma membranes and phycobiliprotein autofluorescence, as well as exhibiting spatially reduced reactive oxygen species accumulation and dehydrogenase activity upon rewetting. Between 10 and 28% of the cells of the Chroococcidiopsis sp. colonies survived (Billi 2009). The EPS of cyanobacteria can take up considerable amounts of water and it is hypothesized that the water trapped in EPS slows the desiccation rate on the one hand and enables cyanobacterial cells to make necessary preparations for desiccation on the other hand (e.g. Mazor et al. 1996; Colica et al. 2014). In the lichenized state, however, all cells of Chroococcidiopsis spp. photobionts seem to survive, accumulations of dead photobiont cells could never be observed after rehydration of different lichens species of the Lichinomycetes, even when kept for several years in the dry state (Büdel 2011). The filamentous, biological soil crust inhabiting cyanobacterium Microcoleus vaginatus, for example, showed recovery of photosynthesis within 1 h, accompanied by upregulation of anabolic pathways. At the onset of desiccation, genes for oxidative and photo-oxidative stress responses, osmotic stress responses, and the synthesis of carbon and nitrogen storage polymers were induced (Rajeev et al. 2013). For surviving extreme desiccation, many cyanobacteria produce both compatible solutes at an intracellular level and copious amounts of exopolysaccharides as a protective coat (Urrejola et al. 2019). A preparation of cyanobacteria for dehydration was discovered as signaling of forthcoming dehydration by dawn illumination was observed in the filamentous cyanobacterium Leptolyngbya ohadii. However, so far the mechanisms behind this are unknown. It was found that exposure to far-red light or lack of ground warming during dawn severely reduced revival rates after rewetting and altered the network of gene expression. Light and temperature sensing was attributed to phytochromes and many genes were up‐ or down‐regulated before water content decline, while others were strongly affected by the progression of dehydration and desiccation. Despite the fact that photosynthetic activity was regained during early rewetting, transcription was only barely observed during that phase but continued during the desiccated phase. It was demonstrated that RNA is stabilized during desiccation, possibly by intrinsically disordered proteins. The authors conclude that increasing light and temperature at dawn activates a network of genes preparing the cells towards dehydration. It seems that in addition to preparing towards

B. Büdel

dehydration, the cells also prepare for forthcoming rewetting, during dehydration (Oren et al. 2019). All of these aforementioned processes indicate the synchronization between terrestrial cyanobacteria and their environment. What is good for cyanobacteria to survive in harsh environments is a problem for investigation of them and cyanobacteria from extreme environments are often refractory to cell disruption methods. Nevertheless, recent studies on the unicellular genus Gloeocapsopsis species from the hypolithic environment of the Atacama Desert revealed a genomic arrangement exclusive of the genus that is associated with the recycling of the compatible solute trehalose, a sugar involved in desiccation tolerance of cyanobacteria (Urrejola et al. 2019).

3.7.4.2 Light and Carbon Dioxide-Exchange Like in all other photosynthetic organisms, cyanobacteria include the full range of light and shade adapted species. The range of light intensities (better: PPFD) over which photosynthesis can occur is determined by photosynthetic and metabolic characteristics of each species. These, again, are genetically controlled, meaning that a given species is “genotypically” adapted to a particular range (Richardson et al. 1983). Many, if not most, cyanobacteria species live in aquatic environments. There, light is attenuated exponentially with depth. Additionally, depending on the type (freshwater or marine, suspended load), water filters the light passing through. Generally, in all types of water, red light is essentially undetectable at depths of about 10 m, while blue light penetrates deepest. With their accessory pigment endowment, cyanobacteria can adapt/acclimatize to different light qualities, e.g. in deep ocean waters with predominantly blue light or inside forests with green light, they can increase phycoerythrin and improve the harvesting of blue and green light. For the bulk of microalgae, the LCP (light compensation point) is between 4 and 20 µmol m−2 s−1, most aquatic microalgae have an optimal NP between 400 and 700 µmol (Richardson et al. 1983). The lowest PPFD recorded so far at which still NP was measured is that for the Nostoc photobiont of the temperate rainforest lichen from New Zealand, Pseudocyphellaria dissimilis. The lower light compensation point (NP compensates DR) was at 1 µmol m−2 s−1 PPFD and photosynthesis was saturated at 20 µmol m−2 s−1 PPFD only (Green et al. 1991). Maximal PPFD measured at a cyanobacterial community under natural conditions was 2700 µmol photons m−2 s−1 at 2400 m altitude on top of the Auyan Tepui, Guayana Highland, Venezuela. Patches of the filamentous and truly branched cyanobacteria Stigonema ocellatum and S. panniforme still performed positive NP, despite the unusually high PPFD (Büdel 1999).

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3.7.4.3 Temperature and Carbon Dioxide-Exchange Aquatic cyanobacteria found in geothermal habitats worldwide are those that at 57–74 (Fig. 3.52) °C tolerate the highest temperatures where they still perform NP (Ward et al. 2012). Among them, Synechococcus cf. lividus (high temperature form, HTF) is the taxon that tolerates the most extreme temperatures and is found in geothermal areas of North America (Fig. 3.51), Latin America, Thailand, and China. Other prominent species from geothermal habitats are Cyanothece cf. minervae (max. 62 °C), Chlorogloeopsis HTF (max 64 °C), Leptolyngbya spp. (*62 °C), Fischerella cf. laminosus (max. 58 °C, Leptolyngbya cf. amphigranulata (*56 °C), Geitlerinema cf. terebriformis (max. 55 °C), Spirulina cf. labyrinthiformis (max. 51 °C), Calothrix spp. (*53–55 °C), and Pleurocapsa spp. (max. 57 °C), (Ward et al. 2012). In the Mushroom Spring of the Yellowstone National Park, ten putative ecotypes of Synechococcus form mats along hot water channels/gradients ranging in temperature from 56 to 68 °C. At all temperatures of these Synechococcus mats, photosynthetic oxygen release was measured from the mat surface to 2 mm depth (Ward et al. 2006).

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Terrestrial cyanobacterial soil crusts, for example, at the Boodjamulla National Park in the Gulf Savannah region of Queensland, Australia, reached surface temperatures close to 50 °C in the dry and 40 °C in the wet state. Although the pure culture of Symplocastrum purpurascens isolated from this biocrust still performed positive NP at 47 °C (Fig. 3.53a), this never happened in the field during a one-year measuring period (Fig. 3.53b). At temperatures above 42 °C, the biocrust was usually dry and inactive (Büdel et al. 2018). In the dry state, surface temperatures of cyanobacteria-dominated rock or soil crusts above 60 °C are not uncommon in tropical regions around the world. Sudden rain events cool down the cyanobacterial crust at the rock/ soil surface immediately to below 40 °C and thus prevent the cells from thermal damage.

3.7.4.4 Thallus Water Content and Carbon Dioxide-Exchange The poikilohydric nature of cyanobacteria allows them to overcome drought periods in general, some can even survive desiccated for many years. Only a few minutes after remoistening, metabolic activity occurs and photosynthesis starts a few minutes later. While aquatic cyanobacteria do

Fig. 3.52 Hot spring cyanobacteria from Yellowstone National Park. ©Don Johnston PL/Alamy Stock Photos

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B. Büdel

Fig. 3.53 CO2 gas exchange of terrestrial cyanobacteria from a biological soil crust in Queensland, Australia; a laboratory measurement of temperature-related NP of a Symplocastrum purpurascens pure culture isolated from the biocrust (n = 3); b field measurement of the S. purpurascens dominated biocrust under natural conditions; all NP data points of one year plotted in the diagram

live with the restrictions of CO2 availability in water and often possess a CCM to overcome at least partly the CO2 shortage, colony-forming terrestrial cyanobacteria suffer considerably from not enough, but even more from a surplus of water (Lange et al. 1993; Büdel 1999; Büdel et al. 2018). In contrast to many eukaryotic algae and bryophytes, cyanobacteria cannot be reactivated by high air humidity alone; they need more water than can be gained from high air humidity within a certain time (Lange et al. 1986). So far only one exception is known, the desert soil cyanobacterium Sociatus tenuis reached 30% of its maximum NP rate with an air humidity as low as 96% at 15 °C under experimental conditions in the laboratory (Lange et al. 1994); conditions that would probably never be met in nature. The colony-forming lifestyle of terrestrial cyanobacteria might be of advantage during desiccation as it slows down the water loss and thus allows an extended period of photosynthetic activity. However, at full water saturation CO2 diffusivity is reduced roughly 10,000 times compared to air and thus slows down photosynthetic CO2 fixation drastically (e.g. Badger et al. 1993). This however complicates the measurement of NP rates, not only in cyanobacteria but in all organisms with a colonial or thallus organizational type. They do not only have an optimal temperature range for NP and DR but also an optimal thallus water content (TWC)! Below and above the optimal TWC NP rates drop, either because there is not enough water available or there is excessive water acting as an additional diffusion resistance. Structure and organization of the colony/thallus also influence optimal TWC for maximum NP rates. Wet Nostoc commune thalli have an enormous amount of EPS compared to the colonies of the unicellular

Chroococcidiopsis sp. and need fivefold the amount of water to reach optimal TWC and thus optimal NP (Fig. 3.54a, b).

3.7.4.5 Effects of Phosphorous Regardless of marine, freshwater, or terrestrial ecosystems, phosphorous availability can considerably influence the growth of cyanobacteria. Cyanobacteria of the family Rivulariaceae react especially sensitively to low phosphorous availability. As early as in 1969 an inhibitory effect of low inorganic phosphate concentrations on the growth of Gloeotrichia sp. was reported from Lake Berkelse near Delft, Netherlands (Fogg 1969). Many species of the Rivulariaceae can form multicellular elongated hair-like structures void of chlorophyll at the apical end of the tapering trichomes (Figs. 3.37h, 3.46h). Other species, for example, the genus Calothrix, develop tapering “hairs” that retain their chlorophyll but do not elongate. It was observed that phosphorous limitation enhanced tapering in most Rivulariaceaen species but only one-third of them developed hairs. In addition, cultures growing under increasingly phosphorous limited conditions even stopped hormogonium production (for a comprehensive overview, see Whitton and Mateo 2012). Accumulation of phosphorus apparently also plays an important role in the ability of the bloom-forming cyanobacterium Nodularia spumigena to grow even when dissolved inorganic phosphorus is depleted. A recent study investigated phosphorus incorporation and distribution in cyanobacterial filaments using scanning electron microscopy in combination with energy-dispersive X-ray analysis and nanoscale secondary ion mass spectrometry (NanoSIMS). After the addition of phosphate to a phosphorus-depleted

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Fig. 3.54 CO2 gas exchange of terrestrial cyanobacteria; a Nostoc commune, isolated from a biocrust; b Chroococcidiopsis sp., isolated lichen photobiont. The yellowish rectangle marks the range of optimal NP (90% of max.); both species were measured in the range of their optimal light and temperature. TWC given as equivalents of mm water column (like precipitation values). Figure (a) courtesy of Michelle Szyja

population, the phosphate concentration decreased in the water, and isochronally intracellular polyphosphate accumulated. At the cellular level, phosphorus was stored in the form of polyphosphate granules in vegetative cells but heterocytes remained low in phosphorus. This demonstrates that the division of tasks between vegetative cells and heterocytes is not only restricted to nitrogen fixation (Braun et al. 2018).

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99 Zhang P, Allahverdiyeva Y, Eisenhut M, Aro EM (2009b) Flavodiiron proteins in oxygenic photosynthetic organisms: photoprotection of photosystem II by Flv2 and Flv4 in Synechocystis sp. PCC 6803. PLoS One 4:e5331 Zhu C, Guo G, Ma Q, Zhang F, Ma F, Liu J, Xiao D, Yang X, Sun M (2017) Diversity in S-layers. Prog Biophys Mol Biol 123:1–15 Zilliges Y (2014) Glycogen, a dynamic cellular sink and reservoir for carbon. In: Flores E, Herrero A (eds) The cell biology of cyanobacteria. Caister Academic Press, Norfolk, pp 189–210

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Algae from Primary Endosymbioses Burkhard Büdel and Thomas Friedl

Contents 4.1 Rhodophyta, Red Algae ...................................................................................................................... 102 4.1.1 Origin of Red Algae................................................................................................................. 102 4.1.2 Morphology and Cell Structure................................................................................................ 104 4.1.3 Reproduction, Life Cycle ......................................................................................................... 107 4.1.4 Phylogeny, Systematics, and Diversity .................................................................................... 112 4.1.5 Genome Reductions and Gains: The Ecological Imprint ........................................................ 126 4.1.6 Ecology ..................................................................................................................................... 128 4.1.7 Phylogeography ........................................................................................................................ 130 4.2 Chloroplastida—Green Algae ............................................................................................................ 131 4.2.1 Ecological and Economic Importance ..................................................................................... 132 4.2.2 Origin of Green Algae ............................................................................................................. 133 4.2.3 Defining Characters of the Green Algae.................................................................................. 134 4.2.4 Reproduction and Life Cycle ................................................................................................... 142 4.2.5 Systematics and Classification of the Chloroplastida .............................................................. 143 4.2.6 Phylum Prasinodermophyta...................................................................................................... 146 4.2.7 Phylum Chlorophyta and Prasinophytes .................................................................................. 147 4.2.8 The Core Chlorophyta, Chlorodendrophyceae, and Pedinophyceae ....................................... 148 4.2.9 Class Trebouxiophyceae........................................................................................................... 148 4.2.10 Class Chlorophyceae ................................................................................................................ 158 4.2.11 Class Ulvophyceae ................................................................................................................... 167 4.2.12 Phylum Streptophyta—The Streptophyte Algae Grade........................................................... 177 4.3 Glaucophyta ......................................................................................................................................... 198 4.3.1 Origin of the Phylum Glaucophyta.......................................................................................... 198 4.3.2 Morphology and Cell Structure/Function ................................................................................ 198 4.3.3 Genome..................................................................................................................................... 200 4.3.4 Classification and Systematic Arrangement of the Glaucophyta ............................................ 200 4.3.5 Ecology ..................................................................................................................................... 202 4.4 Cercozoa—A Second Primary Endosymbiosis................................................................................. 203 4.4.1 Primary Endosymbiosis? .......................................................................................................... 204

B. Büdel (&) RPTU Kaiserslautern, Department of Biology, Erwin-Schrödinger-Str. 13, 67663 Kaiserslautern, Germany e-mail: [email protected] T. Friedl Department of Experimental Phycology and Culture Collection of Algae (EPSAG), Georg August University Göttingen, Nikolausberger Weg 18, 37073 Göttingen, Germany e-mail: [email protected] © Der/die Autor(en), exklusiv lizenziert durch Springer-Verlag GmbH, DE, ein Teil von Springer Nature 2024 B. Büdel et al. (eds.), Biology of Algae, Lichens and Bryophytes, https://doi.org/10.1007/978-3-662-65712-6_4

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Origin ........................................................................................................................................ 204 Morphology and Ultrastructure ................................................................................................ 204 Classification and Systematic Arrangement............................................................................. 206

References ..................................................................................................................................................... 207

4.1

Rhodophyta, Red Algae

Burkhard Büdel The red algae are a group of important photoautotrophic organisms that include unicellular microalgae as well as large macroalgae with a size of more than one meter. Phylogenetically they are descendants of a primary endosymbiosis event between a heterotrophic eukaryote and a cyanobacterial endosymbiont roughly 1,200–1,600 million years ago (Yoon et al. 2004, 2010; Rodríguez-Espeleta et al. 2005; Bengtson et al. 2017). They are primary plastid-bearing organisms. As one of their major innovations, the morphologically more complex red algae evolved a triphasic life cycle, including one haploid and two diploid phases, the tetrasporophyte and the carposporophyte. The diploid carposporophyte is borne on the haploid female gametophyte. Red algae occur in both, fresh-and marine waters (Fig. 4.1) but reach their highest diversity along the coastal waters of the World’s oceans. True aero-terrestrial species are unknown. Today, the red algae include more

than 7,100 species (Yoon et al. 2017) with high diveristy especially in tropical and temperate marine waters. Red algae have a longstanding history of human use and today they are cultivated for the production of food, food processing and laboratory use in microbiology in the form of gel-forming agar, agarose, and carrageenan, for example.

4.1.1 Origin of Red Algae The first eukaryotic organisms acquired their ability to convert light into chemical energy through endosymbiosis with a cyanobacterium (see Chap. 2). This first endosymbiosesis gave rise to the nowadays named “primary” plastids, which are present in all green plants (including algae), red algae, and Glaucophytes. The origin of the Rhodophyta itself is still a matter of debate, nevertheless, based on a six gene phylogeny from 46 taxa, an estimated age of 1,300 million years was determined (Yoon et al. 2004). Red algae were traditionally classified as plants, thought to be the most ancient eukaryotic organisms.

Fig. 4.1 Red algae: Corallina officinals (gracefully build shrubs) and Lithospermum sp. (pink rock crust) in their natural marine environment at Helgoland, Germany

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The first molecular studies indicated similarities between red and green plastids, suggesting a single endosymbiotic origin of the red algal plastids with those of the green plants. On the basis of sequencing of the largest subunit of RNA polymerase II from red and green algae, phylogenetic analyses provided strong support for an early emergence of the Rhodophyta, preceding the origin of the line of green plants (Stiller and Hall 1997). Initially, it was a widely accepted view that primary plastids arose only once (Palmer 2003). Following this line of argumentation, the predictions that (1) plastids form a monophyletic group and (2) all primary photosynthetic eukaryotes are implied. A phylogenomic analyses that compared 50 genes from 16 plastid and 15 cyanobacterial genomes with 143 nuclear genes from 34 eukaryotic species unraveled a strong significant support for a single endosymbiotic event giving rise to the primary photosynthetic eukaryotes (Rodríguez-Ezpeleta et al. 2005). It is interesting to note that there is evidence for a close relationship with plastids of heterokont algae from plastid genome phylogenies, indicating that they all derive from the same endosymbiosis event via incorporation of a red alga. This also supports the view of a relatively simple path of linear descent for the evolution of photosynthesis in a large proportion of algae, and emphasizes plastid loss in several lineages (Janouškovec et al. 2010). Based on molecular clock analysis, Yoon et al. (2004) suggested 1,274 million years ago as the date for red algal secondary endosymbiosis.

4.1.1.1 Fossil Record Because of their ability to induce calcium carbonate precipitation in- and outside their cell walls, calcareous red algae have quite a good fossil record. The so far oldest record of a supposed red alga is that of Rafatazmia chitrakootensis from the 1,600 million years old Tirohan Dolomite of the Lower Vindhyan in central India that contains phosphatized stromatolitic microbialites. These fossils are extremely well preserved and interpreted by the authors as belonging to the crown-group red algae. Rafatazmia chitrakootensis is a uniseriate filamentous alga with large cells of diffusely distributed septation (Fig. 4.2). Each cell has a centrally suspended, conspicuous rhomboidal disk, thought to be the remains of a pyrenoid. The septa between the cells have central structures resembling pit connections and pit plugs. Other filamentous species found in the Tirohan Dolomite are Denaricion mendax, with coin-like cells that are reminiscent of the large sulfur-oxidizing bacteria or members of the oscillatoriacean cyanobacteria and the lobate sessile fossil Ramathallus lobatus with pseudoparenchymatous thalli and apical growth. The morphology of the latter even suggests affinity with the morphologically more complex Florideophyceae. If these inferences are correct, the

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genera Rafatazmia and Ramathallus represent the crown group of multicellular rhodophytes, antedating the oldest previously accepted red alga in the fossil record by about 400 million years (Bengtson et al. 2017). The formerly oldest fossil red algal record dates back 1,000 million years (Gibson et al. 2017) and was the filamentous species Bangiomorpha pubescens originating from the Hunting formation, Mesoproterozoic deposits on Somerset Island of arctic Canada (Butterfield et al. 1990; Butterfield 2000, 2015). With the thalli formed by a combination of uni- and multiseriate filaments, this fossil strongly resembles the modern genus Bangia (Fig. 4.3). The red algae of the subclass Corallinophycidae have the ability of cell wall calcification and thus are also predestined to leave a fossil record. Indeed, there are calcified remains from the latest Proterozoic Nama Group in Namibia (550 million years old) that may represent red algae of the Corallinales because of their conceptacle-like structures (Grant et al. 1991). There is also strong evidence for coralline red algal fossils in the lower Ordovician (roughly 500 million years ago). For example, the fossil genus Arenigiphyllum closely resembles vegetative parts of coralline red algae of the common modern genera Lithophyllum or Lithothamnion (Riding et al. 1998). The fossil record of coralline red algae dates back to at least 500 million years ago and is uninterrupted until today (Pena et al. 2020).

104 Fig. 4.3 Fossil red alga Bangiomorpha pubescens from the Mesoprozterozoic Hunting formation of arctic Canada. a Multiseriate filament; b uniseriate filament with basal holdfast. From Butterfield (2000) with permission of Palaeobiology

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4.1.2 Morphology and Cell Structure 4.1.2.1 General Morphology The red algae have achieved unicellular, colonial, filamentous, and complex thallus organizational levels. They do not develop mobile cells, even during sexual reproduction. Their bodies (thalli) are either heterotrich (unequally filamentous) or subdivided into three distinct morphological sections rhizoid (adhesive disc), cauloid (stem), and phylloid (leaf-like structure). The thalli are generally pseudoparenchymatic, which means that independently of the complexity of their bodies, they are always composed of filaments and do not form a true tisue, as for example in certain green algae, brown algae, bryophytes, and vascular plants (Fig. 4.3). Pseudoparenchymes are formed by clustering of filaments merging with their gelatinous cell walls. The less complex multicellular taxonomic groups (e.g., the class Bangiophyceae) do not have specialized meristematic cells and growth occurs anywhere in the thallus. In the morphologically more complex class of the Florideophyceae, apical growth by means of a terminally located apical cell is realized. Besides branched filamentous thalli, also shoot- or leaf-like types exist. Two main general structures occur with thalli that are either uniaxial, composed of a single main branched filament in the center of the thallus (Fig. 4.4a), or they are multiaxial and then composed of many filamentous axes, each of which are derived from an own terminal apical cell (Fig. 4.4b). The central filament of the uniaxial type forms numerous whorled side branches in equispaced distances along the main axes, finally forming a smooth outer texture like in the genus Lemanea (Fig. 4.4c), or resulting in a moniliform habitus, for example, in the genus Batrachospermum. The leaf-like thallus of, for example, the genus Delesseria is a two-dimensional subtype of the uniaxial growth form where the leafy structure is derived by a peculiar mode of

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branching. The first-order branches on both sides of the central filament only form second-order branches at the lower side that congenitally merge with their cell walls (Fig. 4.4d, e). In the multiaxial type, the parallel filaments branch dichotomously with the side branches continuing branching and form a closed outer texture (Fig. 4.4a). Calcified thalli Red algae with calcified cell walls often expose a somehow multiaxial thallus structure. Their thallus is either articulated (Fig. 4.21a, b) or non-articulated, geniculate or non-geniculate (Graham et al. 2016). The non-geniculate thalli of coralline algae separate into a lower hypobody, functioning as a holdfast to the substratum in the lowermost part, and a peribody forming the upper thallus part (Fig. 4.21c–e). Filaments of the hypobody are arranged horizontally with their marginal cells being meristematic. Filaments of the peribody are arranged vertically, having their meristematic cells subterminal, forming the epithallus, a one to several cells thick upper layer. Calcification occurs in growing regions of the red algae, and calcium carbonate crystals become deposited either in the form of calcite or of aragonite onto an organic matrix outside their cell walls. The deposition of calcium carbonate in the red algal thallus is controlled in a way that allows a defined precipitation of calcium carbonate at specific locations in the thallus. All but three cell types in the coralline algae are calcified; (1) cells of reproductive structures, (2) branch joins (genicula), and (3) lesion sites of thalli undergoing reparation (Borowitzka and Vesk 1978; Bilan and Usov 2001; Pueschel et al. 2005). The calcification process can be indirectly supported by the photosynthetic rise of the pH, but in general, it is not known how calcium carbonate precipitation in red algae is controlled (see also review of McCoy and Kamenos 2015). It was observed that in the geniculate Corallina pilulifera,

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Fig. 4.4 Morphology of red algae. a Multiaxial body of robust and somehow fleshy red algae; b uniaxial body, composed of single branched filaments with subapical cells that divide radially and form whirls of branch filaments; c uniaxial body of Lemanea sp., longitudinal section showing the central single branched filament and its lateral branching that leads to a cortex like outer structure; d, e phylloid development from filaments in Caloglossa leprieurii, primary and secondary filaments are indicated by grey color in e. From Esser (2000), Kryptogamen 1 with permission of Springer Nature

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4.1.2.2 Cell Structure As red algae never have flagella for locomotion, they are consistently unable to move during all developmental stages of their life history. However, gliding motility is possible to some extent for many unicellular red algae and spores (monospores, tetraspores, carpospores, and zygotospores). Red algal cell walls consist of cellulose microfibrils embedded in an amorphous matrix containing sulfated galactan polymers and mucilage. Because their cell walls are less rigid than that of other algae, they are often referred to as extracellular matrix (ECM). The gelatinous mucilage is usually composed of agars and carrageenans, whose major constituents are highly hydrophilic, sulfated polygalactans that are polymers of b-(1–4) galactose and a-(1–3) linked 3,6 anhydrogalactose. While the carrageenans are characterized by the presence of D-galactose and anhydro-D-galactose, the agars are also more heavily methoxylated. There are several types of carrageenans and agars present in different species. Both, agars and carrageenans, are of considerable economical value and are used, for example, in food production and microbiology. Mucilages are produced in the Golgi apparatus and are polymers of D-xylose, D-glucose, D-glucuronic acid, and galactose. Their proportions differ among taxa. Red algae of the corallines and some other taxa have heavily calcified cell

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walls (ECM’s). There, calcium carbonate is deposited in the crystalline form of calcite, while in some Nemaliales it is deposited as aragonite crystals. Among cell organelles, the Golgi bodies are often associated with other organelles. Three types are distinguished: (1) Golgi bodies are associated with the nucleus in the Rhodellophyceae, (2) Golgi bodies are associated with the endoplasmatic reticulum in the Cyanidiophyceae, Compsopogonophyceae, Stylonematophyceae, and the genus Rhodella of the Rhodellophyceae, (3) Golgi bodies are associated with mitochondria in the Porphyridiophyceae, Bangiophyceae, and Florideophyceae. Unlike in green algae, food reserves like starch in the chloroplasts are not present in red algal plastids. Instead, granules of a differently branched glucan, the Floridean starch, are stored outside the plastid in the cytoplasm (Fig. 4.5). Cell–cell connections After the formation of the new cell wall between two neighboring red algal cells, there is only one single pore space left, the pit connection which is immediately plugged up by a pit-plug. The characteristic spherical pit-plug core consists of proteinaceous material. Besides the pit-plug core, further elements can be associated with the core. Additional components can be present at the cytoplasmic face of the core and those are cap membranes and inner and outer cap layers. At least 8 different types of pit-plug connections can be distinguished in red algae, and they are often taxon specific (Fig. 4.6).

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Fig. 4.5 Ultrastructure of a red algal cell (Agardhiella subulata). From Kamiya et al. (2017), courtesy of Nakayama and Kamiya with permission of Borntraeger Verlagsbuchhandlung (www.schweizerbart.de)

Fig. 4.6 Cell–cell pores in red algae (pit plugs). a Plug core without any additional structure, Compsopogonales, Rhodochaetales, and Ahnfeldiales; b with inner cap layers, Bangiales; c with inner cap layers and cap membranes, Hildenbrandiales and Gelidiales; d with cap membranes only, Pihiellales and most Rhodymeniophycidae; e with cap membranes between inner and outer cap layers, Palmariales, Acrochaetiales, Colaconematales, Entwisleiales, and Nemaliales; f two cap layers only, Rhodachlyales and Thoreales; g like E but outer cap layers dome-shaped expanded, Colaconematales; h same as G but without cap membranes, Balbianiales, Batrachospermales, and Corallinophycidae (Kamiya et al. 2017). From Kamiya et al. (2017), courtesy of M. Kamiya with permission of Borntraeger Verlagsbuchhandlung (www. schweizerbart.de)

Multinucleate cells and polyploidy Because of relatively high diffusion resistances for all sorts of molecules in liquid cytoplasm, transport of material information is limited. The larger a cell is, the longer the transport processes require. So far, two ways to overcome this restriction in cell size exist, or are suggested: (1) improvement of cellular transport by cytoplasmic streaming and (2) nuclear omnipresence due to many nuclei via mitosis. It is known from the red algal genus Porphyra that cytoskeletal motor proteins are restricted to a small set of kinesins, appearing to be the only universal cytoskeletal motors within the red algae. The surprisingly minimal cytoskeleton apparently offers a potential explanation for why red algal cells and multicellular structures are more

limited in size than in most other multicellular algal lineages (Brawley et al. 2017). As an alternative, endoreduplication, a repeated replication of the entire nuclear genome without intervening mitosis, resulting in polyploidy is realized. This common process of genome amplification in red algae is also known as nuclear polygenomy, endopolyploidy, or polyteny. In some red algae, the apical meristematic cell of filaments is also multinucleate, although apical cells are not typically polyploid (Goff and Coleman 1990). The typical variation among red algae in numbers of nuclei and polyploid nuclei and their combination in apical and other cells of filaments is summarized in Fig. 4.7. There is apparently often a tight correlation observed between the number of nuclei and the number of plastids in red algae (Goff and Coleman 1990).

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Fig. 4.7 Variation of nuclear patterns in apical and derived cells of Florideophycidae. Modified from Goff and Coleman (1990)

Nuclear status of: Apical cell

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Plastids and red algal photosynthesis

4.1.3 Reproduction, Life Cycle

Red algal chloroplasts are surrounded by a double-membrane envelope and can be with or without one or more pyrenoids. They are of a stellate, ribbon-like, or discoid shape. Their thylakoids are unstacked and evenly spaced. One or sometimes 2 thylakoids run around the periphery, parallel to the chloroplast envelope. However, this feature is not observed in the classes of the Porphyridiophyceae, Bangiophyceae, and a few genera of the Rhodellophyceae (Scott et al. 2010). The photosynthetic pigments are chlorophylla, the phycobiliproteins phycoerythrin, phycocyanin, and allophycocyanin as well as several xanthophylls. The biliproteins are united in phycobilisomes that are attached to the stromal surfaces of the thylakoids and are associated with the light-harvesting antennae of PS II (Gantt 1990). Most living organisms with an oxygenic photosynthesis accumulate either glycogen or starch as energy storage polymers. While chloroplast starch synthesis in green algae and plants occurs via ADP‐glucose, the Floridean starch synthesis in red algae proceeds via uridine diphosphate‐ glucose in semblance to eukaryotic glycogen synthesis and occurs in the cytosol rather than the plastid. Nevertheless, the pathways of starch synthesis in green and red algae represent chimeras of the host and endosymbiont glycogen synthesis pathways. But, in contrast to green algae, the endosymbiont‐derived proteins function in the cytosol of the red algae (Patron and Keeling 2005).

Like in many other algal groups, red algal vegetative and sexual reproduction modes exist. However, asexual reproduction via thallus disintegration, like in other thallus forming algal groups, is rarely observed. Vegetative reproduction occurs via secondary cycles by mito-aplanospores also termed monospores and may be found in gametophytes and sporophytes. Sexual reproduction is usually connected with the alternation of a haploid (gametophyte) and a diploid (sporophyte) generation. Gametophytes are either monoecious1 (all sexes occur on one plant) or dioecious (both sexes occur on two different plants (male and female) that can be isomorphic (both sexes look the same) or heteromorphic (both sexes have different shapes). Fertilization starts with the adhesion of a generally non-flagellated spermatium (male gamete) on a trichogyne, a hair-like extension of the carpogonium (egg cell) developing from a female gametophyte.

4.1.3.1 The Biphasic Life Cycle In the class Bangiophyceae, an alternation of two generations occurs, the usually macroscopic gametophyte (haploid, n) is followed by a normally microscopic sporophyte (diploid, 2n) after fertilization. By fertilization the carpogonium transforms into the zygote, that then divides via 3 mitotic divisions completely into 4, to 32 carpospores (more 1

Monoecious/dioecious is the correct terminology given that the sex is determined in the haploid phase (see Vranken et al. 2023).

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generally “mitospores”, but also named “zygotospores” because they are formed by the zygote). The carpospores grow to form the sporophytes, also named the “conchocelis phase”. The sporophyte produces sporangia (conchosporangia) in which meisosis takes place forming the meioaplanospores (conchospores) that then produce the male and female gametophytes (Fig. 4.8a).

4.1.3.2 The Triphasic Life Cycle In the Florideophyceae the female gametangium (oogonium or carpogonium) caries an elongated conception cell (trichogyne). Together with some basal cells it forms the

procarp (Figs. 4.9 and 4.10). Ahead of fertilization, a single-layered covering (pericarp) develops and together with the procarp the cystocarp is developed. The zygote remains in the carpogonium and develops diploid spore-forming filaments. They are the diploid intermediate generation and are termed the carposporophyte or in the older literature the gonimoblast. Via mitoses the carpospores are formed from which the tetrasporophyte arises. In the sporangia of the tetrasporophyte, four tetraspores (haplo-aplanospores) are formed after a meiotic division. From the carpospores the new gametophytes develop.

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Fig. 4.9 Triphasic life cycle of Polysiphonia sp.: Mating type is a gameto-gametangiogamy. Original drawing Spindler & Büdel

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Fig. 4.10 Reduced life cycle of Batrachospermum sp. The carpospores form the diploid sporophyte (the so-called Chantransia-stage) that does not form tetrasporangia and thus tetraspores, but rather undergoes meiosis in their terminal cells from which the new gametophyte arises. The mating type is a gameto-gametangiogamy. The formation of monospores in the Chantransia-stage is a means of vegetative reproduction. Original drawing Spindler & Büdel

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Fig. 4.11 Stages of the triphasic and reduced triphasic life cycle. a Pterothamnion plumula, stained with cotton-blue, carposporophyte with carposporangia (arrows); b tetrasprophyte with tetrasporangia (arrows); c tetrasporangia. d, e, g, h Batrachospermum cf. gelatinosum, gametophyte with carposporophytes (d, e, arrows), carpogonia with trichogynes (g, h, arrows). Batrachospermum sp., new carpogonium (arrow) on gametophyte (f); courtesy of Johanna Knappe, Marburg, Germany. Scale bar in this figure is approximation

The reduced life cycle of Batrachospermum

Variation of the carposporophytes

In species of the genus Batrachospermum, the triphasic life cycle is somewhat reduced as there are no tetrasporangia developed in which the meiosis takes place. Rather, in terminal cells of the Chantransia phase meiosis occurs and the new gametophyte grows from the terminal cell (Figs. 4.10 and 4.11).

There are several ways realized in the Florideophyceae by which carposporophytes develop after fertilization (taken from Kamiya et al. 2017): (1) The fertilized carpogonium directly produces small filaments which in summary form the “gonimoblast”. These filaments form the carposporangia at their apices (e.g., in the genus Nemalion).

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(2) Many carpogonia with elongated trichogynes are formed within a dome-shaped structure named “conceptacle”. After fertilization, the zygotes and their adjacent cells fuse to form a single “fusion cell”. The fusion cell produces the carposporophyte (gonimoblast in the older literature) filaments with each of them forming a terminal carposporangium (e.g., the genus Corallina). (3) The fertilized carpogonium transfers its divided diploid nuclei by means of a connecting filament, originating from the zygote to several auxiliary cells (supporting cells). Each of the auxiliary cells then develop gonimoblasts and from this the carposporangia emerge terminally (e.g., Dudresnaya).

(4) A supporting cell connected with the zygote demerges an auxiliary cell, later fusing with the supporting cell and then forming the gonimoblast. Finally, carposporangia are developed at terminal ends of the gonimoblast filaments (e.g., Aglaothamnion).

The diploid nucleus moves into the supporting cell via an anastomosis (secondary cell–cell connection) between the fertilized carpogonium and the supporting cell. Later, the supporting cell and the auxiliary cell merge and form the gonimoblast.

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4.1.4 Phylogeny, Systematics, and Diversity Red algae form a monophyletic entity and are characterized by unique features, for example, the constant lack of flagella and centrioles, a very special type of cell–cell connection the so-called pit connections, and the occurrence of a triphasic life cycle in the class Florideophyceae. Extensive detailed and recent works on red algae are those of Kamiya et al. (2017) in the book series “Syllabus of Plant Families” and of Yoon et al. (2017) in the Handbook of Protista (Archibald et al. 2017). Many diverse molecular data support the monophyly of the Rhodophyta, the Viridiplantae (green algae and land plants), and the Glaucophyta. They are collectively referred to as the Archaeplastida (for an overview see Adl et al. 2019). There are several possibilities for the evolution of the Archaeplastida: (1) the multiple primary endosymbiosis hypothesis, where two or three different eukaryotic heterotrophs undergo endosymbiosis with a cyanobacterium, or (2) the widely accepted single primary endosymbiosis hypothesis, with the origin of the plastid by acquisition of a cyanobacterium by a common ancestor of Archaeplastida.

B. Büdel and T. Friedl

For the evolutionary history of many other algal groups, one of the most important evolutionary contributions of the red algae has been their function as a plastid donor through secondary endosymbiosis. A multigene phylogeny identified several well-supported lineages, the earliest to diverge was the Cyanidiophyceae, a strong monophyly of the Bangiophyceae and Florideophyceae was suggested and a seven class system proposed (Yoon et al. 2006). However, the internal relationships among the four classes Compsopogonophyceae, Porphyridiophyceae, Rhodellophyceae, and Stylonematophyceae still remain unresolved. Interestingly, results from mitochondrial and plastid genome analyses strongly suggest that organellar genome data can provide sufficient phylogenetic information to resolve most phylogenetic relationships in the Rhodophyta (Box 4.1; e.g., Yang et al. 2016; Yoon et al. 2017). A current classification scheme for the Division Rhodophytaas used in the following subchapters, is presented in Box 4.1.

4.1.4.1 Classification and Systematic Arrangement of the Rhodophyta A current classification scheme for the Division Rhodophyta, as used in the following subchapters, is presented in Table 4.1

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Table 4.1 Overview of the current classification of the Division Rhodophyta

Subdivisions Classes Subclasses Cyanidiophytina Cyanonidiophyceae Rhodophytina Porphyridiophyceae Stylonematophyceae Compsopogonophyceae

Rhodellophyceae

Bangiophyceae Florideophyceae Hildenbrandiophycidae Nemaliophycidae

Orders

Families

Cyanidiales

Galdieraceae

Porphyridiales Rufusiales Stylonematales Compsopogonales

Porphyridiaceae Rufusiaceae Stylonemataceae Boldiaceae, Compsopogonaceae Erythrotrichiaceae Rhodochaetaceae Dixoniellaceae Glaucosphaeraceae Rhodellaceae Bangiaceae

Erythropeltidales Rhodochaetales Dixoniellales Glaucosphaerales Rhodellales Bangiales Hildenbrandiales Acrochaetiales Balbianiales Baliales Batrachospermales Colaconematales Entwisleiales Nemaliales

Rhodoachlyales Thoreales Corallinales Hapalidiales Rhodogorgonales

Hildenbrandiaceae Acrochaetiaceae Balbianiaceae Balliaceae Batrachospermaceae Colaconemataceae Entwisleiaceae Galaxauraceae, Liagoracae, Liagoropsidaceae, Nemaliaceae, Scinaiaceae, Yamadaellaceae Meiodsicaceae, Palmariaceae, Rhodophysemataceae, Rhodothamniellaceae Rhodoachlyaceae Thoreaceae Coralinaceae Hapalidiaceae Rhodogorgonaceae

Ahnfelthiacles Pihiellales

Ahnfeltiaceae Pihiellaceae

Palmariales

Corallinophycidae

Ahnfeltiophycidae

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Rhodymenio-phycidae

Sporolithales

Sporolithaceae

Acrosymphytales Bonnemaisoniales

Acrosymphytaceae Bonnemaisoniaceae, Naccariaceae Callithamniaceae, Ceramiaceae, Dasyaceae , Delesseriaceae, Inkyuleeaceae, Rhodomelaceae, Sacromeniaceae, Spyridiaceae, Wrangeliaceae Gelidiaceae, Gelidiellaceae, Orthogonacladiaceae, Pterocladiaceae Acrotylaceae, Areschougiaceae, Blinksiaceae, Calosiphoniaceae Gracilariaceae Halymeniaceae, Trengiaceae Nemastomataceae, Schizymeniaceae Plocamiaceae, Pseudoanemoniaceae, Sarcodiaceae Champiaceae, Faucheaceae , Fryeellaceae, Hymenocladiaceae, Lomentariaceae, Rhodymeniaceae Sebdeniaceae

Ceramiales

Gelidiales

Gigartinales

Gracilariales Halymeniales Nemastomatales Plocamiales

Rhodymeniales

Sebdeniales

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Box 4.1: Phylogeny of the Rhodophyta

Red underlying: class level, ocher shows the subclass level; arrows indicate inconsistencies of the tree. Large letters in brackets give the type of pit-plugs present in the orders (see Fig. 4.6) and “npp” indicates that there are no pit-plugs realized. * Note that this taxon was assigned to the new order Atractophorales by Saunders et al. 2016. GPI = glycosylphosphat idylinositol. Bold lines indicate robust branches that are supported by bootstrap values  90% (according to Verbruggen et al. 2010, modified from Kamiya et al. 2017, Yoon et al. 2017 and Yang et al. 2016).

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4.1.4.2 Subdivision Cyanidiophytina If not cited otherwise all taxa numbers given in this section are taken from Guiry and Guiry (2023). Class Cyanodiophyceae This class comprises coccoid unicellular species only belonging to the order Cyanidiales with the two families Cyanidiaceae (2 genera, Cyanidium, Cyanidioschyzon, 3 species) and Galdieria ceae (1 genus, Galdieria, 5 species). It can be concluded from molecular phylogeny that they were the first red algal class to diverge. Their generally uninucleate cells can be covered by a cell wall or be naked. Normally, they contain single parietal and blue-green colored chloroplasts that do not have pyrenoids, and have peripherally arranged thylakoids (Fig. 4.12). Sexual reproduction is not known and asexual reproduction is by binary fission. Most species are known from acidic and high temperature environments such as hot springs or acidic sulfur fumes.

4.1.4.3 Subdivision Rhodophytina Class Porphyridiophyceae This class is probably not monophyletic and includes one order, the Porphyridiales which consists of one family, the Porphyridiaceae (5 genera, 10 species). It comprises coccoid unicellular species that are sometimes united in a common mucilage. They are uninucleate with a single central stellate

chloroplast (Fig. 4.13). The cells are usually able to move (by gliding). Their color ranges from blue-green to green or reddish. Sexual reproduction is unknown and asexual reproduction takes place by binary division. They occur in marine or freshwater but also in moist soils. The genus Porphyridium can also act as an endosymbiont of benthic foraminifers. Other genera are Erythrolobus, Flintiella, Rhodoplax, and Timspurckia. Class Compsopogonophyceae The class has three orders, the Comsopogonales (2 families, 4 genera, 10 species), Erythropeltales (2 families, 10 genera, 63 species), and the Rhodochaetales (1 species). In total, the class comprises some 73 species. They are all filamentous or saccate (shaped like a pouch or sack) and live in coastal seawater or freshwater. Their chloroplasts are parietal, discoid or ribbon-shaped, lobed and without pyrenoids. The order Compsopogonales includes three genera, including the well-known freshwater genus Compsopogon (Fig. 4.14). A few species of Compsopogon regularly occur in warm-water aquaria, probably introduced with vascular aquatic plants imported from warm regions. There are a few publications that report the occurrence of C. aeruginosus and C. coeruleus in European freshwaters, most probably from aquarium discharge (e.g., Müller 1960; Heynig 1971; Gärtner 1987; Breton 2014). Class Rhodellophyceae The class has only 7 species and is subdivided into the three orders Dixoniellales (3 genera and species), Glaucosphaerales (1 species), and the Rhodellales (2 genera and 2–3 species; Kamiya et al. 2017). All species are unicellular and sometimes united in a common mucilage. The cells are able to move (by gliding) and are uninucleate with one central stellate or peripheral lobed chloroplast. Cells are blue-green to reddish. Sexual reproduction is not known and asexual reproduction occurs by binary divisions. While the Dixoniellales and Rhodellales occur in marine to brackish water, the Glaucosphaerales occur in freshwater only. Class Bangiophyceae

2 µm

Fig. 4.12 Cyanidium caldarium TEM view of a cell with three endospores; Yellowstone National Park, USA, salty waters, 80 °C, survive desiccation and crystallization of salt. Non-layered thylakoids in plastids. © Science-Photo-Library, with permission

The Bangiophyceae comprise one order, the Bangiales and one family, the Bangiaceae that include some 200 species in 15 genera (Sutherland et al. 2011) and are, with the exception of the type genus Bangia, all marine. They occur as benthic organisms and occasionally epiphytic. Their life cycle is heteromorphic, the gametophyte (n) is foliose (8 genera) and one to two cell layers thick, or it can be an unbranched filament of uniseriate to multiseriate cells (7 genera). The cells contain one or, more rarely, two stellate

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Fig. 4.13 Porphyridium purpureum (SAG 1380-1d), United Kingdom, Scotland, Brixham Harbour, from marine Ulothrix enrichment culture. Courtesy of Tatyana Darienko with permission of the SAG, Univ. Göttingen

Fig. 4.14 Example of Compsopogonales. Compsopogon aeruginosus. a Filament with cortex (left) and non-corticated filament (center). b cortical cells with numerous parietal chloroplasts. Sample from a warm-water aquarium, Germany

a

b

50 µm

20 µm

(sometimes even more) plastids with a central pyrenoid; pit plugs are lacking. The spermatia are formed in special cells named spermatangia by division of the cell content. After being released, the spermatia fertilize female cells and the resulting zygote immediately divides into packets of zygosporangia that via meiosis release the zygospores, forming the sporophyte (2n). The sporophyte is a uniseriate filamentous “conchocelis phase” that can actively bore into shells. From the “conchocelis” filament, conchospores are

formed by mitosis. It is assumed that meiosis occurs upon germination of the conchospore, thus forming the new gametophyte. Order Bangiales Molecular phylogenetic studies have confirmed the monophyly of the Bangiales and the lineage containing both the Bangiales and the Florideophyceae (Oliveira and

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Bhattacharya 2000; Müller et al. 2001; Saunders and Hommersand 2004). The Bangiales comprise the highly valued seaweed aquaculture crops that have been sophistically cultivated, harvested, and traded in Japan, China, Korea, and Southeast Asia for thousands of years (Mumford and Miura 1988). The foliose thalli nowadays are also harvested in Chile, Wales, Pacific North America, and New Zealand (Turner 2003; Sutherland et al. 2011). Red algae of the order are found worldwide from tropical to polar seas, most species occur in the intertidal zone of coastal regions where they grow on rocks, shells, or other algae. Some are found solely in subtidal habitats. Worldwide efforts resulted in a more or less continuous addition of new taxa, demonstrating the still insufficient knowledge about their diversity and distribution (e.g., Sutherland et al. 2011). Until recently, only two genera were recognized in the Bangiaceae on the basis of their gametophyte morphology. Both were unbranched, the uniseriate to multiseriate filaments that were summarized in the genus Bangia and those with leaf-like thalli in the genus Porphyra. Recently, the new genus Pseudobangia with the new species P. kaycoleia, characterized by an unbranched filamentous growth, was added (Müller et al. 2005). Further new unbranched filamentous additions of the order were made with the genera Minerva and Dione (Nelson and Broom 2005). All unbranched filamentous taxa are characterized by bipolar spore germination, rhizoidal attachment cells internal to the cell wall, and a lack of pit connections between cells. The gametophyte thalli all are pseudoparenchymatous and have intercalary cell divisions (Sutherland et al. 2011). There is still an ongoing debate concerning the identity of the two species Bangia atropurpurea and B. fuscopurpurea (Fig. 4.15a, b). It had been thought that B. atropurpurea was the same species as B. fuscopurpurea and because B. fuscopurpurea (described in 1819) is the older name, B. atropurpurea (described in 1824) was synonymized with B. fuscopurpurea. However, doubts came up regarding whether or not the unification of the two species was justified. Firstly, culture studies revealed that B. atropurpurea exposed optimum growth in freshwater, whereas B. fuscopurpurea showed optimum growth in marine media (Belcher 1960). Secondly, a karyological and phylogenetic study revealed that the freshwater filaments of “B. fuscopurpurea” were all monosporic, had three chromosomes and a distinctive chromosome morphology in that the third chromosome is much smaller than those from marine 3-chromosome collections, thus positioning the freshwater Bangia species on a separate and well-supported branch in the phylogenetic tree (Müller et al. 2003). Consequently, the authors proposed the resurrection of the species B. atropurpurea to represent this lineage and suggested that, until final clarification, all freshwater species should be treated as B. atropurpurea and the marine species as B. fuscopurpurea (Müller et al. 2003).

B. Büdel and T. Friedl

Today, within the Bangiales fifteen genera are recognized of which seven belong to the filamentous and eight to the foliose taxa. The foliose taxa have been revised and the genus Porphyra is restricted to five described and several undescribed species (Fig. 4.15c–e). Other foliose taxa previously placed in Porphyra are now recognized as belonging to the genera Boreophyllum, Clymene, Fuscifolium, Lysithea, Miuraea, Pyropia, and Wildemania (Sutherland et al. 2011). Class Florideophyceae This is the most diverse group of the red algae with roughly 6,913 species, 94% of them live in the marine, others in brackish and/or freshwater habitats. The algae live benthically, epilithically and often also epiphytically on other algae or vascular plants, and 166 species from 66 genera are so far known to even live parasitically on other red algal hosts (Kamiya et al. 2017). The class is subdivided into 5 subclasses and *35 orders with the subclass Rhodymeniophycidae being the species richest with *5,200 species. Florideophycean algal thalli are always multicellular, filamentous and branched and often form pseudoparenchymatic structures that are cylindrical, compressed, or foliose. They grow from apical meristems, often supported from additional intercalary meristematic cells in more advanced genera. The cells can be uni- or multinucleate with two sibling cells joined by the characteristic pit plug. Non-sibling cells might be secondarily joined by secondary pit plug connections. Their life cycle is primarily triphasic (Figs. 4.8b, 4.9 and 4.10) and involves a free-living diploid tetrasporophyte and a haploid gametophyte generation with a fully or semi-parasitic diploid carposporophytic phase borne and dependent on the female gametophyte. Tetrasporophytes can be either morphologically similar (isomorphic) or dissimilar (heteromorphic) to the gametophytes (Kamiya et al. 2017). Here we discuss just a few selected orders, for a complete overview please refer to the specific literature, for example, Kamiya et al. (2017). Subclass Hildenbrandiophycidae Order Hildenbrandiales They all have crustose thalli with or without erect branches and are of a cartilaginous (gristly) consistency. Only one family exists, the Hildenbrandiaceae, with two genera and so far 19 species (Guiry and Guiry 2023). Most species occur in the marine environment in the tropical, temperate, and subpolar climatic zones, where they occupy rock substrata in the intertidal to subtidal zones. Some species also occur permanently or temporally inundated in brackish and/or

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b

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c

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Fig. 4.15 Examples of Bangiales. a Bangia fuscopurpurea (white arrow) and Porphyra sp. (black arrow), at the quay wall of Helgoland harbor, North Sea, Germany. b Bangia fuscopurpurea filaments from quay wall. c Porphyra cf. linearis population on sandstone wall, supralittoral zone, Helgoland, Germany. d Porphyra sp., large sample and e herbarium specimen of Porphyra umbilicalis, both from the North Sea littoral, Helgoland, Germany

freshwater, where they always live attached to the rock substratum (Fig. 4.16). Subclass Nemaliophycidae Order Acrochaetiales This order includes algae that live epilithically, epiphytically, or endophytically in marine or freshwater. Their thalli are heterotrichous, monosiphonous with branched or simple

filaments that are attached to the substratum by a single cell of a multicellular prostrate plate-like structure. The raised filaments are monopodial, either singular or often tufted (Fig. 4.17a). The cells are cylindrical or moniliform arranged and uninucleate with parietal or axial chloroplasts with or without one to several pyrenoids. Reproduction is monophasic via monospores (Fig. 4.17b), biphasic with reduced gametophytes or triphasic with morphologically similar or dissimilar gametophytes and tetrasporophytes. Two families are known with presently 224 species. The

120 Fig. 4.16 Hildenbrandiales; a Hildenbrandia sp. from freshwater, on rock surface of the Debengeni waterfalls, Magoebaskloof, Limpopo province, South Africa. b Hildenbrandia rubra on rock from the marine environment, North Sea, Helgoland, Germany. c Hildenbrandia rivularis, covering a quartz rock, Germany; d longitudinal section of the thallus showing the vertical arrangement of the upright filaments. Courtesy of Antje Gutowski, AlgaLab (c, d)

B. Büdel and T. Friedl

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b

3 cm

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c

~1 cm

d

Fig. 4.17 Acrochaetiales. Audouinella cf. hermannii, a tufts (arrow) growing on a vascular aquatic plant leaf in an aquarium; b filament cylindrical cells with parietal chloroplasts and a monospore (arrow)

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1 cm

family Acrochaetiaceae includes almost all species in 12 genera, while the family Ottiaceae is so far known to have one species, Ottia meiospora (Guiry and Guiry 2023). All 45 freshwater species are united in the mostly epiphytic genus Audouinella, that is characterized by producing monosporangia and parietal laminate or ribbon-shaped chloroplasts (Fig. 4.16). Order Balbianiales Members of this order occur in freshwater and form uniseriate, monoecious, and branched filaments. They live

10 µm

epilithically (Rhododraparnaldia) or epiphytically (Balbiania), mostly on members of the family Batrachospermaceae (e.g., Leukart and Knappe 1995). Their cells contain several chloroplasts. Only one family exists, the Balbianiaceae with the two genera, Rhododraparnaldia with one species, and Balbiania with two currently accepted species (Guiry and Guiry 2021). Their triphasic life cycle is characterized by (1) gametophyte carpogonia that have an elongate and thin trichogyne, (2) their carposporophytes that resemble gametophytes with tetrasporangia on short branches, and (3) sporophytes that produce monospores 7–9 µm in diameter (Fig. 4.18).

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Fig. 4.18 Balbianiales. a Balbiania investiens (arrows), growing on Batrachospermum sp. Filaments. Courtesy of Jo Wilbraham, taken from AlgaeVision. b, c, d Balbiania investiens, from the river Ocker, Germany; b tip of Batrachospermum thallus with epiphytic B. investiens (arrows); c, d germinating carpospores with filament production. Pictures b– c courtesy of Antje Gutowsky, AlgaLab

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a

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d 10 µm 10 µm

200 µm

Order Batrachospermales The red algal order Batrachospermales so far comprises some 260 species in one family, the Batrachospermaceae (Entwisle et al. 2009) with 12 genera. All known species occur in freshwater only. The first phylogenetic study on the Batrachospermales found the well-known genus Batrachospermum to be highly paraphyletic (Vis et al. 1998). Expanded studies of taxa with DNA sequence data have established many new genera formerly all included within the genus Batrachospermum. So far, the genera Kumanoa, Sheathia, Torularia, Virescentia, Acarposporophycos, Visia (Entwisle et al. 2009; Necchi and Vis 2012; Necchi et al. 2018, 2019a, b; Rossignolo and Necchi 2016; Salomaki et al. 2014), Lympha (Evans et al. 2017), Nocturama (Entwisle et al. 2016) and Volatus (Chapuis et al. 2017) have been newly described out of the formerly single genus. With the most recent description of the new genus Montagnia that previously belonged to Batrachospermum, the paraphyletic complex seems to have been resolved (Necchi et al. 2019b). The family is characterized by heterotrichous thalli with a heteromorphic life cycle that is triphasic, with a haploid gametophyte generation, diploid carposporophyte, and a diploid sporophyte generation (“Chantransia” stage, looks like the uniseriate genus Audouninella). The pit connections between neighboring cells have two pit plug cap layers, other layers are enlarged. Other common genera, apart from the former Batrachospermum complex, are for example Balliopsis, Lemanea, Paralemanea, Psilosiphon, and Sirodotia. Batrachospermum and its related genera are composed of one main axial filament with or without cortical

filaments originating from the nodia (Fig. 4.19e). The nodia occur in regular sequences and are the origin of the nodular whirls composed of filaments (Fig. 4.19d), thus creating the characteristic habitus of the alga and the reason for the German name “Froschlaichalge”, meaning that it looks like the cords of frog spawn (Fig. 4.19a–c). The genera Lemanea, Paralemanea, and Psilosiphon have tubular, cartilaginous, and pseudoparenchymatous thalli with a smooth outer cortex and inner medullary layer, the latter is often composed of only loosely interwoven filaments (Figs. 4.4c and 4.19f–h). Order Palmariales The order of the Palmariales consists of the four families Meiodiscaceae (4 genera, 8 species), Palmariaceae (4 genera, 23 species), Rhodophysemataceae (5 genera, 11 species), and the Rhodothamniellaceae (2 genera, 3 species). Their life history consists of either an alternation of generations with macroscopic tetrasporophytes, male gametophytes, and microscopic female gametophytes or by directly formed tetrasporangia, presumably derived by an apomeiotic process (see glossary). Thalli of the family Meiodiscaceae are filamentous, forming a monostromatic pseudoparenchymatous disc or an endozoic network of anastomosing filaments. The family Palmariaceae thalli are solid or hollow, elliptical, palmate, or tubular. The cortical cells are in one or more layers and the medullary cells are large and spherical to subspherical and also in one or more layers. A well-known representative of this family is the edible seaweed Palmaria palmata (Fig. 4.24f).

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0.5 cm

c 1 mm

1 cm

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f

g

h

1 cm 1 cm

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Fig. 4.19 Batrachospermales. a Herbarium specimen of Batrachospermum sp., calcareous freshwater creek, Germany. b Thallus of Batrachospermum gelatinosum, freshwater creek in sandstone, Spessart midlands, Germany. c B. gelatinosum from a tufa cascade of a little creek, Germany; calcium carbonate depositions on whirls. d, e Batrachospermum sp., Letsitele River, Limpopo Province, South Africa; main axis with nodial whirls and internodial main axis (white arrows) coated with cortical filaments (black arrows). f–h Lemanea cf. rigida, freshwater creek, Spessart midlands, Germany; herbarium specimen (f), subaqueous view of the thalli bases (g), thalli growing on sandstone rock take off the river (h)

Order Thoreales This order has one family, the Thoreaceae with the two genera Thorea (17 species) and Nemalionopsis (3 species). The gametophytes are large and rope-like, 5–20 cm (rarely 2 m) long with a diameter of 0.5–3 mm. They are multiaxial, differentiated into an inner medulla composed of numerous colorless, heavily interwoven filaments. The outer part

persists of numerous regularly out-bound, short assimilation filaments (Fig. 4.20). Sexual reproduction is only known from Thorea, Nemalionopsis that is currently known to only reproduce by monospores. The carpogonial branch of Thorea consists of the carpogonium only, with a trichogyne that has an expanded base. The gonimoblast filaments form carposporangia, sometimes monosporangia are also produced. From the carpospores, a branched, uniseriate

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a

b ~1000 µm

0.5 cm

Fig. 4.20 Thoreales. a, b Thorea hispida, habitus as found floating in the river Oker in Braunschweig, Germany (a); detail showing the assimilatory filaments originating perpendicular to the man filaments axis (b)

Chantransia-stage emerges, producing an attached gametophyte via meiosis. The spermatangia cluster on the assimilatory or on specialized filaments. Tetrasporangia are not formed. They grow epilithically in freshwater rivers and channels at depths of 3 m, sometimes they may be found free-floating. Subclass Corallinophycidae The subclass Corallinophycidae includes the four orders Corallinales (7 families, *24 genera, 600 species), Hapalidiales (2 families, *16 genera, 117 species), Rhodogorgonales (1 family, 3 genera, 4 species), and the Sporolithales (1 family, 4 genera, 44 species) and harbors in total roughly 770 species. Calcification of the cell walls is a typical feature for the whole subclass. The members of the Corallinales either live as epiliths, epiphytes or partly or entirely endophytes. Three major growth forms can be differentiated; the two non-geniculated types of rhodoliths and crustose thalli, and those with geniculate-articulated thalli (McCoy and Kamenos 2015). The rhodoliths have a rock-like structure, but are formed by red algae that grow free and unattached (Figs. 4.21c, d and 4.25). The articulated thalli are realized with branches composed of alternating uncalcified ring-like sectors (= genicula) and elongated calcified sectors (= intergenicula; Fig. 4.21a, b). Non-geniculate thalli are completely calcified. The thalli are of multiaxial and pseudoparenchymatous structure. Some are composed of unconsolidated filaments with meristematic cells either located terminal or intercalary. The meristematic cells produce epithelial cells (cortex) with partly calcified walls outwardly and vegetative cells inwardly. Gametophytes are either monoecious or dioecious. Order Corallinales The Corallinales include the single family Corallinaceae, comprised of 42 genera with some of them having an unclear phylogenetic status. Well-known genera are Corallina with

so far 28 species, Lithothamnion with 83 species, and Lithophyllum with 126 accepted species. Order Hapalidiales The Hapalidiales are epilithic, free-living as rhodoliths or partly or entirely endophytes. They are non-geniculate and calcified, and do not have distal walls of epithelial cells. The Hapalidiaceae is the only family with 56 supported species. Probably the best-known genus is Phymatolithon with 21 accepted species (Jeong et al. 2019) which includes the prominent, Maerl-forming species P. calcareum (Figs. 4.21e and 4.26). New genera being still described and defined within this order, for example, recently the genus Phymatolithopsis with three species. Their geographical range is from Southeast Atlantic and Indian Oceans and the temperate Northwest and Southwest Pacific Ocean (Jeong et al. 2022). Subclass Rhodymeniophycidae Order Ceramiales This species rich and marine habitat restricted order (roughly 2,700 species) comprises 5 families, the Callithamniaceae (37 genera, 210 species), Ceramiaceae (*50 genera, 450 species), Delesseriaceae (*100 genera, 640 species), Rhodomelaceae (*125 genera, *1,100 species), and the Wrangeliaceae (*56 genera, 300 species). Thallus morphology varies from small filamentous tufts to delicate membranous and large foliose thalli, which are sometimes cartilaginous (Fig. 4.22c, g). The life cycle is triphasic with isomorphic gametophytes and sporophytes. The tetrasporangia are tetrahedrally or cruciformally divided and borne on whorl or lateral branches. The carpogonial branches are 4-celled and are borne on supporting cells. After fertilization, auxiliary cells develop from the supporting cells, or the supporting cells act directly as auxiliary cells. The diploidization of the auxiliary cells occurs by the

124 Fig. 4.21 Corallinales. a Corallina elongata, lower tidal zone, Atlantic coast Brittany, France. b Corallina sp., mediterranian coast at Cala Montgo, Spain; thallus branches exposing the geniculate nature with alternating calcified filaments and uncalcified genicula (arrows). Hapalidiales. c Phymatolithon calcareum, calcified thalli forming Maerl, coastal region, Roscoff, France. Corallinales. d Lithothamnion glaciale, rhodolith forming red alga in 20–70 m depth of Mosselbukta in the arctic Svalbard Archipelago; courtesy of Solvin Zankl. e Lithophyllum incrustans, covering rocks in the tidal zone, Roscoff, France

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development of a connecting cell arising from the fertilized carpogonium, or by fusion of the carpogonium and the fusion cells. The gonimoblast arises from an auxiliary cell. Asexual reproduction is by unicellular or multicellular propagules. The family Rhodomelaceae represents the genus and species richest family within the order and comprises species with usually erect, multiply branched or in some genera foliose and/or prostrate thalli (Fig. 4.22c, f, g). The Delesseriaceae often have attractive thalli composed of foliose phylloids, formed by primary and secondary filaments (Figs. 4.4d, e and 4.22d, e), having a more or less pronounced cauloid (stem) and are attached to the substratum by a discoid holdfast (rhizoid). The Ceramiaceae form thalli that are usually erect and uniaxial. Each axial cell bears two or numerous normal or reduced determinate laterals or whorl branchlets. Their cells are uninucleate, and the gametophytes are dioecious. The filaments of Ceramium are small, growing to no more than 30 cm. They consist of a uniseriate axis of cells surrounded by smaller cells forming a cortex (Fig. 4.22a). The genus Pterothamnion in contrast

forms thalli that are composed of a uniseriate main axis alternately branched in one plane; branching occurs in the upper part of the axial cells (Fig. 4.22b). Order Plocamiales The Plocamiales are red algae that live as epiliths, epiphytes, or parasites and are exclusively found in marine habitats. In terms of species richness they are a small group. They include 2 families, the Plocamiaceae (3 genera, 48 species) and the Sarcodiaceae (3 genera, 25 species). Their life cycle is triphasic with isomorphic gametophytes and tetrasporophytes. The thalli are filamentous, uniaxial, or ramisympodial (bifurcate branching, see glossary) and the carpogonial branches are 3-celled and are borne by an intercalary cortical supporting cell that also functions as the auxiliary cell. The thalli of the Plocamiaceae consist of a stoloniferous (producing or bearing stolons) base and a complanate (put into one plane) erect thallus with alternating marginal series of 2– 6 compressed lateral branches (Fig. 4.23).

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Fig. 4.22 Ceramiales. a Ceramium cilliatum, habitus of sparsely branched thallus, inset: close up of the complex filament with large celled main axis and small cortical cells, coastal Britanny, Roscoff, France; courtesy of Solvin Zankl. b Pterothamnion plumula, habitus of terminal branch, intertidal, Helgoland, Germany. c Laurencia botryoides, herbarium specimen, intertidal, Indian Ocean, West Australia. d, e Delesseria sanguinea, Herbarium specimen (d) and living material, North Sea, Helgoland, Germany. f Vertebrata fucoides, gametophyte with carposporophytes (arrows), intertidal, Roscoff, France. g Osmundea pinnatifida, cartilaginous thallus, intertidal, coastal Britanny, Roscoff, France

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Order Gigartinales The Gigartinales are a species-rich order of marine red algae with roughly 940 species in 36 families and 170 genera. Apart from the genus Rhodopeltis, their thalli are not calcified. They are monoecious or dioecious and are characterized by an alternation of isomorphic and heteromorphic generations. In heteromorphic species the gametophyte is macroscopic. Generally, their thalli are erect or crustose,

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terete, flattened or foliose, branched or unbranched and sometimes proliferous. The members of the Dumontiaceae family have erect, terete to flattened, radially to distichously branched, or foliose and unbranched thalli. The thalli are generally mucilaginous, the thallus structure is uniaxial, rarely multiaxial with a filamentous medulla and cortex (Fig. 4.24b). The Dumontiaceae include 19 genera with about 70 species. The Gigartinaceae comprise 9 genera with about 150

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Fig. 4.23 Plocamiales. Plocamium cf. cartilagineum, coastal Britanny, Roscoff, France

species. Their thalli are erect and grow from a discoid or crustose base. The mature thalli are cylindrical, compressed or flattened and dichotomously or pinnately branched or unbranched. Species of the genus Chondracanthus (e.g., C. exasperatus, the Turkish towel, Fig. 4.24e) can reach a size of 1 m or more. The species Chondrus crispus, colloquially named Irish moss, is widely known because of its cultivation to gain the polysaccharide carrageenan (Fig. 4.24c). The family of the Phyllophoraceae consists of 15 genera with 134 species. They are monoecious or dioecious, the life cycle is iso- or heteromorphic. The gametophyte thalli are erect, terete, compressed, or foliose (Fig. 4.24a) and the tetrasporophyte of heteromorphic species is crustose. Some species lack an independent tetrasporophyte, their tetrasporangia develop on the female gametophyte in place of carposporangia. The species Mastocarpus stellatus, the false Irish moss, is morphologically closely related to Chondrus crispus, but has typically channeled fronds (Fig. 4.24d). Both species overlap in their geographical distribution.

4.1.5 Genome Reductions and Gains: The Ecological Imprint A remarkable feature of red algal evolution is their relatively limited gene inventory (*5–10 thousand genes) compared to those of other free‐living algae. Apparently, the common ancestor of red algae has undergone extensive genome reductions, probably resulting in the specialization of certain lineages to a symbiotic or parasitic lifestyle, or may have adapted to an extreme or oligotrophic environment. In order to study this important aspect, genome evolution was investigated using 14 red algal genera representing the major branches of the Rhodophyta (Qiu et al. 2015). In this

analysis of gene gains and losses, two putative major phases of genome reduction were identified: (1) in the basal lineage of all red algae, where genome reduction leads to the loss of flagellae and basal bodies, the glycosyl‐phosphatidylinositol anchor biosynthesis pathway, and the autophagy regulation pathway; and (2) in the common ancestor of the extremophilic Cyanidiophytina. In addition, red algal genomes are also characterized by the recruitment of hundreds of bacterial genes through horizontal gene transfer that have taken on multiple functions in shared pathways and have replaced eukaryotic gene homologs. The results of Qiu et al. (2015) suggest that Rhodophyta may trace their origin to a gene depauperate ancestor. It appears that a limited gene inventory is sufficient to support the diversification of the major eukaryote lineage of the red algae that are characterized by sophisticated multicellular reproductive structures and an elaborate triphasic sexual cycle. A comprehensive study on the genome of the florideophyte Chondrus crispus (Irish Moss, Fig. 4.24c) sheds lights on the early evolution of the Archaeplastida but also helps to delineate the innovations necessary for the emergence of aero-terrestrial algae and other land plants and their adaptation to the terrestrial environments (Collén et al. 2013). The presence of two cellulose families (glycoside hydrolase 5 and 45, GH 5 and 45) in C. crispus for example, supports the assumption that the ancestor of the Archaeplastida was a protist feeding on exopolysaccharides such as cellulose. This is consistent with the ancient origin of family GH9 cellulases in eukaryotes (Davison and Blaxter 2005). The red algae only kept GH5 and GH45 cellulases while green algae and plants lost these genes and conserved GH9 cellulases. This includes that cellulose biosynthesis was acquired independently in red and green algae, and that this independent acquisition can partially explain the structural diversity of cellulose-synthesizing enzyme complexes and cellulose microfibrils in Archaeplastida (Tsekos 2002). It was suggested by Collén et al. (2013) that the red algal lineage went through an evolutionary bottleneck. Early in their evolution, but after their divergence from green algae, selective pressure for small physical size or low-nutrient requirements probably caused a reduction of the genome, with loss of introns and intergenetic material. This might also explain the lack of flagella in all life-cycle stages in red algae as a result of the loss of the referring genes. The unicellular red algae Cyanidioschyzon merolae or Galdieria sulphuraria live in a high temperature-low pH environment, and for some reason, these conditions favor compact genomes in red algae, probably indicating that the ancestral red algae were acido- and thermophilic organisms. The evolutionary bottleneck also might explain the high numbers of orphan genes in the genome, as the more derived red algae were forced to reinvent gene functions that were lost during the genome reduction (Collén et al. 2013).

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4.1.6 Ecology Red algae are the most diverse seaweeds in the marine environment, only a few occur in freshwater and with a very few exceptions of small coccal taxa, they never made their way into aero-terrestrial habitats. It seems that they are the great “losers” of photoautotrophic thalloid eukaryotes (Frazer 2015). Desiccation tolerance in order to survive drought conditions is not well understood in macrophytic marine red algae. Many of them grow in the intertidal zone on rocky shores where they experience more rapid and severe losses of water than most vascular plants. An investigation on the two thalloid species Porphyra umbilicalis and P. yezoensis revealed quite different tolerances to desiccation. Both species loose about 95% of their thallus water content in the first two hours of dehydration and their final relative water content is virtually the same. However, massive membrane leakage, reduced respiration and reduced oxygen evolution was observed in P. yezoensis after desiccation, but not in P. umbilicalis, and extensive membrane disruption was found in P. yezoensis after desiccation, but not in P. umbilicalis. Reactive oxygen species defense, the repression of membrane phase transition, and formation of cellular glass (vitrification) are the three major desiccation tolerance mechanisms known from land plants. However, light has been found to stimulate reactive oxygen species damage in P. yezoensis, while the mechanisms providing a higher desiccation stability of P. umbilicalis remains unclear. The latter species probably has a more stable vitrification when desiccated and a lower molecular mobility during the drying process (Liu 2009).

4.1.6.1 Freshwater The roughly 200 freshwater red algal species are generally more diverse in tropical rather than in temperate and polar zones. However, some warm-water taxa, for example, Compsopogon and Thorea, have made their way to temperate European environments during warm summers (Eloranta et al. 2011). Compsopogon occurs regularly in warm-water aquaria, apparently throughout the world. Microscopic coccal red algae of the genera Cyanidium, Rhodospora, Chroothece, and Porphyridium commonly grow terrestrially on moist soil and rock, but also in peat bogs. Hot acid springs are the typical environment of the genera Cyanidium and Galdieria while Porphyridium species prefer habitats with high nutrient availability. The microscopic and short filamentous species of the genera Chroodactylon and Kyliniella occur on rocks and epiphytically in standing waters. The epiphytic genus Balbiania is known only epiphytically from Batrachospermum species (B. gelatinosum and B. keratophytum; Starmach 1977).

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The typical habitat for macroscopic freshwater red algae is running waters, but some species in genera such as Batrachospermum and Hildenbrandia can also be found in lakes and ponds. In temperate North America and Europe maximum growth of freshwater red algae is reached from late autumn to early summer. However, when water temperature remains low (7.0, but many taxa prefer soft and acidic water (Eloranta et al. 2011). Apparently, this type of distribution pattern is associated with the ability of the species to use either free carbon dioxide as the only inorganic carbon source and the typical carbon source of acidic water, whereas others can make use of bicarbonate and carbonate, the characteristic inorganic carbon sources of alkaline water (Raven and Berdal 1981; Raven et al. 1982, Eloranda et al. 2011).

4.1.6.2 Marine The vast majority of red algal taxa are macroscopic forms occurring in marine water bodies where they are distributed along rocky shores of the oceans, colonizing rocks or other

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solid substrata down to depths of more than 250 m. More than 100 species grow free floating or entangled in other substrates (vegetation, natural or anthropogenic litter) and many species also occur epiphytically on other macroalgae or seagrasses. As there is a large body of publications available (see for example regional and general floras of red algae), we restrict ourselves here to the relatively recent field of rhodolith research. Rhodoliths are mostly red or reddish crustose benthic marine algae that in shape resemble corals but are constructed by red algae depositing calcium carbonate in their cell walls, forming hard rocky structures. Rhodoliths Coralline red algae which form rhodoliths in Polar Regions have received little attention concerning their potential as ecosystem engineers and carbonate factories. They were recently found to be much more widespread in polar waters than previously thought. The northernmost rhodolith communities were discovered in 2006 at Nordkappbukta (North Cape Bay, 80° N) at Nordaustlandet, Svalbard (Fig. 4.25). The perennial coralline algae experience extreme seasonality with a light regime including 4 months of winter darkness, and highly varying sea ice coverage, nutrient supply, turbidity of the water column, temperature, and salinity. In addition to intensive field and laboratory research, the rhodolith communities and their environment were explored by means of a manned submarine (JAGO). The coralline flora was composed mainly of Lithothamnion glaciale together with a lesser amount of Phymatolithon tenue. The rhodolith communities occurred between 30 and 51 m, while coralline algae attached to cobbles were present as deep as 78 m. The surrounding waters were always saturated with calcite and

Fig. 4.25 Rhodoliths in the arctic sea, Mosselbukta, Svalbard Archipelago. Courtesy of Solvin Zankl with permission of Natur-Forschung-Museum (Wisshak 2019)

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aragonite for the whole area. The rhodolith-associated macrobenthic fauna determined was large and included 59 species. The concomitant appearance of corallines and grazers apparently keeps the corallines free from epiphytes and coequally provides an optimal food source for the grazers (Teichert et al. 2012). Maerl Maerl is a Breton word that describes sand or gravel on the sea floor at sublittoral sites, which is formed by the calcified remains of red algae (rhodoliths). It is distributed in the Mediterranean, along the Atlantic coast (Fig. 4.26) from Portugal to Norway, into the English Channel, the Irish Sea, and the North Sea. In Brittany, maerl beds form a > 5500-year-old habitat. However, maerl has been so heavily used as an agricultural fertilizer, that it is considered to be a threatened habitat in Brittany (Grall et al. 2003). Commonly recorded maerl-forming species in the NE Atlantic are Lithothamnion corallioides, L. glaciale, L. tophiforme and Phymatolithon calcareum, but maerl formed by Lithophyllum spp. is also occasionally found. Lithophyllum fasciculatum forms globular to sub-globular unattached thalli up to 10 cm in diameter and forms maerl beds at 0–2 m depth in the eastern part of the Bay of Brest, France. Apart from maerl beds in Brittany, L. fasciculatum maerl has also been recorded in Kingstown Bay, Ireland where it occurs at 0–3 m depth. There, L. fasciculatum co-occurs with the closely morphologically similar maerl species L. dentatum. The taxonomy of NE Atlantic maerl-forming Lithophyllum spp. is considered as somewhat messy, and their ecology has not yet been studied in detail (Peña et al. 2013).

Fig. 4.26 Maerl bed at several meters depth, Atlantic Ocean, Falmouth, south west UK. The Maerl beds are mainly shallow, between 5 and 9 m. Courtesy Mark Milburn, UK

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4.1.6.3 Ecophysiology Depth zonation and photosynthesis The red algae are distributed from the upper intertidal to the deep littoral zones. Crustose coralline algae have even been observed at a depth of 268 m, where they receive less than 0.001% of the surface irradiance (Littler et al. 1986). For the temperate marine red algae Chondrus crispus and Porphyra purpurea light compensation points for growth were found to be as low as 0.4–1.8 µmol m−2 s−1 PAR (at 7 °C), respectively (Markager and Sand-Jensen 1992), which is among the lowest light compensation points known for cryptogamic organisms (see also Green et al. 1991). The researchers found that there is no surplus of energy to balance grazing and mechanical losses, and these factors must be of minor importance for macroalgae growing at great depths. A literature review found depth limits of about 0.5% of surface light for marine macroalgae of the upper zone including mainly leathery algae, followed by an intermediate zone of foliose and delicate algae with depth limits at about 0.1% surface light, and a lower zone of encrusted algae extending down to about 0.01% surface light (Markager and Sand-Jensen 1992). The carbon source of red algae is either HCO
3 or CO2. The two stable carbon isotopes on Earth are 12C (99%) and 13 C (1%). Because of a number of kinetic and equilibrium processes that discriminate differentially against the heavier isotope, the natural abundance of these isotopes in the organic carbon of plants differs from that in the source (Farquhar et al. 1989). The major potential discriminant in carbon fixation is the carboxylation reaction carried out by RUBISCO. Consequently, algae that mainly assimilate 13 HCO
3 should have less negative d C values than those that exclusively use CO2. This theoretical consideration was confirmed in two ecologically different populations of red alga (together with green and brown algae), those from below the lowest tide line, which are never exposed to atmospheric 13 carbon and can use HCO
3 , had d C values in the range of −9.6 to −22.6‰, while 6 red algal species, which are probably only able to use CO2, had d13C values in the range of −29.9 to −34.5‰ (Maberly et al. 1992). This method elegantly allows not only recognition but also discrimination, at least to some extent, between red algal taxa along a depth gradient and their possible photosynthetic performance.

4.1.7 Phylogeography The red algae are diverse in pantropical regions and relatively sparse in polar seas. A number of molecular studies discuss the origin of many red algal lineages or extant members of lineages in the southern hemisphere and suggest

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that many species are actually species complexes. It is evident that the resolution of species boundaries is not clearly evident from morphological examination only. On the basis of morphological identifications, many red algal species are reported to have circumglobal distributions. As mentioned before, molecular data have revealed a range of issues with morphologically defined species boundaries. Consequently, the real distribution of such circumglobal species must be questioned. A recent study investigated distribution patterns of nine species of the Rhodomelaceae by molecular data sets (rbcL gene) from widely spaced geographical locations. The authors identified three distinct patterns (Fig. 4.27): (1) Species with a strong phylogeographic structure (e.g., phylogenetic similarity correlates with geographical provenance) often in the pattern that populations from different locations were considered as different species (Lophosiphonia obscura, Ophidocladus simpliciusculus, Polysiphonia villum, and Xiphosiphonia pinnulata). (2) Species with a broad distribution partly explained by putative human‐mediated transport (Symphyocladia dendroidea and Polysiphonia devoniensis). (3) Non‐monophyletic complexes of cryptic species, most of which have a more restricted distribution than previously thought (Herposiphonia tenella, Symphyocladia dendroidea, and the Xiphosiphonia pennata complex that includes the species Xiphosiphonia pinnulata and Symphyocladia spinifera). The species Polysiphonia devoniensis was originally described from southern England (Maggs and Hommersand 1993) and later recorded from other locations in Atlantic Europe (Díaz-Tapia and Bárbara 2013). More recently, P. kapraunii was described from North Carolina (Stuercke and Freshwater 2010). The phylogenetic analysis revealed that these two species form a clade and are closely related. It is unclear so far whether or not they should be considered a single species. This study is a nice example showing that widely distributed species are the exception in marine red algae, unless they have been spread by humans recently, thus explaining their modern disjunctive distribution (Díaz-Tapia et al. 2018). Red algae of the genus Porphyra sensu lato (Bangiaceae) are important seaweeds that are also used in aquaculture for human food production. The genus is genetically the most diverse group of the Bangiophyceae. On the basis of a genetic study, the biogeographic population structure in two abundant Porphyra species in the Northeast Atlantic was investigated recently (Varela-Álvarez et al. 2022). Investigated were the dioecious annual species Porphyra dioica and the protandrous, hermaphroditic and winter annual species Porphyra linearis. Both species occupy distinct niches. Firstly, P. linearis exposed a strong genetic differentiation of the north-central eastern Atlantic populations,

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Fig. 4.27 Distribution and unrooted distance phylogram based on rbcL sequences of Polysiphonia villum (a, b) and P. devoniensis (“P. kapraunii”in North Carolina) (c, d). In panels a and c, circles indicate the regions from which sequences are available, colors indicate the distribution of each haplotype when multiple haplotypes were present. Areas outlined in red are regions where the species is recorded for the first time. Coastline in black shows the reported distribution of P. villum (a) and P. devoniensis (c), red coastlines that of “P. kapraunii” (c), (Guiry and Guiry 2022). Courtesy of Díaz-Tapia et al. (2018) with permission from Wiley & Sons Publisher

occurring from Iceland to the Basque region of Northeast Iberia. However, the southern populations, occurring from Galicia in Northwest Iberia and Portugal, showed a higher genetic diversity in the south, while a lower diversity was characteristic of the northern, homogenous population. Secondly, for P. dioica a weaker genetic differentiation was found and all the haplotypes mixed across its distribution. From their results, the researchers hypothesized a northward colonization from southern Europe, where the ancestral populations reside for P. linearis. But, for P. dioica the signature of this colonization is not as obvious due to hypothetical higher gene flow among populations. The latter probably linked to its reproductive biology and annual life history (Varela-Álvarez et al. 2022).

4.2

Chloroplastida—Green Algae

Frederik Leliaert, Olivier De Clerck, Tatyana Darienko, Jan de Vries, Thomas Friedl The Chloroplastida, also known as Viridiplantae (“green plants”) or Chlorobionta, are arguably the most dominant group of primary producers on earth. They include green

algae (containing around 500 genera and 8,000 extant species) and embryophytic land plants (Fig. 4.28). Green algae have played a fundamental role in the global ecosystem for hundreds of millions of years. The evolution of embryophytic land plants from a green algal ancestor in the mid-Ordovician was a key event in the history of life and has led to dramatic changes in the earth’s environment by initiating the development of terrestrial ecosystems (Kenrick and Crane 1997). The green algae appear to form a natural division, well-differentiated from all other groups of algae. It is much more difficult, however, to separate green algae from the embryophytic land plants. Because of their shared evolutionary history, many defining features are also shared with land plants, or at least those plants with flagellate stages in their life cycle. The green algae were traditionally termed Chlorophyta, but it has become clear from phylogenetic studies that separating the embryophytic land plants leaves the Chlorophyta paraphyletic (Leliaert et al. 2012). Thereto, the traditional division Chlorophyta has been split into Chlorophyta sensu stricto and the Streptophyta. Recently a third phylum was recognized, the Prasinodermophyta (Li et al. 2020). The Streptophyta includes the land plants and a paraphyletic

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Fig. 4.28 Evolutionary relationships among the main (class-level) lineages of Chloroplastida. The tree shows the “green algae” as a paraphyletic group. Together with the embryophytic land plants, they form a monophyletic group, the Chloroplastida

assemblage of green algae, termed charophytes or streptophyte algae grade (Figs. 4.28 and 4.69).

4.2.1 Ecological and Economic Importance Green algae are ecologically important in aquatic environments worldwide. They are especially abundant and diverse as microalgae and macrophytes in freshwater environments such as lakes, ponds, streams, and wetlands, where some may form nuisance blooms when confronted with excessive nutrients. Other green algal groups are well represented in marine environments. These include the green seaweeds, macroscopic green algae which are abundant in coastal

environments. Some green seaweeds (e.g., Ulva) can form extensive, free-floating coastal blooms, called “green tides” (Gao et al. 2017); other genera including Caulerpa and Codium are infamous for their invasive nature (Verlaque et al. 2003; Provan et al. 2008). Another group of green algae that is widespread in marine habitats are the prasinophytes, which are planktonic algae that are mainly found in oceanic environments and are especially abundant in more eutrophic, near-shore waters, where they can form monospecific blooms (Guillou et al. 2004). Several green algae are also found in highly specialized and extreme environments, such as deserts, hypersaline habitats, acidic waters, and mesophotic marine waters. Others have adapted to living in snow or ice. Some green

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algal groups (e.g., Trentepohliales) are exclusively terrestrial and never found in aquatic environments. Several lineages engage in symbiosis with a diverse range of eukaryotes. The association of green algae to associate with fungi to form lichens is well known, but green algae are also found as endosymbionts in certain ciliates, foraminifera, cnidarians, molluscs, and even vertebrates (Fig. 2.8, see Chap. 6). The large majority of green algae are photoautotrophic. Mixotrophy, however, is fairly common among green algae. Numerous green algae are capable of supplementing photosynthesis by uptake and utilization of exogenous dissolved organic carbon, such as sugars, amino acids, and other small molecules (osmotrophy). Other species (e.g., Pyramymonas, Cymbomonas) even supplement photosynthesis with phagotrophy (Bell and Laybourn-Parry 2003; Maruyama and Kim 2013). Several green algae have served as model systems in scientific research. For example, Melvin Calvin used Chlorella to study the light-independent reactions of photosynthesis, now known as the Calvin cycle. Joachim Hämmerling, by conducting transplant experiments with the giant-celled Acetabularia, showed that the nucleus of a cell contains the genetic information that directs cellular development. Studies using Chlamydomonas, a unicellular green flagellate, were pivotal in the first half of the twentieth century, demonstrating that different genes were responsible for different enzymatic reactions, thereby laying the foundation of molecular genetics (Salomé and Merchant 2019). In addition, green algae have served as model organisms by combining experimental and genomic data for studying ancient eukaryotic features such as the function and biogenesis of chloroplasts, flagella, and eyespots, and regulation of photosynthesis (e.g., Chlamydomonas reinhardtii), symbiosis (e.g., Chlorella), multicellularity and cell differentiation (e.g., Volvox, Ulva). Genomes of charophyte green algae are important sources of information for studying the origins of plant traits because of their evolutionary relationship to land plants (Cheng et al. 2019). Several green micro- and macroalgae are commercially important. Dunaliella salina is one of the commercially most important green algae because of its high b-carotene content. Haematococcus pluvialis (Fig. 4.1) is cultivated for its production of astaxanthin. This carotenoid is used as an antioxidant food supplement or as a food coloring, for example, in salmon aquaculture (Ambati et al. 2019). Some green microalgae, including Dunaliella salina, Ettlia oleoabundans, and Botryococcus braunii, contain high amounts of lipids and are potentially valuable for producing biofuels (Chen et al. 2011). Some green seaweeds, including Ulva, are used as fertilizer for agriculture. Others are harvested or cultivated for utilization in various food preparations, mainly in East and South East Asia, including Ulva reticulata, Monostroma oxyspermum (green laver or aonori),

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and Caulerpa lentillifera (sea grapes or lato). Some green seaweeds, including species of Aegagropila, Caulerpa, and Halimeda, are sold as ornamental macrophytes for marine or freshwater aquaria, which brings a risk of the introduction of alien species (Vranken et al. 2018). A few green algal species are pathogenic, including heterotrophic Prototheca species causing infections in humans and farm animals known as protothecosis (Lass-Flörl and Mayr 2007). Cephaleuros and Stomatochroon (Trentepohliales) are notorious plant pathogens causing algal red rust diseases, damaging economic crops worldwide (Joubert and Rijkenberg 1971).

4.2.2 Origin of Green Algae The Chloroplastida originated following an endosymbiotic event. A heterotrophic eukaryotic cell engulfed a photosynthetic cyanobacterium-like prokaryote that became stably integrated and eventually evolved into a membrane-bound organelle, the plastid (see Chap. 2). This single event marked the origin of photosynthetic eukaryotes and gave rise to three extant lineages with primary plastids: the Chloroplastida, Rhodophyta, and Glaucophyta (Fig. 2.5). From this starting point, photosynthesis spread widely among the eukaryotes via secondary endosymbiotic events that involved the capture of either green or red algae by diverse non-photosynthetic eukaryotes, thus transferring the once captured cyanobacterial endosymbionts (i.e., the plastids) laterally among eukaryotes. Some of these secondary endosymbiotic partnerships have in turn been captured by other eukaryotes, known as tertiary endosymbiosis, resulting in an intricate history of plastid acquisition (Keeling 2010; Fig. 5.47). Three groups of photosynthetic eukaryotes have plastids derived from a green algal endosymbiont: the chlorarachniophytes (see Sect. 5.3), a small group of mixotrophic algae from tropical seas, the euglenids (see Sect. 5.4), which are common in freshwater, and some green dinoflagellates (Lepidodinium spp., see Sect. 5.2). It is generally recognized that the green algae are ancient, but dating their origin is difficult because of the sparse fossil record (Fig. 4.29). The earliest fossils attributed to green algae include Proterocladus species from Neoproterozoic deposits (750 mya − 1bya) with filamentous thalli that resemble siphonocladous green algae (Butterfield et al. 1994; Tang et al. 2020). However, whether these ancient fossils represent green seaweeds remains uncertain (Del Cortona et al. 2020; Hou et al. 2022). For example, the resistant outer walls of prasinophyte cysts (phycomata) are well preserved in fossil deposits and abundant from the Palaeozoic era onward only (ca. 250–540 mya) (Colbath 1983). The oldest reliable records of siphonous seaweeds (Bryopsidales, Dasycladales) and stoneworts (Charophyceae) are from the

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Fig. 4.29 Green algal fossils. a Caulerpa-like fossil from the Permian (Gustavson and Delevoryas 1992); b Archaeobatophora typa, a dasycladalean green alga from the Upper Ordovician (image retrieved from Biodiversity Heritage Library) (from Nitecki 1976); c Proterocladus antiquus, a species interpreted as a benthic siphonocladalean green alga from the Meso- Neoproterozoic transition (from Tang et al. 2020)

Palaeozoic also (Hall and Delwiche 2007; Verbruggen et al. 2009). Molecular clock analyses based on these fossil records have estimated the origin of the green lineage between 700 and 1500 mya (Herron et al. 2009; De Clerck et al. 2012; Sánchez-Baracaldo et al. 2017; Del Cortona et al. 2020; Nie et al. 2020; Hou et al. 2022). Molecular phylogenetic evidence indicates that early in the evolution of Chloroplastida two principal lineages emerged: the Chlorophyta and Streptophyta, which have followed different evolutionary paths (Leliaert et al. 2012). A third species-poor lineage of marine planktonic and benthic green algae, the Prasinodermophyta, likely emerged before the origin of the Chlorophyta and Streptophyta (Li et al. 2020). The Chlorophyta, initially diversified as unicellular planktonic algae in oceans and freshwater environments, gave rise to the modern prasinophytes and the morphologically diverse core Chlorophyta which radiated in marine coastal, freshwater, and terrestrial environments. The Streptophyta evolved in freshwater and damp terrestrial habitats and colonized dry land approximately 476–432 million years ago, giving rise to the embryophytic land plants. The early evolutionary history of the Chlorophyta in the oceans and possibly freshwater environments of the Mesoand Neoproterozoic is marked by a radiation of planktonic unicellular green algae (prasinophytes), which played a vital role in the eukaryotic “greening” of our planet (Falkowski et al. 2004). The microfossil record indicates that these planktonic green algae dominated the eukaryotic phytoplankton of the Paleozoic. During the Neoproterozoic, the

prasinophytes gave rise to the morphologically and ecologically diverse core chlorophytes. The green seaweeds (Ulvophyceae) radiated in marine benthic habitats, possibly in the Meso-Neoproterozoic, and formed key components in coastal environments (Del Cortona et al. 2020; Hou et al. 2022). Two lineages, the Chlorophyceae and Trebouxiophyceae, radiated in freshwater and damp terrestrial habitats and came to dominate these environments during the Palaeozoic and Mesozoic Eras. In oceanic environments, dinoflagellates, coccolithophores (Haptista), and diatoms increased in abundance and largely displaced the green algae from the end-Permian onward. Similarly, the decline of green algae in freshwater phytoplankton started with the rise of freshwater dinoflagellates during the Cretaceous, and the appearance of diatoms and chrysophytes during the Cenozoic. Unicellular Streptophyta were likely the first eukaryotic algae in Neoproterozoic freshwater environments (Becker and Marin 2009). During the Palaeozoic, two multicellular charophyte lineages, the Zygnematophyceae and Charophyceae, diversified and dominated freshwater macrophytic communities between the Permian and Early Cretaceous. These macrophytes were largely replaced by freshwater angiosperms in the Late Cretaceous and Tertiary.

4.2.3 Defining Characters of the Green Algae The green algae possess several unifying traits that sets them apart from other groups of algae. As mentioned above, many

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of these characteristics are shared with embryophytic land plants. The most important defining characters, which will be elaborated on in the sections below, include: 1. A double membrane encloses the chloroplasts. Thylakoids are grouped in grana. Chloroplasts are not associated with an encircling layer of the endoplasmic reticulum. 2. The photosynthetic pigments chlorophyll a and b are present, along with accessory pigments such as carotenes and xanthophylls. 3. Pyrenoids, when present, are embedded within the chloroplast, penetrated by thylakoids, and (often) surrounded by starch. 4. Starch is the most important reserve polysaccharide, which can be clustered around the pyrenoids or scattered through the chloroplast stroma. 5. Flagellate cells typically possess two (or a multiple of two) flagella that are similar in structure (isokont), although they may differ in length and behavior. 6. The flagellar transition zone (i.e., the region between the flagellum and its basal body) is characterized by a star-like (stellate) structure, a nine-pointed star, visible in cross section using an electron microscope, linking nine pairs of microtubules. 7. Cell walls, when present, are generally composed of a fiber matrix of cellulose.

4.2.3.1 General Morphology Green algae exhibit a remarkable morphological diversity, with thallus architectures ranging from coccoid or flagellate unicellular algae to colonies and large giant-celled macrophytes. The different organizational levels are summarized in Fig. 4.30. 4.2.3.2 Chloroplasts All green algal cells contain at least one plastid (or the remnant of a plastid). Plastids generally contain photosynthetic pigments (chloroplasts), or they may serve as storage organelles (e.g., unpigmented starch-rich plastids in some siphonous green algae, resembling land plant amyloplasts). A double membrane surrounds plastids without enclosing periplastidal endoplasmic reticulum (Fig. 4.31). Chlorophyll a and b are located in the thylakoids, which are stacked and form typical grana. The photosynthetic reserve (starch) production and storage occurs inside the plastid. This is unique to the green algae (and, by extension, the Embryophyta); in other eukaryotic algae the photosynthetic storage product, whether starch or some other material, is found primarily in the cytoplasm. The light-harvesting complexes are categorized into LHCA and LHCB, which serve as the antennas for photosystem I (PSI) and photosystem II (PSII),

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respectively (in contrast, in red algae, Lhcr subunits function as the light-harvesting antenna for PSI) (Qin et al. 2019). The shape and number of chloroplasts per cell are very variable among green algae but are usually uniform within families or genera (Fig. 4.32). For this reason, green algal plastid shapes and numbers are widely used as taxonomic characters. Many flagellate cells of green algae contain an eyespot or stigma (Fig. 4.33), which appears as a red spot and is always located inside the chloroplast, never outside like in dinoflagellates and euglenoids (see Sects. 5.2 and 5.4). The eyespot consists of one to eight rows of closely packed carotenoid globules inserted between thylakoids. The photoreceptor is rhodopsin, located in the plasma membrane overlying the eyespot. The eyespot has several functions; it enhances the contrast, increases the intensity of the light, and shields the photoreceptor. The eyespot and photoreceptor function in light perception, controlling the direction of motility of flagellate cells (phototaxis). Depending on the light intensity, cells swim toward a dim light source (positive phototaxis) or away from bright light (negative phototaxis). In many green algae, the chloroplast contains one to several pyrenoids, which are the center of starch production. Pyrenoids appear as round or oval granular structures penetrated by several thylakoids or tube-like extensions of thylakoids and surrounded by several plates of starch, but without a definite boundary separating it from the rest of the chloroplast (Fig. 4.34). The formation of starch occurs within the chloroplast but is not restricted to the pyrenoids, as starch grains are also found elsewhere in the chloroplast (Fig. 4.31). The starch deposition is particularly extensive in algae growing in nutrient-poor conditions. Most green algae have a typical green color resulting from abundant chlorophylla and chlorophyllb that are not masked by large amounts of differently colored accessory pigments. However, a number of green algae occupying dry or sunny habitats may accumulate photoprotective orange or red pigments in amounts sufficient to obscure chlorophyll (Fig. 4.35).

4.2.3.3 Flagella Most green algae have flagellate cells at least in some stage of the life cycle (exceptions include species in the Zygnematophyceae, Prasinodermophyta, Chloropicophyceae, Picocystophyceae, some Mamiellophyceae, and some Trebouxiophyceae, where flagella are absent). Flagellate cells generally have two or a multiple of two flagella (up to 16 in Pyramimonas), which are generally inserted apically or subapically, or more rarely laterally (some prasinophytes and Mesostigma). The flagella of a cell are isokont, which means they are similar in structure, although they may differ in length or behavior. Few green algae have a single flagellum

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Fig. 4.30 Diversity of thallus organizations in the green algae Cladophora and Chara represent examples of coenocytic and of parenchymatous thalli, respectively

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Fig. 4.31 Ultrastructure of the flagellate cell of Chlamydomonas, showing the main intracellular structures, including the central nucleus with the nucleolus, surrounded by the cup-shaped chloroplast containing stacked thylakoid membranes, starch granules, and pyrenoid within the stroma. An eyespot is positioned against the inner envelope membrane of the chloroplast. Two flagella project from the apical region of the cell. Figure modified from Engel et al. (2015) licensed under CC BY 4.0, https://doi.org/10.7554/eLife. 04889.001

only (e.g., the prasinophytes Micromonas and Monomastix). A rare type of flagellate cells is characterized by a ring of flagella around the anterior end of the cell (stephanokont zoospores) and occurs in asexual reproductive cells of Oedogoniales (Chlorophyceae; Fig. 4.60) and Derbesia (Ulvophyceae). Such flagellate cells have presumably evolved independently in the two lineages. Flagella have a 9 + 2 arrangement of microtubules (axoneme), typical for flagella and cilia in other eukaryotes. Towards the base of the flagellum, just above where it emerges from the cell, the axoneme passes into a transition zone, which typically contains a structure that appears star-shaped (stellate) in cross-section, and H-shaped in longitudinal section, visible using an electron microscope (Fig. 4.36). The flagella terminate inside the cell by basal bodies, composed of nine interconnected triplets of microtubules, and function as a template for the assembly of the flagellar axoneme. Basal bodies are ultrastructurally similar to centrioles, which are present at each pole of the mitotic spindle during nuclear division. The significance of the centrioles, which replicate during cell division, is now thought to be that they transmit the capacity to form flagella from one cell generation to another (Carvalho-Santos et al. 2011).

In most species of Chlorophyta, the flagella are anchored in the cell by four microtubular roots, which are connected to the basal bodies and run beneath the plasma membrane to the posterior end of the cell (Fig. 4.37). When the cell is viewed from the top, these microtubular roots are symmetrically arranged in a cross (cruciate root system). Two of these roots contain two microtubules, and the two others contain three to eight microtubules (X-2-X-2 configuration). In cruciate root systems, the relative position of the basal bodies can be positioned directed oppositely (DO), shifted clockwise (CW), or shifted counter-clockwise (CCW). Basal body orientation is relatively conserved, and the different types of orientation characterize some of the main groups of green algae (van den Hoek et al. 1995; Graham et al. 2016). In the Streptophyta and some prasinophytes (e.g., Halosphaera and Pterosperma) the flagellar roots are arranged asymmetrically, with basal bodies positioned parallel to one another, and with a broad unilateral band of approximately 60 microtubules (multi-layered structure) anchoring the flagella in the cell (O’Kelly 1992; van den Hoek et al. 1995). Some green algae have a deviant flagellar root system Those include Nephroselmis species (Nephroselmidophyceae) that only have three microtubular roots with a multi-layered structure associated with one of them (Moestrup and Ettl

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Fig. 4.33 Eyespot in Chlamydomonas. a Transmission electron micrograph of the eyespot apparatus of Chlamydomonas, composed of carotenoid globules (asterisks) inserted between thylakoids (arrowheads) and the chloroplast envelope (large arrows). The plasma membrane (small arrow) is closely attached to the chloroplast envelope in the region overlying the eyespot. Scale bar = 300 nm (from Schmidt et al. 2006); b Schematic drawing of the eyespot apparatus, based on Ueki et al. (2016)

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Fig. 4.34 Chloroplasts with pyrenoids. a Light micrograph of plastids forming a reticulate network with numerous pyrenoids in Boodlea; b Transmission electron micrograph of a pyrenoid within the chloroplast (CH) with starch plates (s) and protruding thylakoids (t). The nucleus (n) is also indicated. Figure b from Miyaji (1999)

Fig. 4.35 Examples of green algae occupying dry or sunny habitats and accumulating photoprotective orange or red pigments which obscure the green chlorophyll pigments. a Purple-red films of Haematococcus in bird baths, courtesy of James Morgan; b orange-colored Trentepohlia growing in terrestrial habitats; c and red growths of Chlamydomonas nivalis in mountain snows

Fig. 4.36 Flagellar apparatus of Chlamydomonas. a Transmission electron micrographs of longitudinal section, from Dutcher and O’Toole (2016), licensed under CC BY 4.0, https://doi.org/10.1186/s13630-016-0039-z; b stellate structure of the transition zone in cross-section, from Lechtreck et al. (2013), licensed under CC BY 4.0, https://doi.org/10.1186/2046-2530-2-15

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Fig. 4.37 Schematic representation of the flagellar root system. A Female gamete of Bryopsis maxima, with flagella and microtubular roots observed using anti-a tubulin immunofluorescence, from Miyamura et al. (2010); B Cell viewed from the side with two flagella (red) terminating in basal bodies (blue), anchored by four microtubular roots (purple) connected to the basal bodies. Two roots are composed of two microtubules (r-2), the two other by three to eight microtubules (r-X); C Cell viewed from the top showing the four microtubular roots arranged symmetrically in a cross (cruciate root system); D Four main types of basal body orientations. DO, cruciate root system with directly opposed basal bodies. CW, cruciate root system with clockwise displaced basal bodies. CCW, cruciate root system with counter-clockwise displaced basal bodies. Asymmetrical, parallel basal bodies and multi-layered structure (MLS)

1979). Species in the Trentepohliales (Ulvophyceae) and some species in the Trebouxiophyceae that have four cruciately arranged microtubular roots along with a multi-layered structure (Melkonian and Peveling 1988; López-Bautista et al. 2002). Although it remains uncertain whether the multi-layered structure in green algae with asymmetrical and cruciate root systems are homologs, these multi-layered structures may present an ancestral feature of flagellate green algae, which disappeared in various lineages (van den Hoek et al. 1995). Other flagellar structures found in green algae include two transversely striated robust fibers, which connect the flagella, and a contractile striated fiber (rhizoplasts) that connects the flagellar apparatus to the nucleus (e.g., Chlamydomonas) or the chloroplast (e.g., Pyramimonas). Transversely striated connective fibers are present in most green algae. Rhizoplasts are common in the Chlorophyta, but rare in the charophytes.

4.2.3.4 Mitosis and Cytokinesis The processes of mitosis and cell division (cytokinesis) are variable among the main green algal groups (van den Hoek et al. 1995). During early prophase, the basal bodies duplicate and move towards the poles of the future mitotic spindle, where they function as centrioles. An unusual type of mitosis is present in species of Trebouxiophyceae, where the centrioles are not positioned at the poles of the mitotic spindle but at the metaphase plate of the chromosomes (metacentric mitosis). Two main types of mitosis can be distinguished. Closed mitosis occurs in most Chlorophyta, and is characterized by a nuclear envelope that remains

nearly intact during metaphase (although at the poles, there are polar perforations, openings in the nuclear membrane through which the microtubules of the mitotic spindle penetrate in the nucleus). Open mitosis occurs in most charophyte green algae and features a nuclear envelope broken down during nuclear division. The behavior of the mitotic spindle during telophase also varies from group to group. In most green algae, the mitotic spindle degenerates quickly during telophase (non-persistent telophase spindle). In some groups, including Ulvophyceae, the mitotic spindle persists, resulting in a typical dumbbell-shaped telophase configuration. Mitosis is followed by cytokinesis in most green algae, with the notable exception of several green seaweeds with multinucleate cells (e.g., Cladophorales and Bryopsidales), where mitosis is uncoupled from cytokinesis. In most green algae, cell division is accomplished by furrowing, in which a cleavage furrow is usually already present at the early prophase, and develops further during telophase (Fig. 4.38a). Green algae with giant, multinuclear cells, including the coenocytic cells of Cladophorales and siphonous cells of Bryopsidales and Dasycladales, exhibit specialized modes of cell division. In several groups of the core Chlorophyta, cell division is mediated by a phycoplast, which is an array of microtubules oriented parallel to the plane of cell division, determining the formation of a new cell wall, which is either formed through cell wall ingrowth or outgrowth (Fig. 4.38b). In the charophyte classes Charophyceae, Zygnematophyceae, and Coleochaetophyceae, cell division is mediated by a phragmoplast, in which an array of microtubules is oriented perpendicularly to

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Fig. 4.38 Main types of cell division in the green algae. a microtubule-independent cleavage furrow; b phycoplast microtubules oriented perpendicular to the spindle axis are involved in cell wall formation through cell wall ingrowth; c phragmoplast microtubules are oriented parallel to the spindle axis associated with centrifugal vesicle fusion leading to the development of a cell plate with plasmodesmata. Modified from Graham et al. (2016)

the plane of cell division, determining the formation of the cell plate and new cell wall (Fig. 4.38c). This type of cell division usually results in the formation of plasmodesmata, which are channels through the cross walls that enable intercellular communication, and the development of complex tissues. In Zygnematales, an intermediate situation occurs whereby furrowing of the plasma membrane is accompanied by phragmoplast formation (Buschmann and Zachgo 2016), and no plasmodesmata are formed.

4.2.3.5 Cell Walls The cells of most green algae, including many unicellular and all multicellular or siphonous species, are surrounded by a cell wall (Domozych et al. 2012; Domozych 2019). Several unicellular green algae, however, either lack a cell wall (“naked” cells) or have other types of cell covering, including organic body scales (van den Hoek et al. 1995; Becker et al. 1994). Most green algal cell walls have a polysaccharide fibrils-matrix organization, while some species have cell walls composed mainly of glycoproteins (Domozych 2019). In Chlorophyta, the cell wall is often composed of cellulose or chitin-like polysaccharides, xylan, or mannan. The cell walls in prasinophytes are highly diverse. Many prasinophytes are naked (e.g., Ostreococcus, Micromonas). In contrast, others (e.g., Pyramimonas, Nephroselmis) are covered by one to several layers of organic body scales of various forms, including plate-like, hair-like, and complex, three-dimensional structures (Melkonian 1990). These organic structures are mainly composed of monosaccharides and are produced within the Golgi apparatus. Cells in the Chlorodendrophyceae are covered by a coherent wall composed of fused body scales. The Chlorophyceae include a wide diversity of cell walls ranging from cellulose-pectin complexes to walls composed

of hydroxyproline-rich glycoproteins. Cell wall composition may remain identical or change during the life cycle. Cell walls in the Ulvophyceae are also highly variable and are composed of cellulose, mannans, glucan, xylans, and sulfated and/or pyruvylated polysaccharides (Ciancia et al. 2020). In the charophyte classes Charophyceae, Zygnematophyceae, and Coleochaetophyceae, cell walls contain assemblages of polymers similar to land plants, including cellulose, pectins, hemicelluloses, arabinogalactan proteins, extensin, and lignin. Some members of the ulvophycean orders Bryopsidales (e.g., Halimeda) and Dasycladales (e.g., Neomeris) and some species of Charophyceae (e.g., Chara) are calcified by depositing calcium carbonate as aragonite crystals on the outside of the cells, forming a structural defense against grazing.

4.2.3.6 Giant Multinucleate Cells The thalli in some green seaweed orders and the Charales are characterized by large multinucleate cells. Species in the orders Cladophorales and Blastophysa and some species in the Ulotrichales (e.g., Urospora and Acrosiphonia) have a siphonocladous thallus organization, which is characterized by multicellular thalli composed of multinucleate cells with nuclei organized in regularly spaced cytoplasmic domains (Motomura 1996; McNaughton and Goff 1990). Species in the orders Bryopsidales and Dasycladales have a siphonous thallus organization, characterized by thalli consisting of a single giant tubular cell. Local constrictions (e.g., side branches in Bryopsis) or rings of cell wall material (e.g., Codium) may subdivide these tubular cells into compartments. Still, essentially the cytoplasm is continuous over the entire thallus. In most species, the siphonous cells contain thousands of nuclei. Still, in several species of Dasycladales, the siphonous thallus remains uninucleated throughout much

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of its life cycle with a giant diploid nucleus that only divides at the onset of reproduction (Berger and Kaever 1992). Siphonous cells typically exhibit cytoplasmic streaming, transporting organelles, nutrients, and transcripts across the thallus (Menzel 1994, 1987; Mine et al. 2005). Some siphonous species form large seaweeds with thalli functionally differentiated into distinct structures, including rhizoids, stolons, and blades. The giant cells of siphonocladous and siphonous species are characterized by several cytological specializations, such as unique mechanisms of cell differentiation, cell division, and wounding response (Menzel 1988; La Claire 1992; Kim et al. 2001; Mine et al. 2008). At the opposite end of the green tree of life, the internodal cells of the Charales, which measure up to 1 mm in diameter and up to 10 cm in length, also display vigorous cytoplasmic streaming with flow speeds up to 50–100 lm/s. The particular pattern of streaming observed in Chara, which involves rotational streaming in two oppositely directed longitudinal stripes, is driven by myosin-coated vesicles that slide along actin cables (Woodhouse and Goldstein 2013).

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Fig. 4.39 Two types of haplontic life cycles in which mitotic divisions are limited to the haploid life stage, and haploid vegetative cells produce haploid gametes that fuse and form a zygote (the only diploid phase in the life cycle) followed by a meiotic division. A Life cycle of Chlamydomonas (Chlorophyceae) in which two isogametes (+ and −) attach to their anterior ends and fuse to form a zygote, followed by fusion of the gametic nuclei to form a diploid zygotic nucleus. The diploid zygote becomes thick-walled and dormant (hypnozygote). Meiosis takes place in the diploid zygote, followed by the formation of four haploid zoids, which are released and develop into vegetative cells. Vegetative cells divide mitotically (not shown in figure), or form gametes. B Haplontic life cycle of Ulothrix (Ulvophyceae) in which vegetative, haploid, different sexualized filaments (+ and − gametophytes) produce biflagellate isogametes (+ and −) which come together and attach to form a quadriflagellate planozygote, which attaches to the substratum and germinates into a stalked cell (Codiolum stage). Under the right conditions, the content of the stalked zygote divides up to give 4–16 quadriflagellate meiospores, which germinate and grow into haploid vegetative filaments, which grow by mitosis and cell division. Vegetative filaments also produce asexually through quadriflagellate zoospores, which grow directly into new vegetative filaments (not shown in figure). Based on Lokhorst and Vroman (1972) and van den Hoek et al. (1995)

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Fig. 4.40 Two types of diplohaplontic life cycles in which there are two free-living vegetative phases that undergo mitotic divisions. A Isomorphic diplohaplontic life cycle of Ulva (Ulvophyceae) with alternation of two morphologically similar multicellular generations: a haploid gametophyte and a diploid sporophyte. Biflagellate female and male gametes (female gametes being slightly larger than the male ones) are formed by mitotic cell divisions in the female and male gametophytes, respectively. After anisogamous fusion, diploid zygote germinates and develops into a diploid sporophyte. Cells in the sporophyte undergo meiosis and produce quadriflagellate, haploid meiospores, which germinate and develop into female or male gametophytes, based on (Koeman and Van den Hoek 1981); B Heteromorphic diplohaplontic life cycle of Derbesia (Ulvophyceae) with alternation of two morphologically dissimilar generations. Biflagellate female and male gametes are formed in the female and male gametophytes, and are released through pores of the vesicular thalli. After anisogamous fusion (plasmogamy) a binucleate cell is formed, containing one male and one female nucleus, which grows in a siphonous sporophyte with numerous male and female nuclei. The sporophyte develops pear-shaped sporangia, in which karyogamy between male and female nuclei takes place, followed by meiosis and the development of stephanokont meiospores. Meiospores contain several nuclei and develop into female or male gametophytes, based on Eckhardt et al. (1986) and van den Hoek et al. (1995)

environment (marine versus freshwater). The marine Ulvophyceae mainly have diplohaplontic life cycles, while most freshwater green algae have a haploid vegetative phase and a single-celled, often dormant zygote as the diploid stage. The haplontic life cycle associated with a dormant zygote which characterizes many freshwater green algae is often viewed as an adaptation toward adverse ecological conditions such as desiccation of shallow pools and ponds (Cavalier-Smith 2002). The encysted zygote is well-adapted to desiccation and its pigmentation reduces radiation damage by UV. Furthermore, the diploid nature of the cell would enable recombinational repair of double-strand breaks of the DNA, a repair mechanism that is absent in haploid cells. In several green algal groups (mostly prasinophytes, Mesostigmatophyceae, Klebsormidiophyceae, Prasinodermophyta, and several terrestrial members of the core Chlorophyta), sexual reproduction appears to be absent or at least goes undocumented. Information from genome sequences however usually does reveal a full complement of genes involved in meiosis (Grimsley et al. 2010; Fučíková et al. 2015; Speijer et al. 2015), indicating that sexual reproduction is probably cryptic rather than absent. Asexual reproduction occurs by binary fission, fragmentation, or production of flagellate cells (zoospores), or non-motile autospores (Fig. 4.41).

4.2.5 Systematics and Classification of the Chloroplastida Early classification schemes of green algae and hypotheses on their evolutionary history were based on the classical notion that evolution follows trends in levels of complexity. In this view, it was thought that ancestral unicellular green algae have given rise to distinct lines of increasing size and morphological complexity. However, data on the ultrastructure of green algal cells, gathered by electron microscopy from the 1970s onwards, led to a reconsideration of green algal classification. The fine structure of flagella and variation in the processes of mitosis and cell division (see Sects. 4.2.3.3 and 4.2.3.4) were believed to more accurately reflect evolutionary relationships because of their involvement in fundamental and conserved processes of cell motility and replication, and are thus less liable to convergent evolution than gross morphological traits, such as thallus architecture. Phylogenetic hypotheses based on ultrastructural data postulated an early diversification of unicellular flagellates, resulting in several lineages of flagellate green algae, some of which then evolved into more complex colonial, multicellular, or siphonous forms (van den Hoek et al. 1995). From the 1990s onward, DNA sequence data has provided a new framework for revising the relationships among green

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Fig. 4.41 Asexual reproduction in coccoid green algae by the formation of autospores and zoospores (schematic example). a Coccoid vegetative cell with prominent cell wall, chloroplast appressed to the cell wall with a pyrenoid, nucleus with nucleolus at interphase; b Two stages of division of protoplast by mitosis within the parental cell wall, a pulsating vacuole may have appeared, the protoplasts are naked, i.e., without own cell wall; c Autosporangium with numerous autospores. After multiple successive divisions of the protoplast numerous daughter cells are formed which become encircled by their own walls within the parental cell wall, as the sporangial (parental) wall disintegrates it ruptures and passively releases the autospores (daughter cells); d Autospores (enlarged for the graphic), they look like small copies of the vegetative cell, in some cases the shape of the autospores may be different from that of the vegetative cell; e Zoosporangium, multiple divisions of the protoplast result in numerous daughter cells without a cell wall, the daughter cells start to develop emergent flagella and other characteristics of motile cells, e.g., pulsating vacuoles; numerous motile zoospores with emergent flagella are released as the sporangial (parental) wall disintegrates. f Zoospore with two equally long emergent flagella and a red eyespot at the periphery of the chloroplast (enlarged for the graphic), the zoospore settles following ceasing the flagella after a period of active swimming and being positively phototactic, it starts to develop a cell wall thus representing a young coccoid vegetative cell which enlarges

algae. In general, molecular phylogenetic analyses have validated higher-level classification of the green algae based on ultrastructural data. Initially analyses were based on individual genes, including mainly ribosomal rDNA and to a lesser extent plastid genes. More recently, studies unraveling deep relationships in the green algae now make use of multigene sequence data (Lemieux et al. 2014a, b; Leliaert et al. 2016; Lopes dos Santos et al. 2017; Cheng et al. 2019; One Thousand Plant Transcriptomes Initiative 2019; Del Cortona et al. 2020; Li et al. 2020; Hou et al. 2022). Current hypotheses on

green algal evolution posit the early divergence of tree lineages: the Chlorophyta, including the majority of described green algal species, the Streptophyta comprising of the charophytes, a paraphyletic assemblage of freshwater green algae, and the land plants, and a small clade including a few marine species, the Prasinodermophyta (Fig. 4.28). Below is an overview about the current classification and the systematic arrangement of the Chloroplastida (Table 4.2). It is adapted from Leliaert et al. (2012), Leliaert et al. (2015), Neustupa (2015), Lopes dos Santos et al.

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Table 4.2 Overview of the current classification of the Chloroplastida Phylum

Class

Order

Families

Prasinodermophyta

Prasinodermophyceae

Prasinodermales

Prasinodermaceae

Palmophyllophyceae Chlorophyta

Prasinococcaceae Palmophyllaceae

Nephroselmidophyceae

Nephroselmidales

Nephroselmidaceae

Mamiellophyceae

Mamiellales

Bathycoccaceae, Mamiellaceae

Dolichomastigales

Dolichomastigaceae, Crustomastigaceae

Monomastigales

Monomastigaceae

Pyramimonadophyceae

Pyramimonadales

Pyramimonadaceae

Chloropicophyceae

Chloropicales

Chloropicaceae

Picocystophyceae

Picocystales

Picocystaceae

Pseudoscourfieldiales

Pycnococcaceae

Chlorodendrophyceae

Chlorodendrales

Chlorodendraceae

Pedinophyceae

Marsupiomonadales

Marsupiomonadaceae, Resultomonadaceae

Pedinomonadales

Pedinomonadaceae

Trebouxiophyceae

Streptophyta

Prasinococcales Palmophyllales

See the less formal clade system of the Trebouxiophyceae, Table 4.1 and Fig. 4.52

Chlorophyceae

See the less formal clade system of the Chlorophyceae, Table 4.2 and Fig. 4.53

Ulvophyceae

Bryopsidales

Bryopsidaceae, Caulerpaceae, Codiaceae, Derbesiaceae, Dichotomosiphonaceae, Halimedaceae, Ostreobiaceae, Pseudobryopsidaceae

Ulvales

Bolbocoleonaceae, Cloniophoraceae, Ctenocladiaceae, Kornmanniaceae, Phaeophilaceae, Ulvaceae, Ulvellaceae

Ulotrichales

Acrosiphoniaceae, Binucleariaceae, Gomontiaceae, Hazeniaceae, Helicodictyaceae, Kraftionemaceae, Monostromataceae, Planophilaceae, Sarcinofilaceae, Tupiellaceae, Ulotrichaceae

Chlorocystidales

Chlorocystidaceae

Oltmannsiellopsidales

Oltmannsiellopsidaceae

Scotinosphaerales

Scotinosphaeraceae

Ignatiales

Ignatiaceae

Cladophorales

Anadyomenaceae, Cladophoraceae, Okellyaceae, Pithophoraceae, Pseudocladophoraceae, Siphonocladaceae

Dasycladales

Dasycladaceae, Polyphysaceae

Trentepohliales

Trentepohliaceae

The classes and orders discussed in the Streptophyta do not form a monophylum and, therefore, are referred to the “streptophyte algae grade” (Fig. 4.69). Due to ongoing molecular phylogenetic and phylogenomic works, no family-level classification appears to be appropriate yet for the streptophyte algae grade. Mesostigmatophyceae1

Mesostigmatales Chlorokybales2

Klebsormidiophyceae

Klebsormidiales

Charophyceae

Charales

Coleochaetophyceae

Chaetosphaeridiales Coleochaetales

Zygnematophyceae3

Desmidiales Zygnematales

1

The Mesostigmatophyceae also encompass Spirotaenia, which may represent an additional order or even a distinct class Recently, transcriptomic analyses revealed deep phylogenetic divergences with the genus Chlorokybus, and this substantiated the Chlorokybales to represent an independent class Chlorokybophyceae (Irisarri et al. 2021) 3 Recently, transcriptomic analyses revealed five independent origins of the filamentous growth within the Zygnematophyceae, which led to the proposal of a new five-order system for the Zygnematophyceae (Hess et al. 2022) 2

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(2017), Caisová and Melkonian (2018), Škaloud et al. (2018), and Li et al. 2020. The list contains extant taxa only.

4.2.6 Phylum Prasinodermophyta The phylum Prasinodermophyta includes marine planktonic coccoid green algae, as well as benthic macroscopic thalli composed of distantly spaced cells embedded in a gelatinous matrix (Li et al. 2020). Cell division is unequal, and flagellar stages have never been observed. Only asexually reproducing species are known so far. The class includes a few species summarized in two classes: the Prasinodermophyceae and Palmophyllophyceae. The class Prasinodermophyceae is represented by the single genus Prasinoderma (Prasinodermales), including a few picoplanktonic coccoid species with cells surrounded by

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a thick multi-layered cell wall (Fig. 4.42a) (Hasegawa et al. 1996; Jouenne et al. 2011). The class Palmophyllophyceae (Leliaert et al. 2016) includes two orders. The Prasinococcales with the single genus Prasinococcus, representing marine planktonic species characterized by coccoid cells with a thin cell wall perforated by pores, and surrounded by a thick gelatinous capsule (Fig. 4.42b) (Miyashita et al. 1993). The Palmophyllales include benthic species growing in dimly lit marine habitats (Zechman et al. 2010). Palmophyllales have a unique type of multicellularity, forming well-defined macroscopic thalli composed of coccoid cells embedded in a firm gelatinous matrix (Fig. 4.42c–e). Species in the genus Verdigellas have been recorded from depths down to 200 m (Ballantine and Norris 1994). These low-light ecosystems present a challenging environment for photosynthetic organisms and relatively few algae live in such habitats (Leliaert et al. 2011).

Fig. 4.42 Morphological diversity in the Prasinodermophyta. a Prasinoderma coloniale (Prasinodermales), a picoplanktonic unicellular marine green alga, from Li et al. (2020). b Prasinococcus capsulatus (Prasinococcales), non-motile cells, each embedded in gelatinous capsules, courtesy of Daniel Vaulot, Station Biologique de Roscoff. c Palmophyllum crassum (Palmophyllales) from deep water, forming irregularly lobed crusts, tightly fixed to the substrate, courtesy of Véronique Lamare. d Cross section of Palmophyllum, showing the coccoid cells embedded in a gelatinous matrix. e Palmoclathrus stipitatus (Palmophyllales) from deep-water habitats of Southern Australia, characterized by perennial stalks from which seasonal, net-like blades grow, courtesy of Kevin Branden, Botanic Gardens & State Herbarium, Adelaide

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4.2.7 Phylum Chlorophyta and Prasinophytes The Chlorophyta contain most of the green algal species, occurring in diverse marine, freshwater, and terrestrial environments. In aquatic habitats, species may be planktonic or benthic. The Chlorophyta display a large morphological diversity, including unicellular, colonial, multicellular, and siphonous forms. Mitosis is generally closed with a nuclear membrane persisting throughout the metaphase stage. The flagellar apparatus is typically characterized by a symmetrical cruciate root system wherein rootlets of variable (X) numbers of microtubules alternate with rootlets composed of two microtubules to form a “X-2-X-2” arrangement. The orientation of the flagellar root system (Fig. 4.37d) has served as an important character for defining the main groups of Chlorophyta. At present 10 classes are recognized within the Chlorophyta. Continuous taxonomic effort has resulted in the formal description of several lineages at class-level, especially within the former Prasinophyceae. Prasinophytes. The early diverging lineages of the Chlorophyta form a paraphyletic group, collectively termed prasinophytes (Fig. 4.28). They are a diverse group of planktonic unicellular green algae in predominantly marine, but also freshwater environments. Cell shapes are highly variable. Cells can be naked, or covered by cell walls or organic body scales. The fine structure of these scales is equally diverse, and has traditionally been used to differentiate the major groups of prasinophytes (Melkonian 1990; Leliaert et al. 2011; Sym and Pienaar 1993; Sym 2015). Some species possess up to seven distinct scale types, others have a single type. Mitotic processes vary between groups, and flagella are present or absent. The number of flagella and their behavior is also very variable. Some cells push with

undulating flagella, while others swim with flagella forward. Most species have four flagella, some species have one or eight, and some other eight or 16. Biochemical features, such as photosynthetic pigments and photorespiratory enzymes, are also diverse. Nearly all species are known to reproduce asexually (Nephroselmis species being the exception), although sexual reproduction has been indicated indirectly in Ostreococcus and Micromonas based on the presence of sex-related and meiosis-specific genes in their genomes (Derelle et al. 2006; Worden et al. 2009). Most prasinophyte lineages are relatively species-poor compared to the core Chlorophyta. Molecular phylogenetic data have identified several lineages of prasinophytes, many of which are currently recognized at the class level. Some of these lineages are only known from environmental sequences, and have not been formally described (Lepère et al. 2009; Lopes dos Santos et al. 2016). Below, the main prasinophyte lineages are described.

4.2.7.1 Class Mamiellophyceae The Mamiellophyceae is the morphologically and ecologically most diverse clade of prasinophytes, including approximately 20 species, occuring as phytoplankton in marine and freshwater habitats (Tragin and Vaulot 2019; Marin and Melkonian 2010). Species may have scaly or naked (i.e., lacking a cell wall) cells that are non-motile (coccoid) or motile with one or two flagella. The class includes the order Mamiellales and two smaller orders, Monomastigales (genus Monomastix) and Dolichomastigales (genera Crustomastix and Dolichomastix). The Mamiellales mainly include marine coccoid and flagellate species. Ostreococcus (Fig. 4.43) and Micromonas, which are among the smallest eukaryotes known (cell sizes of 0.5–2 µm), and form important components of marine picoeukaryotic

250 nm Fig. 4.43 Ostreococcus tauri (Mamiellophyceae, Mamiellales), a picoeukaryotic coccoid marine alga. Reconstruction from EM tomography after cryo-fixation (Henderson et al. 2007), nuclei (n, red), nuclear envelope (ne), chloroplasts (c, green), mitochondria (m, dark purple), Golgi bodies (g, yellow), peroxisomes (p, orange), granules (gr, dark blue), inner membranes including ER (er, light blue), microtubules (light purple), and ribosome-like particles (r). From Lechtreck et al. (2013), licensed under CC BY 4.0, https://doi.org/10.1186/2046-2530-2-15

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communities (Courties et al. 1994; Demir-Hilton et al. 2011). Sexual reproduction is unknown in the class, but indirect evidence for sexual reproduction in Ostreococcus and Micromonas is based on the occurrence of sex-related and meiosis-specific genes in their genomes (Derelle et al. 2006; Worden et al. 2009; Grimsley et al. 2010).

4.2.7.2 Class Pyramimonadophyceae The Pyramimonadophyceae includes species with relatively large flagellate cells that are covered by complex body scales occurring in multiple layers, and with 4, 8, or even 16 flagella (Fig. 4.44c–f). More than 50 species are described from marine and freshwater habitats (Daugbjerg et al. 2020). Some species of Pyramimonas and Cymbomonas are mixotrophic and possess a food uptake apparatus. Other species produce resting cysts surrounded by two wall layers (phycomata) (Fig. 4.44a, b) (Parke et al. 1978), which possibly result from sexual reproduction. These phycomata are abundant in the fossil record from the Cambrian onward (Kustatscher et al. 2014). 4.2.7.3 Class Nephroselmidophyceae The Nephroselmidophyceae include relatively large, asymmetrical flagellates with bean-shaped to ovoid flattened cells and two laterally inserted, unequal and heterodynamic flagella (Fig. 4.44h) (Nakayama et al. 2007; Yamaguchi et al. 2011). The flagellar apparatus is atypical by having three flagellar roots only. Cells and flagella are covered by diverse scales in multiple layers (Fig. 4.44h, i–k). Roughly some 30 species are known from marine and freshwater habitats. Nephroselmis is one of the few prasinophytes where sexual reproduction has been observed in culture. Marine Nephroselmis species have been identified as endosymbionts in the katablepharid Hatena arenicola (Okamoto and Inouye 2006; Yamaguchi et al. 2014). Endosymbiontic Nephroselmis cells show extensive structural changes and are ultrastructurally tightly associated with its host, but the integration of the symbionts is not entirely stable as the division of the symbiont is not coordinated with that of the host with some daughter cells having to capture new Nephroselmis symbionts from the environment. 4.2.7.4 Class Chloropicophyceae The Chloropicophyceae comprises scale-less coccoid picoplanktonic algae surrounded by a thin cell wall. The class includes 6 known species, several of which are important components of marine phytoplankton, especially in moderately oligotrophic waters (Tragin and Vaulot 2018; Lopes dos Santos et al. 2017). 4.2.7.5 Class Picocystophyceae Other prasinophyte clades, that are phylogenetically distinct from the aforementioned classes include the

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Picocystophyceae, containing the coccoid species Picocystis salinarum from inland saline lakes and ponds (Lewin et al. 2000; Lopes dos Santos et al. 2017), the Pycnococcaceae, including marine flagellates (Pseudoscourfieldia) and coccoids (Pycnococcus), and a number of clades that are known only from environmental sequencing.

4.2.8 The Core Chlorophyta, Chlorodendrophyceae, and Pedinophyceae The core Chlorophyta include three diverse and species-rich classes, Trebouxiophyceae, Chlorophyceae, and Ulvophyceae, and two smaller classes, Pedinophyceae and Chlorodendrophyceae. The core Chlorophyta are characterized by a mode of cell division that is mediated by a phycoplast which is a system of microtubules that develops parallel to the plane of nuclear division (a phycoplast is absent in the Pedinophyceae and Ulvophyceae). The orientation of the flagellar basal bodies is generally conserved within these classes. The Pedinophyceae, Chlorodendrophyceae, Trebouxiophyceae, and Ulvophyceae have counter-clockwise (CCW) displaced basal bodies. The Chlorophyceae has clockwise (CW) displaced basal bodies as well as directly opposite basal bodies (Fig. 4.37).

4.2.8.1 Class Chlorodendrophyceae The Chlorodendrophyceae is a small clade uniting the marine or freshwater scaly quadriflagellates Tetraselmis (Fig. 4.45) and Scherffelia (Guillou et al. 2004). These unicellular algae were traditionally regarded as members of the Prasinophyceae but they share several features with the core Chlorophyta clades, including closed mitosis and a phycoplast (Mattox and Stewart 1984). The close relationship with the classes of the core Chlorophyta has been confirmed by phylogenetic data (Del Cortona et al. 2020). 4.2.8.2 Class Pedinophyceae The Pedinophyceae include marine and freshwater uniflagellate algae. Cell division is not mediated by a phycoplast. Molecular phylogenetic data have provided evidence for an independent class Pedinophyceae, but the exact relationship with the phycoplast-containing core chlorophyte clades (Chlorodendrophyceae, Trebouxiophyceae, Ulvophyceae, and Chlorophyceae) remains uncertain (Del Cortona et al. 2020).

4.2.9 Class Trebouxiophyceae The class Trebouxiophyceae encompasses predominantly unicellular green algae of coccoid organization. Most of the so far known species have a clear preference for terrestrial

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Fig. 4.44 Diversity of Pyramimonadophyceae and Nephroselmidophyceae. a Pterospermella sp., a microfossil from the Lower Devonian consisting of a vesicle with one or more wings, interpreted as phycomata of prasinophycean green algae, similar to phycoma stages of the extant Pterosperma (Pyramimonadales), from Kustatscher et al. (2014); b Phycoma stage of Pterosperma, courtesy of Johannes Rick, Plankton*Net, Alfred Wegener Institute; c Flagellate cell of Pterosperma sp. characterized by four very long flagella, from PhycoKey (Baker et al. 2012); d Pyramimonas sp. (Pyramimonadales), flagellate cell with four similar flagella emerging from an anterior depression, with a large cup-shaped chloroplast and stigma, courtesy of David Patterson and Bob Andersen, Provasoli-Guillard National Center for Culture of Marine Phytoplankton); e Pyramimonas sp., scanning electron micrograph showing body and flagella covered with different types of organic body scales, courtesy of Rick van den Enden, Australian Antarctic Division; f Diversity of body scales in two species, Pyramimonas parkae (top) and P. amylifera (bottom), including intermediate cell body scales (left), outer cell body scales (middle), and outer flagellar scales (right), based on McFadden et al. (1986); g Nephroselmis sp. (Nephroselmidophyceae), flagellate cell with two laterally inserted unequal flagella and a cup-shaped chloroplast, courtesy of William Bourland; h Nephroselmis sp., scanning electron micrograph showing a complex covering of organic body scales, courtesy of Shoichiro Suda, University of the Ryukyus; i, j Nephroselmis, multiple layers of various organic body scales on the cell surface; k Nephroselmis, flagellum covered by scales and hairs, from Yamaguchi et al. (2011)

150 Fig. 4.45 Tetraselmis (Chlorodendrophyceae). a Tetraselmis indica from salt pans in India, characterized by flagellate cell with four equal flagella; b SEM image showing the flagella covered by hairs and scales, from Arora et al. (2013)

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habitats. However, a surprising diversity of lifestyles not found in other green algae is present in the Trebouxiophyceae. Members of the class are frequently involved in symbioses, i.e., exosymbiosis such as lichens (Sect. 6.2), endosymbiosis with ciliates and some metazoa, thrive as endophytes within plant tissue or are obligate heterotrophs with a parasitic lifestyle. In most genera of Trebouxiophyceae, the vegetative cells can withstand periods of drought without forming special dormant stages. It follows that sexual reproduction, during which a zygote as a resting stage would be developed, is of less importance, i.e., it has been observed in just very few trebouxiophytes. Likely, the class is among the most species-rich lineages of the Chlorophyta. Since the class was first described based on molecular phylogenetic analyses about 30 years ago, comprising just a few genera, its known diversity has increased fast up to now. The discovery of new species and lineages of the Trebouxiophyceae is still going on. Several members of Trebouxiophyceae are of economic importance or serve as model organisms, e.g., Botryococcus (Fig. 4.51) and Chlorella (Fig. 4.48a). Vegetative morphology There is only a narrow diversity of vegetative morphology. It may typically be referable to three different growth forms. Most trebouxiophytes are unicellular coccoid cells, e.g., Chlorella (Fig. 4.48a), Coccomyxa (Fig. 4.47h, i), Chloroidium (Fig. 4.47c, d), and Trebouxia (Fig. 4.46a, b). Many unicellular coccoids are rather minute, with vegetative cells smaller than 5 lm, e.g., Choricystis (Fig. 4.48h), Diplosphaera (Fig. 4.49a), and Muriella (Fig. 4.48b). The cells of Eremosphaera, however, are unusually large, with a diameter of up to about 200 lm (Fig. 4.49i). The shape of the trebouxiophyte unicells is mostly globose, e.g., Lobosphaera (Fig. 4.46e) and Elliptochloris (Fig. 4.47g). Other unicells are more ovoid, e.g., Chloroidium (Fig. 4.47d),

Coccomya (Fig. 4.47h, i); Symbiochloris (Fig. 4.48i), and Watanabea (Fig. 4.47a, b). The cells of Stichococcus are elongated and cylindrical (rod-shaped, Fig. 4.49b). Most trebouxiophyte cell walls are smooth. Some genera produce mucilage (extracellular polymeric substances, EPS), likely to support desiccation tolerance, e.g., in Coenochloris (Fig. 4.48f, g). Trochisciopsis is known for its sculptured cell wall (Fig. 4.46f). The sarcinoid growth (cell packets) is formed when daughter cells are kept together by remnants of the parental cell wall for some while following autospore formation (Fig. 4.41). Typical examples are found in Apatococcus (Fig. 4.46i), Desmococcus (Fig. 4.49e), and Pleurastrosarcina (Fig. 4.49h). The three genera are distributed in four independent lineages of the Trebouxiophyceae in molecular phylogenetic analyses (Fig. 4.52). Filaments are formed in Prasiola (Fig. 4.49c), Geminella (Fig. 4.49f), and Microthamnion. Stichococcus can also form short filaments, but they disintegrate rapidly. Prasiola can even form a thallus of blade-like shape where the cells occur in packets (Graham et al. 2016). In Microthamnion, the filaments can be highly branched (Graham et al. 2016). So far, no flagellate vegetative form has been identified in the Trebouxiophyceae (Lemieux et al. 2014a, b). Reproduction Asexual reproduction is predominant in the Trebouxiophyceae, i.e., sexual reproduction has not been observed in almost all genera. Most trebouxiophytes presumably are haplonts. Reproduction by autospores (Fig. 4.41) prevails; that is, in most known members, no motile cells (zoospores, Fig. 4.41) are formed. Those are called autosporin. Examples of reproduction by autospores are shown here with Myrmecia (Fig. 4.46d), Dictyochloropsis (Fig. 4.46h), and Leptosira (Fig. 4.49g). There are only a few lineages of

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Fig. 4.46 Trebouxiophyceae, unicellular coccoids of the lineages Trebouxiales (a–d), Lobosphaera clade (e, f), and Apatococcus (g–i); a, b Trebouxia angustilobata, lichen photobiont, culture strain SAG 2204); c Asterochloris echinata, lichen photobiont, culture strain SAG 2282; a– c spherical unicells with star-like chloroplast suspended in the middle of the cell and containing a pyrenoid, the nucleus lies in a cavity of the chloroplast; d Myrmecia biotorellae, culture strain CCAP 250/1, ovoid to pear-shaped vegetative cells with flat chloroplast appressed to the cell wall, autosporangia that are releasing numerous small autospores; e Lobosphaera tiroliensis, culture strain SAG 2007, spherical cells with cup-shaped chloroplasts; f Trochisciopsis tetraspora, culture strain SAG 19.95; vegetative cells with a wall sculpture, flat parietal chloroplast, nucleus central; g, h Dictyochloropsis splendida, culture strain SAG 2305 isolated from tree bark; g the same cell at two different focus views, left, the optical section with the central nucleus, right, the surface view showing net-like chloroplast structure; h autosporangium with gelatinizing wall, releasing many small autospores; i Apatococcus fuscideae, lichen photobiont, culture strain SAG 2523, large cell packets that slowly disintegrate, flat chloroplasts appressed to cell wall, nucleus central; a–c, e–i courtesy of Tatyana Darienko, with permission of SAG culture collection; d with permission of the CCAP culture collection, Oban, Scotland

Trebouxiophyceae where reproduction is also by zoospores, e.g., in Trebouxiales, and Microthamniales (Fig. 4.52). In Stichococcus, the rod-shaped cells divide into two progeny (Fig. 4.49b). Sexual reproduction is known for Eremosphaera (Fig. 4.49i). It exhibits oogamy, i.e., sperms fertilize egg cells to form dormant zygotes (haplontic life cycle, Fig. 2.9 a; Kies 1967). In Prasiola, sexual reproduction involves a diplohaplontic life cycle alternating haploid and diploid generations (Fig. 2.9c; Friedmann 1959).

Ecology and diversity of lifestyles There is a large number of Trebouxiophyceae species with aerophytic (aero-terrestrial) lifestyles. They live in thin biofilms or form crusts on the surfaces of hard substrates at the contact zone of the substrate with air. Examples are rock surfaces (Darienko and Friedl 2021), tree bark, and roof tiles or similar artificial substrates (Hallmann et al. 2016). Many trebouxiophytes can be found in the upper layer of soils, for example, in arid environments (e.g., Fučíková et al. 2014;

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Fig. 4.47 Trebouxiophyceae, unicellular coccoids of the lineages Watanabales (a–e) and Elliptochloris clade (f–i); a, b Watanabea reniformis, culture strain SAG 211-19b; a ovoid vegetative cells with flat parietal chloroplast that fills about half of the cell; b formation of autospores, i.e., the cells contain multiple protoplasts with chloroplasts; c Chloroidium ellipsoideum, culture strain SAG 3.95, vegetative cells with star-like shaped chloroplast, note autosporangia with autospores of different size (arrows); d Chloroidium antarcticum, culture strain SAG 2583, isolated from soil of a glacier forefield, King George Island, Antarctica, ovoid vegetative cells, some cells undergo protoplast division to form autospores; e Jaagichlorella luteoviridis, culture strain SAG 211-9a, spherical unicells and autosporangia that contain 2 or 3 autospores; f, g Elliptochloris, different species, spherical vegetative cells with flat chloroplasts appressed to the cell wall (parietal); f E. subsphaerica, culture strain SAG 2117, isolated from a roof tile (Germany); g E. perforata, culture strain SAG 2509, isolated from cells growing as epiphytes on a lichen, Karadag Nature reserve, Crimea, Ukraine; h, i Coccomyxa, inconspicuous differences in vegetative morphologies; h C. simplex, culture strain SAG 216-9a, ovoid to elongated vegetative cells with flat parietal chloroplasts, note many empty autosporangial walls; i C. arvernensis, culture strain SAG 216–1, vegetative cells are more spherical; a–i c courtesy of Tatyana Darienko, with permission of SAG culture collection

Darienko et al. 2019), where they can be important constituents of Biological Soil Crusts. Many members of Trebouxiophyceae are commonly engaged in different types of symbioses. The genera Trebouxia (Fig. 4.46a, b) and Asterochloris (Fig. 4.46c) are the most common lichen photobionts (Chap. 6; Muggia et al. 2018); they may not even be found free-living outside of symbioses. Symbiochloris, Coccomyxa, Elliptochloris, and

Stichococcus are widespread as lichen symbionts but are also frequently encountered in the free-living state. Some species of Elliptochloris can also live in symbiosis with cnidarians in marine environments (Letsch et al. 2009). More trebouxiophytes have been recognized as lichen symbionts only recently, e.g., certain species of Apatococcus (Fig. 4.46i, Zahradníková et al. 2017) and Chloroidium (Muggia et al. 2018). There is a respectable diversity of those

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Fig. 4.48 Trebouxiophyceae, unicellular coccoids, and colonies of the lineages Chlorellales (a–e), Prasiola clade (f, g); Choricystis/Botryococcus clade (h), Symbiochloris (i), and Xylochloris clade (j); a Chlorella vulgaris, culture strain SAG 9.95, small spherical cells with a cup-shaped chloroplast with prominent pyrenoid and empty wall of an autosporangium; b Muriella terrestris, culture strain SAG 2435; minute spherical vegetative cells, some undergo protoplast division to form autospores; c Dictyosphaerium ehrenbergianum, culture strain TRV-13-11, isolated from sandstone, Trakhtemyriv Natural Park, Ukraine, cells are embedded in mucilage and connected by a branching structure which originated from older cell wall material; d, e Heynigia riparia, culture strain SAG 2448; the same spherical colony with cells connected by remnants of cell walls at different optical views; d surface view; e optical section of the colony showing connection of the spherical cells by remnants of parental walls; f, g Coenochloris sp., cultured strain isolated LC 21, isolated from the surface of granite rock, La Campana, Chile, spherical vegetative cells embedded in mucilage, chloroplast fills about half of the cells; g mucilage stained with methylene blue; h Choricystis minor, culture strain SAG 251–2, minute spherical cells with flat chloroplast, division stage to form autospores and empty sporangial wall; i Symbiochloris irregularis, culture strain SAG 2154, ovoid vegetative cells with nucleus located in the center; j Xylochloris sp., culture strain SAG 2382, isolated from forest soil, Swabian Alp, Germany, spherical to ovoid vegetative cells with star-like shaped chloroplast; a-i c courtesy of Tatyana Darienko, with permission of SAG culture collection

trebouxiophytes that form minute spherical cells found in endosymbioses with ciliates, Hydra (freshwater Hydrozoa), and freshwater sponges. Some of those species seem to be restricted to symbiosis, i.e., are not found free-living outside of their hosts. Common examples of those specialized species can be found in Auxenochlorella (Fig. 4.50a), Chlorella (Fig. 4.48a), Choricystis (Fig. 4.48h), Coccomyxa

(Fig. 4.47h, i), and Micractinium (Pröschold et al. 2011; Darienko and Pröschold 2015; Pröschold and Darienko 2020). Only a few members of Trebouxiophyceae are known from aquatic habitats, e.g., phytoplankton. A prominent example is Botryococcus (Fig. 4.51). Botryococcus forms oil-producing aggregates of cells and is a common member of phytoplankton in alkaline freshwaters (Fig. 4.51). The

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Fig. 4.49 Trebouxiophyceae, unicellular coccoids, filaments, and cell packets (sarcinoid growth form) of the lineages Prasiola clade (a–e), Geminella (f), Leptosira clade (g), Pleurastrosarcina (h), and Oocystaceae (i); a Diplosphaera chodatii, culture strain SAG 49.86, minute vegetative cells, many undergo protoplast division; b Stichococcus bacillaris, culture strain SAG 249.80, rod-shaped vegetative cells that can form short filaments of 2–3 cells; c Prasiola crispa, culture strain SAG 43.96, filamentous stage with one stellate and axial chloroplast per cell; d Prasiolopsis calcarius, culture strain SAG 10.95, isolated from a loam wall in Basel, Switzerland, vegetative cells forming cell packets; e Desmococcus olivaceus, culture strain SAG 1.94, isolated from a granitic boulder, Northern Victoria Land, Antarctica; vegetative cells forming cell packets; f Geminella minor, culture strain SAG 22.88, filaments of cylindrical vegetative cells, with flat ribbon-shaped (laminate) chloroplast with a pyrenoid; g Leptosira terrestris, culture strain SAG 463–2, large spherical vegetative cells with a chloroplast appressed to the wall; h Pleurastrosarcina terriformae, culture strain SAG 2590, isolated from soil of the Atacama Desert, Chile, vegetative cells forming cell packets; i Eremosphaera viridis, culture strain SAG 228-4d, large spherical cells with several small discoid chloroplasts arranged in strands of cytoplasm, empty disintegrating autosporangium (remains of parental cell walls) forming mucilage; a–i courtesy of Tatyana Darienko, with permission of SAG culture collection

excreted oil efficiently contributes to the buoyancy of the colonies. The alga contains resistant polymers in its cell walls, and, therefore, can be fossilized (e.g., Schiller et al. 2022). It is believed to have contributed substantially to the deposition of high-grade oil shales and coals worldwide (Graham et al. 2016). Other examples of trebouxiophytes with aquatic lifestyle are Parachlorella and Micractinium. Several members of the Oocystaceae, a distinct lineage of

Trebouxiophyceae (Fig. 4.52), are known only from aquatic habitats, e.g., Eremosphaera (Fig. 4.49i). Several genera thrive in aquatic and terrestrial habitats. For example, Dictyosphaerium (Fig. 4.48c), frequently found in phytoplankton, has also been reported from terrestrial habitats (Mikhailyuk et al. 2020). Only very few members of Trebouxiophyceae are known from marine environments. A prominent example is

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Fig. 4.50 Trebouxiophyceae, Chlorellales, AHP clade; a Auxenochlorella protothecoides, culture strain SAG 211-7a, the strain is one of the oldest maintained in culture collections, i.e., in axenic condition since 1892 (Darienko and Pröschold 2015), minute spherical cells with flat parietal chloroplast without pyrenoid; b Prototheca zopfii, culture strain SAG 263-1, spherical unicells that reproduce by autospores, the species is an obligate heterotroph that contains chloroplasts which, however, are without photosynthetic pigments; a courtesy of Tatyana Darienko, with permission of SAG culture collection; b courtesy of Thomas Friedl, with permission of SAG culture collection

Fig. 4.51 Trebouxiophyceae, Botryococcus; a–f B. braunii; a– c, e, f culture strain Bot22; d culture strain Showa; a, b colonies of oval or spherical cells embedded in tough, irregular mucilage, numerous lipid globules are excreted from the cells; c, d overviews of irregularly shaped colonies; c vegetative cells at the periphery of a colony; d fluorescence microscopy, colony stained to show the excretion of oil droplets; e, f two neighboring ovoid cells with an oil droplet excreted between the cells; e bright field microscopy; f fluorescence microscopy, stained (yellow) showing small oil droplets formed in the cytoplasm of the cell, red color is due to the autofluorescence of chlorophyll; a–f courtesy of Christian Kleinert, Mario Salisch, and Carola Griehl, Competence Center Algal Biotechnology, Anhalt University of Applied Sciences, Köthen, Germany

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Pleurastrosarcina*

Prasiola clade* Coenocystis Neocystis clade Chloropyrula Leptosira* Apatococcus Leptochlorella unidentified

Parietochloris clade* Xylochloris clade Eremochloris Microthamniales*

Symbiochloris

Watanabea clade

Elliptochloris clade Choricystis/Botryococcus clade Lobosphaera clade* Xerochlorella Geminella Trebouxiales* outgroup Fig. 4.52 A schematic phylogenetic tree of the Trebouxiophyceae based on analyses of the 18S rRNA gene. Triangles represent collapsed major clades of the class. An asterisk marks a lineage where asexual reproduction by zoospores, in addition to that by autospores, is known. The phylogeny is redrawn from Fučíková et al. (2014), and the position of additional lineages (Darienko et al. 2019) is indicated by arrows. Names of lineages follow Li et al. (2019a, b) and Ueno et al. (2005). Most lineages are also resolved in a phylogeny of multiple chloroplast-encoded proteins (Lemieux et al. 2014a, b). For outgroup, sequences of Chlorophyceae, Volvocales, have been used

Prasiola which lives attached to hard substrate surfaces openly exposed to the air and salt spray. Other trebouxiophytes reported from the marine environment seem confined to a single lineage, Oocystaceae. For three genera of Trebouxiophyceae, a kind of endophytic lifestyle within plant tissue has been reported. There is an unprecedented “endosymbiosis” between the trebouxiophyte Coccomyxa and a higher plant, Ginkgo biloba (Tremouillaux-Guiller and Huss 2007). Within Ginkgo host cells, the alga resides in a “premature” state where the nucleus and mitochondria cannot be observed, and the chloroplast is still not functional. The alga becomes fully functional only after necrosis (damage) of the plant tissue when it is released. It may grow as an epiphyte on the plant surface and be retracted together with the pollen inside the pollen chamber of Ginkgo. How Coccomyxa escaped digestion by the plant tissue and finally became an inherent part of the host is still unknown. Species of Phyllosiphon occur in various subaerial microhabitats, e.g., on the surface of plants. They can also penetrate the intercellular matrix of the plant leaf parenchyma and thus cause leaf necrosis (Ma et al. 2013; Procházková et al. 2016). Within the plant tissue, they form filamentous structures that grow in the intercellular matrix of the leaf parenchyma. The filaments contain “endospores,” which serve the proliferation of the alga within the tissue. In culture, those “endospores” develop into autosporangia which release coccoid cells with a morphology that resembles other members of the Watanabales (Procházková et al. 2016). Similarly,

also species of Heveochlorella can occur as epiphytes (e.g., on the bark of the palm Roystonia regia), but one species, H. hainangensis, has also been found to grow between the bark and xylem of young rubber trees, Hevea brasiliensis (Zhang et al. 2008; Ma et al. 2013). Growth of the endophytic species is enhanced by organic compounds, but not in the epiphytic species, which may be regarded as an adaptation to the endophytic lifestyle (Ma et al. 2013). The Trebouxiophyceae also comprises species that have lost photosynthetic capacity and have evolved free-living or parasitic heterotrophic lifestyles, e.g., Prototheca (Fig. 4.50b) and Helicosporodium (Ueno et al. 2005; Lemieux et al. 2014a, b). Both depend on organic sources. Helicosporidium lives parasitically as an invertebrate pathogen. Prototheca (Fig. 4.50b) occurs in soils and organically enriched freshwater environments (Graham et al. 2016); it can cause skin infections in humans and cattle mammary glands. Though photosynthetic pigments are lacking in both genera, they contain chloroplasts. In molecular phylogenetic analyses, they share a common origin with the photosynthetic Auxenochlorella, forming the AHP (Auxenochlorella/Helicosporidium/Prototheca) clade (Ueno et al. 2005). Despite being photosynthetic, Auxenochlorella is dependent on organic compounds. Auxenochlorella also occurs in soils. It assumes a position basal to the obligate heterotrophs Prototheca and Auxenochlorella (Ueno et al. 2005; Darienko and Pröschold 2015; Plieger and Wolf 2021). This reveals the obligate heterotrophs

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Prototheca and Helicosporidium descendants of the photosynthetic Auxenochlorella (Fig. 4.50). Recognition and classification The scarcity of characteristic morphological features would make a traditional classification impossible so that the class could not be recognized as a distinct lineage of green algae before molecular phylogenetic analyses (Friedl and Rybalka 2012; Leliaert et al. 2012). However, there are a few morphological features seen in the chloroplasts that make the recognition of members of Trebouxiophyceae from cultures likely. Most members of Trebouxiophyceae are described from cultured material, which makes it one of the best-studied groups of green algae. Many genera of Trebouxiophyceae have chloroplasts with a stellate structure, e. g., Asterochloris (Fig. 4.46c), Trebouxia (Fig. 4.46a), and Xylochloris (Fig. 4.48j). In others, the chloroplast is flat and plate-like and appressed to the cell wall (parietal), e.g., Leptosira (Fig. 4.49g), Lobosphaera (Fig. 4.46e), and Myrmecia (Fig. 4.46d). Dictyochloropsis is an example of a net-like (reticulate) chloroplast (Fig. 4.46g). Often, the chloroplast is without a pyrenoid (e.g., Auxenochlorella, Fig. 4.51a, Botryococcus, Fig. 4.51, Coccomyxa, Fig. 4.47h, and Microthamnion), or the pyrenoid can be easily overlooked because its matrix is not covered by starch grains, e.g., in Asterochloris (Fig. 4.4c, g), Trebouxia (Fig. 4.46a), and Leptosira (Fig. 4.49g). Many of the morphological features of vegetative cells, as well as the growth forms, seem to have evolved independently in several not closer related lineages within the Trebouxiophyceae. Table 4.3 Examples of the various lineages of Trebouxiophyceae that are discussed in this chapter (see Fig. 4.52)

Ultrastructural features to define the class may be found in CCW-oriented flagellar bodies (Fig. 4.37d), a non-persistent metacentric spindle, and a phycoplast at cytokinesis (Fig. 4.38b). However, with those features alone, the class could be recognized as a lineage distinct from Ulvophyceae. Also, in only a few lineages of Trebouxiophyceae known today, motile cells (zoospores) are formed (Fig. 4.52). There is still debate about the phylogenetic position of the Trebouxiophyceae among the core Chlorophyta (Fig. 4.28), i.e., whether it is the sister group with Chlorophyceae or Ulvophyceae (see Fig. 2.8). Also, it has been discussed whether the two lineages of Trebouxiophyceae, Oocystaceae, and Chlorellales, together form a single monophyletic lineage which eventually is a separate lineage of green algae not belonging to the Trebouxiophyceae (Lemieux et al. 2014a, b). Molecular phylogenetic analyses show a remarkable phylogenetic breadth within the Trebouxiophyceae, despite the relatively simple vegetative morphology of trebouxiophytes (Fig. 4.52). However, the phylogeny of trebouxiophycean algae remains largely unresolved at deep levels. They are distributed on several monophyletic and well-resolved clades and lineages within the class. There is an array of about 22 distinct monophyletic lineages of Trebouxiophyceae known until now (Fig. 4.52). A current overview can be found in Fučíková et al. (2014), Lemieux et al. (2014a, b), Hallmann et al. (2016), Mikhailyuk et al. (2020), and Li et al. (2021). The examples for trebouxiophyceae genera discussed in this chapter are listed in Table 4.3. The relationship among the various clades and lineages, however, is mostly only poorly resolved

Phylogenetic lineages Chlorellales + Oocystaceae

Example genera Chlorellaceae

Chlorella (Fig. 4.48a), Dictyosphaerium (Fig. 4.48c), Heynigia (Fig. 4.48d, e), Muriella (Fig. 4.48b)

AHP clade

Auxenochlorella (Fig. 4.50a), Prototheca (Fig. 4.50b)

Oocystaceae

Eremosphaera (Fig. 4.49i)

Prasiola clade

Leptosira (Fig. 4.49g)

Apatococcus

Apatococcus (Fig. 4.46i), Dictyochloropsis (Fig. 4.46g, h)

Xylochloris clade

Xylochloris (Fig. 4.48j)

Watanabeales

Chloroidium (Fig. 4.47c, d), Jaagichlorella (Fig. 4.47e), Watanabea (Fig. 4.47a, b)

Elliptochloris clade

Elliptochloris (Fig. 4.47), Coccomyxa (Fig. 4.47h, i)

Choricystis/Botryococcus clade

Botryococcus (Fig. 4.51), Choricystis (Fig. 4.48h)

Lobosphaera clade

Lobosphaera (Fig. 4.46e), Trochisciopsis (Fig. 4.46f)

Geminella

Geminella (Fig. 4.49)

Trebouxiales

Asterochloris (Fig. 4.46c), Myrmecia (Fig. 4.46d), Trebouxia (Fig. 4.46a, b)

Pleurastrosarcina

Pleurastrosarcina (Fig. 4.49h)

Symbiochloris

Symbiochloris (Fig. 4.48i)

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(Fig. 4.52). It may be summarized as a large polytomy (Leliaert et al. 2012). Few of the lineages have formally been described as orders of Trebouxiophyceae, i.e., the Chlorellales, Microthamniales, Trebouxiales, and Watanabales (for an overview, see Li et al. 2021). Others are called informal clades, e.g., the Prasiola clade and Choricystis/Botryococcus clade (Karsten et al. 2005; Darienko et al. 2019), or have been given the name of a genus that they contain (Fig. 4.52). So far, only single gene analyses, i.e., the 18S rRNA gene, have widely been used for Trebouxiophyceae phylogeny (e.g., Fučíková et al. 2014). Analysis of 79 chloroplast DNA (cpDNA)-encoded proteins spanning the broad range diversity of the Trebouxiophyceae has supported most of the lineages of the 18S rRNA gene phylogeny (Lemieux et al. 2014a, b). It is anticipated that a better resolution of the phylogenetic structure within the Trebouxiophyceae will be achieved by combining multiple genes or phylogenomic analyses with the selection of a broad range of trebouxiophycean species (Fig. 4.52)

4.2.10 Class Chlorophyceae The Chlorophyceae is one of three most species-rich green algal classes. The class includes some of the most familiar microscopic green algae, including many model organisms. Its monophyletic distinction from other green algae is well supported by molecular and ultrastructural data. It is the only class in core Chlorophyta whose monophyly remains uncontested as gene and taxon sampling improves (Fučíková et al. 2019). The class is remarkable as its members display the greatest range of vegetative morphology. Flagellated and coccoid forms are present as unicells as well as distinctive colonies (coenobia), there are unbranched and branched filaments as well as coenocytic forms with multiple nuclei per cell. The broad range of body diversity of the Chlorophyceae is depicted here in Figs. 4.54, 4.55, 4.56, 4.57, 4.58, 4.59, 4.60, 4.61 and 4.62. Reproduction is equally diverse, including various asexual and sexual modes. Sexual reproduction involves zygotic meiosis, i.e., all members of Chlorophyceae have a haplobiontic life cycle (see Sect. 2.4, Fig. 4.40a; Lewis and McCourt 2004). The members of Chlorophyceae are especially abundant in freshwater but also occur in terrestrial habitats, e.g., soils. Mitosis is closed, and cytokinesis is mediated by a phycoplast. Among the core Chlorophyta, members of the Chlorophyceae exhibit the greatest variability at the level of flagellar apparatus (Lewis and McCourt 2004; Leliaert et al. 2012). The flagellar basal bodies of most members of Chlorophyceae are displaced in a clockwise direction when the flagellar cell is viewed from the top (CW; Fig. 4.37d), are directly opposed (DO, Fig. 4.37d), or in case of motile cells with four flagella, may have both arrangements, thus

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contrasting with the counter-clockwise arrangement (CCW) observed in the Trebouxiophyceae and Ulvophyceae. The stephanokont flagellation of Oedogoniales (see below) is outstanding among all green algae. Several green algae of the Chlorophyceae serve as model systems or are of economic importance (Leliaert et al. 2012). The biochemistry and physiology of the unicellular, halophilic Dunaliella salina have been studied in detail. This alga is among the most industrially important microalgae because it can produce massive amounts of b-carotene that can be collected for commercial purposes, and because of its potential as a feedstock for biofuels production. The unicellular flagellate Chlamydomonas reinhardtii (Fig. 4.54c) has long been used as a model system for studying photosynthesis, flagellar assembly and function, cell–cell recognition, circadian rhythm, and cell cycle control because of its well-defined genetics. The colonial green alga Volvox (Fig. 4.55h–j) has served as a model for the evolution of multicellularity, cell differentiation, and colony motility. Scenedesmus s.l. (Fig. 4.56a–c) and Pediastrum (Fig. 4.56 d–f) are also important paleoecological or limnological indicators. The broad diversity of vegetative morphology has traditionally been used to circumscribe higher taxa (orders, families) for classification within the class Chlorophyceae. This is especially true for the groups that are morphologically depauperate and exhibit convergent evolution towards a reduced morphology. Molecular phylogenetic and ultrastructural data have identified major clades and lineages within the Chlorophyceae (Fig. 4.53). This rendered several traditional orders originally circumscribed using vegetative morphology to contain phylogenetically unrelated taxa. For example, simply organized unicellular coccoid members of Chlorophyceae are unrelated to each other and distributed in almost all of the lineages recognized at present (Fig. 4.53). For our discussion of the diversity of the Chlorophyceae here we follow a division of the class as revealed from phylogenomic analyses, comprehensive in both the number of marker genes as well as the variety of species (Fučíková et al. 2019). A less formal clade system has been suggested as more useful for the class because dwelling on well-defined, strongly supported Linnean orders is not currently practical in Chlorophyceae. Because Sphaeropleaceae nomenclaturally defines Sphaeropleales, but both represent unrelated lineages, a new name is required for the lineage, and the informal name Scenedesminia has been used (Fučíková et al. 2019). In the following, the three lineages most species-rich and diverse in morphology, the Volvocales, Scenedesminia, and OCC-group (Oedogoniales, Chaetophorales, and Chaetopeltidales), will be discussed. These major lineages provide a good representation of the morphological diversity of the class.

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Scenedesminia Microsporaceae Dictyochloris Golenkinia Jenufa Sphaeropleaceae Treubarinia Spermatozopsis Volvocales OCC

Oedogoniales Chaetophorales Chaetopeltidales

Fig. 4.53 A simplified phylogenetic tree that corresponds to consensus trees inferred from analyses of concatenated nucleotide and amino acid chloroplast data (58 protein-coding genes); dashed lines indicate critical nodes still with low phylogenetic signal; redrawn and modified from Fučíková et al. (2019)

Unicellular flagellates of Chlorophyceae Most prominent examples of unicellular flagellates are the genera Chlamydomonas, Chloromonas, Dunaliella, and Haematococcus (Fig. 4.54). They are all members of the Volvocales. The genus Chlamydomonas as circumscribed in traditional systematics, was one of the largest genera of green algae, with more than 600 species. Molecular phylogenetic studies revealed it to encompass various distinct phylogenetic lineages, which nowadays are regarded as distinct genera, e.g., Lobochlamys and Oogamochlamys (Pröschold et al. 2001). Sexual reproduction is known in only a few species. Most are isogamous, but anisogamy or oogamy has also been observed. Most species are heterothallic, i.e., they consist of an mt+ and mt− strain. Gametes are indistinguishable from vegetative cells, but they do have multiple agglutinins on the flagella, which will promote adhesion of flagella of opposite mating types. The life cycle of Chlamydomonas is shown in Fig. 4.40a. Cellulose is absent from the cell wall of Chlamydomonas, instead, there is a glycoprotein layer contained in the wall. Chlamydomonas produces carbohydrates in a high proportion of dry weight (50%, especially starch), which can be further hydrolyzed using microorganisms (bacteria, fungi) to bioethanol for biofuel production (Saini et al. 2020). Chloromonas (Fig. 4.54e, f), a close relative of Chlamydomonas within the Volvocales, is associated with the phenomenon of “colored snow, i.e., colored blooms of algae on snowfields and glaciers in the high mountains and Polar regions. Biflagellate vegetative cells of

Chloromonas are dominant in greenish snow. Nonmotile, dormant cysts or zygotes with a thick cell wall accumulate orange carotenoid pigments within the cell and then dominate in reddish snow (Matsuzaki et al. 2019). The flagellates Dunaliella (Fig. 4.54g, h) and Haematococcus (Figs. 4.1 and 4.54i–k) are commercially exploited to produce the carotenoids b-carotene and astaxanthin, valued for their colorant and antioxidant properties (Graham et al. 2016; Pagels et al. 2020). Dunaliella occurs in extremely saline waters. It uses glycerol to balance high external osmotic pressures and produces high levels of b-carotene under high salinity and irradiance conditions. Therefore, Dunaliella is used for the industrial production of glycerol and b-carotene. With Dunaliella salina the industrial production of high-value compounds from microalgae began in the 1980s (Borowitzka 2013; Pagels et al. 2020). Haematococcus in nature is found in shallow temporary water bodies (e.g., depressions in rock or on man-made substrates). Under favorable conditions, it occurs as motile biflagellated unicells whose protoplast is connected to the cell wall by multiple strands of cytoplasm (Figs. 4.1 and 4.54i, j). It forms cysts under unfavorable conditions capable of surviving complete desiccation and can be transported in the wind (Fig. 4.54k). The cysts accumulate astaxanthin, responsible for their red coloration. A motile colonial form, Stephanosphaera, is morphologically somehow linked with Haematococcus, having cells with protoplastic extensions (Fig. 4.55a–c). Both genera are also phylogenetically closely linked within the Volvocales (see below; Nakada et al. 2008). Spermatozopsis, which has been used as model organism in various cell biological studies, may represent the only

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Fig. 4.54 Chlorophyceae, unicellular Volvocales; a Chlamydomonas noctigama culture strain SAG 36.72; C. zebra culture strain SAG 39.86; c C. reinhardtii culture strain SAG 34.90; d zygote of C. cf. suboogama (culture strain isolated from soil, Chiloe, Chile), note characteristic reddish coloration due to carotenoid accumulation; e Chloromonas typhlos culture strain SAG 26.86; f field-collected cysts or zygotes of snow-inhabiting C. miwae in optical section (left) and surface view (right); g Dunaliella granulata culture strain SAG 41.89; h D. salina culture strain SAG 184.81; i, j Haematococcus pluvialis, flagellated stage, field material; i swimming cells with flagella; j central nucleus, several pyrenoids and mucilaginous sheath; k cysts of H. pluvialis culture strain SAG 34-1a, red color due to accumulation of the carotenoid astaxanthin; a–c, e, g, h courtesy of Tatyana Darienko, with permission of SAG culture collection; d original Tatyana Darienko; f reproduced from Matsuzaki et al. (2019); i, j courtesy of Christian Linkenheld; k courtesy of Nataliya Rybalka, with permission of SAG culture collection

unicellular flagellate of Chlorophyceae with a phylogenetic position independent of Volvocales (Fig. 4.53). Motile colonial forms of Chlorophyceae A unique feature of Volvocales is that it includes several conspicuous motile colonial forms. In Stephanosphaera the cells are arranged in a ring, and the colony is enclosed in a globular mucilaginous matrix (Fig. 4.55a–c). Other genera of Volvocales occur as motile colonial forms of Chlamydomonas-like cells found in multiples of two. Flagellar action of the cells within a colony must be coordinated to enable a directed movement of the colony towards the light (positive phototactic movement). How the cells coordinate their flagellar beats is still poorly known. The colonial motile forms

define a monophyletic group within the Volvocales (Graham et al. 2016). The number of cells in the colony and colonial shape are genetically determined and specific for each genus. In some genera, all cells of a colony are capable of asexual reproduction by successive bipartition of parental walls to produce new colonies (autocolonies). In others, only certain cells, called gonidia, can generate daughter colonies. Sexual reproduction starts with the formation of gametes by successive division of vegetative cells into gametes. In Volvox, gonidia are also transformed into egg cells. Fusion of gametes is then by isogamy (gametes indistinguishable by size, e.g., Pandorina), anisogamy (with both gametes of unequal size, e.g., Pleodorina, Fig. 4.55g) or oogamy (immotile female gamete by far larger than the tiny motile male gamete, e.g., Volvox, Fig. 4.55h–j). The zygotes are spiny

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Fig. 4.55 Chlorophyceae, colonial Volvocales; a–c Stephanosphaera pluvialis, different views of colonies, arrangement of cells in a ring, colony enclosed in a globular mucilaginous matrix (field material, courtesy of Christian Linkenheld); c phase-contrast microscopy; d Gonium pectorale, planar colony of 16 flagellated cells, note gelatinous strands interconnect the cells (field material, courtesy of B. Büdel); e–f Pandorina morum, colony forming a globular ball-like cluster of few (8–16) flagellated cells, culture strain SAG 60-1b; e close up of the colonies, note dark eyespot visible in two cells, mucilaginous sheath enclosing each colony, and flagella; f colony with formation of daughter colonies (upper half), each cell of a colony divided to form new colonies; g Pleodorina californica, spheroid colony, posterior portion of the colony with larger cells that are reproductive gonidia, the others are somatic cells that cannot divide, culture strain SAG 32.94; h–j Volvox globator, large hollow spheroid colony, culture strain SAG 199.80; h two large colonies that enclose developed smaller daughter colonies; i large colony with small daughter colonies developing from gonidia; j part of a large colony showing the many somatic cells interconnected by protoplasmatic strands; a–c courtesy of Christian Linkenheld; d original Tatyana Darienko; e–g originals of Thomas Friedl, with permission of the SAG culture collection

and thick-walled and germinate by meiosis. The life cycle is haplontic (see Chap. 2; Fig. 2.9). The colonial Volvocales commonly occur in the summer plankton of freshwater lakes. In Gonium, a small number of cells (8–32, the number depending upon the species) forms a planar colony. The cells are interconnected by mucilage strands (Fig. 4.55d). All cells are capable of forming daughter colonies. For sexual reproduction, the colony dissociates into single cells, subsequently functioning as isogametes. Colonies of Pandorina consist of a globular ball-like cluster with 8 or 16 biflagellated cells closely adherent at their bases and embedded in mucilage

(Fig. 4.55e, f). There is already a differentiation of the cells within the colony, i.e., the eyespots of some cells are larger than those of others marking some degree of colony polarity. In contrast to Gonium and Pandorina, the colonies of Eudorina, Pleodorina, and Volvox are spheroids. They have their constituent cells arranged as a single layer at the periphery of the mucilaginous sphere (Pickett-Heaps 1975). A specialization is encountered among those genera as the number of cells per colony increases; it may be regarded as an evolutionary advancement. In Eudorina, all cells are of the same size, and almost all cells except a few (e.g., 24 out of the

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28 in E. elegans) can divide vegetatively, i.e., produce daughter colonies. In Pleodorina the colonies contain around 128 cells, and about half of the cells (in the posterior portion of the colony) are larger and reproductive gonidia (Fig. 4.55g); the other (smaller) cells cannot divide and are called somatic cells. The specialization of vegetative reproduction reaches its peak in Volvox, whose giant colonies may contain more than a few hundred or thousands of cells (Pickett-Heaps 1975). Of these cells, only very few (16 or less) can give rise to daughter colonies. Those gonidia are not flagellated, and, therefore, the flagellar action of the tiny somatic cells is essential to keep the large colony motile. Typically, protoplasmatic strands interconnect adjacent somatic cells, and they function to enable the coordination among the cells (Fig. 4.55j). Volvox forms a hollow sphere, and the daughter colonies originating from the gonidia are formed within the parental colony (Fig. 4.55h–j). The daughter colonies are only released when the parental colony falls apart and dies (Pickett-Heaps 1975). The daughter colonies develop with their cell apices facing inward, and then the colony must invert so that the flagella are oriented toward the outside. This process, called inversion, may be compared with the reversing of a glove when taking it off. For details of the inversion during the daughter colony formation of Volvox, see Graham et al. (2016) and Pickett-Heaps (1975). Coccoid body forms of Chlorophyceae The coccoid body form is widespread in the Chlorophyceae and, like the motile flagellated forms, also occurs as colonies. This body form is of multiple origins within the Chlorophyceae, i.e., distributed on various phylogenetic lineages of the class phylogenetics (Fučíková et al. 2019), i.e., the Volvocales, Scenedesminia, Sphaeropleaceae, Treubarinia, Golenkinia and Jenufa, and Dictyochloris (Fig. 4.53). Scenedesminia describes a monophyletic lineage in phylogenomic analyses whose members have previously been included with the family Sphaeropleaceae in the Sphaeropleales (Fučíková et al. 2019). Coccoid forms of the Volvocales feature motile zoospores formed during asexual reproduction, e.g., Chlorococcum and Chlorosarcinopsis. In Chlorococcum, the biflagellated zoospores resemble very much Chlamydomonas cells, so that Chlorococcum cells may be interpreted just as cysts of Chlamydomonas with the coccoid cyst stage being predominant in the life cycle. Both may be closely connected; see life cycle in Fig. 4.41. A particular feature of coccoid body forms in the Chlorophyceae is that several genera form specialized colonies, i.e., coenobia. A coenobium is a colony where the number of cells is fixed at its origin and not augmented subsequently (Bold and Wynne 1978). A coenobium has a defined shape that is genetically fixed and characteristic of the species. In asexual reproduction, a miniature of the

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parental colony is called autocolony (Fig. 4.56b). Each cell of a coenobium is capable of forming autocolonies. Prominent examples of non-motile coenobia-forming Chlorophyceae are Scenedesmus s.l., Coelastrum, Pediastrum, and Hydrodictyon (Fig. 4.56). In the Scenedesmus sensu latu complex, the cells are cylindrical (elliptical to spindle-shaped) and laterally joined, usually in groups of 4 or 8 in one row (sometimes even in two rows), to form flat coenobia (Fig. 4.56a, c). This morphology is unique among all green algae. The cells have a laminate chloroplast that contains a prominent pyrenoid. Under certain culture or environmental conditions, Scenedesmus s.l. can be entirely unicellular. There are even species known where unicellular stages are predominant. Asexual reproduction is by autocolony formation in which each parental cell forms a miniature colony that is liberated through a tear in the parental wall. A large number of species, i.e., more than 250, are regarded as taxonomically accepted (Guiry and Guiry 2023) in the Scenedesmus s.l. complex. A high degree of morphological plasticity impedes the unambiguous distinction of species in the Scenedemus s. l. complex. The analysis of DNA data is, therefore, the most reliable way to identify and differentiate species in this group (Terlova and Lewis 2019). The quest to separate the complex into phylogenetically meaningful units has been discussed controversially. The genus Desmodesmus has been first distinguished from Scenedesmus by molecular phylogenetic analyses (An et al. 1999). The distinction of Desmodesmus is congruent with morphology. Desmodesmus species have one or several spines on the cells (Fig. 4.56a, b). Their cells have submicroscopic structures on the outermost wall layer, often visible under the light microscope as granulations or ribs on cells (Vanormelingen et al. 2007). Presently, it is generally accepted that there are two genera, Desmodesmus and Tetradesmus, distinct from Scenedesmus sensu strictu. In Tetradesmus species, the cells are spineless, and the wall exhibits a smooth surface. Remarkably, within the same genus Tetradesmus several species are desiccation-tolerant and occur in temperate terrestrial environments and deserts, while others have an aquatic lifestyle (e.g., Terlova and Lewis 2019; Terlova et al. 2021). This allows studying the transition from aquatic environments to land. Otherwise Scenedemus s.l. is common in fresh and brackish waters, particularly in nutrient-rich conditions. They may constitute a major part of the green algal biomass in freshwater phytoplankton. They are important for many applications, e.g., in wastewater treatment studies and producing compounds of commercial interest, such as biodiesel and certain carotenoids (Molino et al. 2020; Turiel et al. 2021). Pediastrum also consists of flat coenobia. The marginal cells have one or two horn-like protuberances and, in some species, even the internal ones (Fig. 4.56d, e). During

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Fig. 4.56 Chlorophyceae, Scenedesminia; coenobia-forming coccoid forms; a, b coenobia of laterally joined cells in species of Desmodesmus, each end cell of a coenobium is with two spines; note prominent pyrenoid in the center of each cell; a D. armatus, culture strain SAG 276-4d; b D. communis, culture strain SAG 276-4b; larger coenobium in the upper half of the image is with two empty cells where the formation of autocolonies happened; c unicellular stage of Scenedemus vacuolatus, culture strain SAG 32.88; note some autosporangia where daughter cells are kept together with empty sporangial walls; d–f Pediastrum duplex, different culture strains; d flat coenobium of strain SAG 261-2, marginal cells with two horn-like protuberances each, note prominent pyrenoid in each cell; e fully developed coenobium, strain AL0403MN; f autocolony formation in strain SAG 28.83; g Hydrodictyon reticulatum, part of a cylindrical young coenobium, field material, scanning electron micrograph; h overview of a large coenobium in culture; i, j Coelastropsis costata, culture strain SAG 32.88, note cell walls are with prominent ridges; j Coelastrum morus, culture strain SAG 2078, spherical cells are connected by blunt processes to form more or less spherical hollow colonies; a–d, h–j courtesy of Tatyana Darienko, with permission of SAG culture collection; e reproduced from McMagnus et al. (2018); g courtesy of Michael Schagerl

autocolony formation (Fig. 4.56f), the protoplast of some or all cells of the coenobium undergoes divisions to form zoospores. They move freely within a vesicle which is the emergent inner layer of the parental wall. After a short period of motility, the zoospores aggregate in one plane and, as they grow, develop the cellular form characteristic of the species (Bold and Wynne 1978). During sexual reproduction, two

isogametes per cell are produced. After a period of dormancy, the zygote germinates by forming zoospores that rapidly transform into non-motile thick-walled polyhedral cells, called polyeders, and new coenobia arise within the polyhedral cells upon their germination. Hydrodictyon forms spectacular cylindrical coenobia that are closed at their ends (Fig. 4.56g, h). The coenobia

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Fig. 4.57 Chlorophyceae, Scenedesminia; unicellular coccoid forms; a Raphidocelis subcapitata, culture strain SAG 61.81, sickle-shaped cells; this strain is a very commonly used test strain for algal growth inhibition tests in ecotoxicology; e.g., Yamagishi et al. (2020); b, c Monoraphidium griffithii, culture strain SAG 202-13; b long, needle-shaped cells with parietal chloroplast; c asexual reproduction by cleavage of parental cells into few parallel autospores that are released by parental wall rupture; d Ankistrodesms densus, culture strain SAG 202-1, long, needle- or spindle-shaped cells aggregated into colonies; e Bracteacoccus minor, culture strain SAG 61.80, cells are spherical unicells that contain multiple nuclei and many discoid plastids without pyrenoids; f Chlorosarcinopsis arenicola, culture strain SAG 14615-2a; large packets of cells with a pyrenoid developed from cell division after which the daughter cells adhere to each other before they dissociate; g Protosiphon botryoides, culture strain SAG 731-1a; cells of different sizes with reticulate chloroplast with a pyrenoid, larger cells are multinucleate (called coenocytes); a– g courtesy of Tatyana Darienko, with permission of SAG culture collection

component cells themselves are cylindrical as well. They contain many nuclei, i.e., are coenocytic, with a large central vacuole, and can be up to several millimeters long. The cells are arranged into end-to-end polygonal configurations in which three to nine cells may be joined (Fig. 4.56g, h). This results in a net-like appearance of the coenobia, and nets up to a meter long occur (Bold and Wynne 1978). Like Pediastrum, asexual reproduction on Hydrodictyon (autocolony formation) is conferred by zoospores as well, but in Hydrodictyon, they move within the cell. Cleavage of the thin cytoplasm layer results in uninucleate zoospores, which move freely within the space provided by the cell wall and vacuolar envelope. The zoospores ultimately retract their flagella, become joined, and aggregate to initiate young coenobia. Therefore, the cell wall and the vacuole comprise a mold in which the young nets are organized before they are liberated after disintegration of the parental cell wall (Bold

and Wynne 1978). Pediastum and Hydrodictyon are close relatives, their sister-group relationship within the family Hydrodictyaceae has been supported by phylogenomic analyses (McManus et al. 2018). Sexual reproduction is accomplished by biflagellated isogamous gametes formed within the cell lumen and shed into the water through a pore in the parental wall. Like in Pediastrum, the zygote forms four zoospores upon germination, which settle down to form polyhedral cells. In Coelastrum, the coenobia are hollow spheres composed of 4 to many cells. Depending on the species the uninucleate cells may be united by extensions of their cell wall (Fig. 4.56i, j). Asexual reproduction is by the formation of autocolonies by any or all the cells of a mature coenobium (Bold and Wynne 1978). The unicellular coccoid members of Chlorophyceae are also rather diverse in morphology. Typical simple unicellular forms are Bracteacoccus (Fig. 4.57e), Dictyochloris, and

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Jenufa (Fig. 4.58a, b). Chlorosarcinopsis forms characteristic cell packets of autospores that adhere to each other for some while (Fig. 4.57f). Asexual reproduction is by biflagellated zoospores without cell walls. Ankistrodesmus and Monoraphidium (Scenedesminia) have needle or spindle-shaped cells (Fig. 4.57b–d). Both genera, as well as Jenufa, are examples of coccoid green algae where asexual reproduction is exclusively by autospores; zoospores have not been observed in those genera. Ankyra and Atractomorpha in the Sphaeropleaceae are other examples of a genus with needle-shaped cells (Fig. 4.58e–g). The vegetative cells of Ankyra have needle-like cell poles (Fig. 4.58e). Atractomorpha has fusiform spindle-like, highly elongated cells (Fig. 4.58f). In young cells, the chloroplast appears as a diffuse reticulum, whereas it disassociates into band-like components in mature vegetative cells (Hoffman 1983). Three other genera of Chlorophyceae are distinct due to their peculiar cell wall extensions. Golenkinia is a common genus characterized by spherical cells with long protrusions of its cell wall (Fig. 4.58c, d). Treubaria has 3–4 appendices on its cell wall so that the cells form a tetrahedron or a triangle (Fig. 4.58g). Trochiscia is characterized by short spines that cover the cell wall (Fig. 4.58i). Coenocytic organization of cells also occurs in several lineages of the Chlorophyceae, i.e., in Dictyochloris (Fig. 4.58a), Hydrodictyon of Scenedesminia (Fig. 4.56g, h), Protosiphon of Volvocales (Fig. 4.57g). Protosiphon is a widely distributed soil alga that has long sac-shaped cells which can reach up to 1 mm with multi nuclei (called coenocytes), which develop when repeated nuclear divisions are not followed by cytokinesis. Those vegetative sacs develop under natural conditions, whereas simple coccoid stages with few or single nuclei prevail in laboratory cultures (Fig. 4.57g). OCC clade—the filamentous Chlorophyceae Most filamentous genera of Chlorophyceae (except Cylindrocapsa, Microspora and Sphaeroplea) are within a well-supported monophyletic clade that comprises three larger lineages, i.e., the Oedogoniales, Chaetophorales, and Chaetopeltidales (OCC). The Oedogoniales is well defined by several outstanding features not found in any other green algae. It comprises three genera, Bulboachaete, Oedocladium, and Oedogonium. The genus Oedogonium comprises a large number of species; Guiry and Guiry (2023) currently list >850 accepted species names indicating that the group is in urgent need of taxonomic revision. Filaments of Oedogonium are composed of elongated cells with reticulate chloroplasts containing pyrenoids, all surrounding a central vacuole. In Oedogonium the filaments are unbranched (Fig. 4.59a–d). Bulbochaete has branched filaments with most of the cells with long, bulbous-based colorless bristles

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or hairs. The nuclei lay closely attached to the wall. Cell division results in distinctive rings that occur in the walls near one of the cells (Figs. 4.59b, c and 4.60a). After mitosis, a ring-like thickening occurs near one end of the cell. Later, the outer wall layer splits in the vicinity of the ring-like structure, and the latter becomes elongated into a cylinder as the protoplasts of the two daughter cells elongate. The expanding ring forms the initial outer wall of the cell. Repeated divisions cause the accumulation of ring-like scars near one end of the cell, i.e., cells that have undergone many divisions will exhibit many such rings (Fig. 4.59c). Asexual reproduction is both by fragmentation and by zoospore production. A single wall-less zoospore develops in the parental cell (Fig. 4.60c). There is a ring of many flagella (approx. 120) that surrounds the cell attached to a colorless, anterior apical dome (Fig. 4.60c, d). This unique type of flagellation is called stephanokontan. The zoospores also possess an eyespot and numerous contractile vacuoles. They emerge through a rupture of the vegetative cell wall (Fig. 4.60c). After a period of motility, the zoospores attach to the substrate at their colorless poles and shed their flagella (Bold and Wynne 1978). Sexual reproduction is by oogamy. The nonflagellate egg cell develops within an oogonium which looks like a somehow swollen, rounded cell (Figs. 4.59d and 4.60b). There are species in which the sperm develops within cells of regular filaments. Then the antheridia are minute, short-cylindrical cells, each producing two multiflagellate sperms which bear about 30 flagella. The sperm are attracted to oogonia by chemical signals (pheromones). In other species, there is a pronounced dimorphism between the female and male filaments, i.e., the male filament being a few-celled dwarf and epiphytic on the female filament, called to be nannandrous in contrast to the macrandrous species with large male filaments. In the nannandrous species special zoospores are formed, intermediate in size between sperms and zoospores, termed androspores. They are formed in special androsporangia which may be present in the same filament as the oogonia or other filaments. The androspore is attracted by chemical signals to the oogonium and settles upon a cell near the oogonium. It undergoes a few mitotic divisions to form a short few-celled antheridium which then produces sperm (Figs. 4.59d and 4.60b). Upon fertilization, a thick-walled zygote develops within the oogonium while the rest of the filament dies. The Chaetophorales includes mostly branched filaments (e.g., the genera Chaetophora, Draparnaldia, Stigeoclonium, Fig. 4.61a–f) with only a few members forming unbranched filaments (e.g., Uronema, Fig. 4.61g, h). The elongated cells in a filament contain a single flat (parietal) chloroplast with prominent pyrenoids (encircled by a starch sheath). A distinct feature is that the branched filaments terminate in multicellular thin colorless hairs (Fig. 4.61f). In

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Fig. 4.58 Chlorophyceae, incertae sedis, coccoid and pseudofilamentous forms; a Dictyochloris fragrans, culture strain SAG 220-1b; mature vegetative cells with an irregular net-like chloroplast and several nuclei; b Jenufa aeroterrestrica, culture strain SAG 2383, spherical or slightly ellipsoidal vegetative cells with a single parietal (appressed to the cell wall) chloroplast and central vacuole; c, d Gloenkinia longispicula, culture strain SAG 73.80, spherical cells with spiny projections; c parietal cup-shaped chloroplast with a prominent pyrenoid surrounded by starch grains, the chloroplast may appear as a bell with a single opening; d focus on spiny projections; e Ankyra judayi, culture strain SAG 17.84, spindle-shaped vegetative cells with needle-like cell poles, chloroplast parietal with pyrenoid; f Atractomorpha echinata, culture strain SAG 70.90, vegetative cells are spindle-shaped with highly elongated cell poles (fusiform), chloroplast is a diffuse net-like structure; g–i Treubarinia; g Treubaria schmidlei, culture strain SAG 36.83, cell with several cell wall appendages (upper part of the image), they form a tetrahedron or triangle with 3–4 appendages surrounded by mucilage (lower part of the image, India ink preparation to visualize the cell wall appendices), note empty cell walls beside life cells; h Cylindrocapsa geminella, culture strain SAG 18.94, chloroplast star-like shaped with a central pyrenoid, thick stratified cell wall (arrow), filaments tend to fragment easily (called pseudofilamentous); i Trochiscia gutwinskii, culture strain SAG102.80, spherical vegetative cells with the wall covered by spines, chloroplast net-like with prominent pyrenoid surrounded by starch grains; a–f, h courtesy of Tatyana Darienko, with permission of SAG culture collection; g, i courtesy of Maike Lorenz, with permission of SAG culture collection

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Fig. 4.59 Chlorophyceae, filamentous forms, Oedogoniales (OCC clade); a–d Oedogonium sp. from freshwater samples; a filament composed of elongated cells with chloroplast in surface view, note net-like (reticulate) structure of chloroplast; b two filaments at different optical focuses, left filament at optical section showing in each cell the chloroplast with several pyrenoids, right filament at the surface view, chloroplast with several pyrenoids encircled by starch grains, arrows indicate the distinctive rings that occur in the walls near one of the cell; c filament with focus on the accumulation of ring-like scars near one end of the cell (arrows); d filament with oogonium (og) and the remnant of an antheridium (arrow); a, b courtesy of Nataliya Rybalka; c, d courtesy of Burkhard Büdel

Stigeoclonium there is a differentiation between prostrate and erect filament systems (Fig. 4.61a, b), the latter terminating in multicellular hairs with only a thin layer of mucilage. Draparnaldia is differentiated into an erect main axis and subordinate lateral branch systems, terminating with multicellular hairs (Fig. 4.61f). The main axis has characteristic band-like chloroplasts (Fig. 4.61e). For Chaetophora, it is very characteristic that in nature, the filaments are embedded in globular gelatinous colonies attached to underwater substrates. After cell division, the daughter cells remain in contact through plasmodesmata (Caisová et al. 2011). Asexual reproduction is by filament fragmentation and zoospores with four flagella (quadriflagellate) of which one or two per cell are formed. In those quadriflagellate motile cells the flagellar cruciate root system displays a mixed form, i.e., the upper basal bodies are directly opposed (DO, Fig. 4.37d), the lower basal bodies are in clockwise absolute orientation (CW, when the flagellated cell is viewed from the top, Watanabe and Floyd 1989; Fig. 4.37d). After a period of motility, they settle the apical side down onto substrates. Most species of Chaetophorales are known from freshwaters. Just a few are terrestrial. The Chaetopeltidales are known from soil or shallow freshwater environments in which the algae can often be exposed to desiccation and strong light (Watanabe et al. 2016). Their thalli consist of unicells, sarcinoid aggregations, or short-branched and unbranched filaments (Fig. 4.62a), sometimes with disk-shaped thalli and pseudocilia (Chaetopeltis, Pseuduvella). A distinct feature is that in freshwater, they live as aquatic epiphytes, e.g., on

freshwater plants (Chaetopeltis, Koshicola, Fig. 4.62, Pseudulvella). In contrast, the genera Hormotilopsis and Floydiella are known from soils. The Chaetopeltidales are characterized by quadriflagellate zoospores (Fig. 4.62b) in which both pairs of flagellar basal bodies are with the directly opposed (DO) arrangement (Fig. 4.37d; O’Kelly et al. 1994). Summary of Chlorophyceae classification A summary of the current classification of the less formal system of Chlorophyceae is given above (Table 4.4) (Fučíková et al. 2019). For most lineages (clades), examples are represented in Figs. 4.54, 4.55, 4.56, 4.57, 4.58, 4.59, 4.60, 4.61 and 4.62. Because a classification based on well-defined, strongly supported Linnean orders has been found not practical for the Chlorophyceae, we also abstain from referring to families here. The lineages listed below may represent taxa of the level of orders.

4.2.11 Class Ulvophyceae The Ulvophyceae include more than 1,700 species (Guiry 2012). Most species are macroscopic and occur in coastal environments (green seaweeds), where they often form ecologically important components (Brodie et al. 2007; Huisman 2015). A considerable diversity comprises microscopic algae and several species are found in freshwater habitats or moist subaerial habitats including soil, rocks, tree

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Fig. 4.60 Unique features of Oedogonium; a–c diagrammatic representations; a cap (ring) formation during cell division, ring-like structure in the cell wall at some distance below last downward-facing cap, the wall breaks at the predetermined site and the material of the ring, now exposed to the outside, is stretched into a cylinder as the daughter cells expand, the newly formed septum (se) moves up until it is past the newly formed upward-facing cap; b sexual reproduction in a nannandrous species, oogonium with egg cell slightly contracted from the cell wall and fertilization pore formed while oogonium secretes an envelope of mucilage, dwarf male filament (antheridium) which is about to release two sperm cells is attached to a cell near the oogonium; c asexual reproduction by zoospores, the whole content of a cell has differentiated into a zoospore with a distinctive ring of flagella and it emerges after the parental wall ruptured circumferentially at the wall weakening, still temporarily enclosed by a hyaline vesicle (left), elongated zoospore going to be attached with its apical dome to the substrate; d scanning electron micrograph of a zoospore showing the ring of flagella at the dome; a–d reproduced from Pickett-Heaps (1975), with permission of Oxford University Press through PLSclear

bark, and leaves (Škaloud et al. 2018). An iso- or heteromorphic diplohaplontic life cycle is found in most groups of ulvophytes, but several species (mainly in freshwater habitats) are only known to reproduce asexually. Traditionally, the class has been circumscribed based on a set of ultrastructural characteristics, including a counter-clockwise orientation of the flagellar root system, cytokinesis by furrowing, a closed persistent mitotic spindle, and the absence of a phycoplast (Mattox and Stewart 1984; O’Kelly and Floyd 1984; Sluiman 1989; Floyd and O’Kelly 1990). Some species have flagellate reproductive cells with cell walls or flagella covered by organic body scales (Sluiman 1989). However, because all these features also occur in other groups of green algae, and thus are not unique to the Ulvophyceae, monophyly of the class has been questioned (Leliaert et al. 2012).

The Ulvophyceae exhibit a large diversity of thallus as well as cellular organizations, which have been categorized into four main types (Cocquyt et al. 2010). The first type consists of flagellate or non-motile unicellular or colonial algae with uninucleate cells, and is present in some species of Ulvales, Ulotrichales, Chlorocystidales, Scotinosphaerales, Oltmansiellopsidales, and Ignatiales (Chihara et al. 1986; Nakayama et al. 1996; Friedl and O’Kelly 2002; Watanabe and Nakayama 2007; Škaloud et al. 2013). The second type comprises multicellular filaments or blades composed of uninucleate cells, and occurs in the Ulvales, Ulotrichales and Trentepohliales. The third type is the siphonocladous thallus organization, which is characterized by multicellular thalli composed of multinucleate cells with nuclei organized in regularly spaced cytoplasmic domains (Motomura 1996; McNaughton and Goff 1990). This type is

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Fig. 4.61 Chlorophyceae, filamentous forms, Chaetophorales (OCC clade); a, b Stigeoclonium aestivale, culture strain SAG 410-1, basal system of branches; a overview; b details at higher magnification, note tapering of side branches; c Chaetophora sp., culture strain SAG 51.89, irregular branching, cells with band-shaped chloroplast and conspicuous pyrenoid; d–f Draparnaldia glomerata, field material (freshwater creek Kaltengrundbach, Germany, Spessart midlands); d overview, showing the habit of growth, main axis consisting of markedly larger cells than those of the branches that originate at one point near the transverse walls of the axial cell (Bold and Wynne 1985), the branches terminate in a gradually tapering multicellular hair; e segment of the main axis with cells containing chloroplasts that are fimbriate and with several pyrenoids; f detail of a branch, each filament with terminal gradually tapering multicellular hair; g, h Uronema confervicola, culture strain SAG 386-2; g unbranched filaments that are fragmenting, with cells that each contain one nucleus and a parietal chloroplast with a prominent pyrenoid; h detail of a filament, basal cell with a thickening of the cell wall; a, b, g, h courtesy of Tatyana Darienko, with permission of SAG culture collection; c courtesy of Anastasiia Kryvenda, with permission of SAG culture collection; d–f courtesy of Burkhard Büdel

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Fig. 4.62 Chlorophyceae, filamentous forms, Chaetopeltidales (OCC clade), Koshicola spirodelophila, culture strain NIES-3575; a filamentous growth in culture, with highly vacuolated cells containing prominent pyrenoids; b different views of quadriflagellated zoospores, anterior vacuole, posterior chloroplast; a, b reproduced from Watanabe et al. (2016)

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Table 4.4 Summary of current Chlorophyceae classification Phylogenetic lineages or orders

Example genera

Volvocales

Chlamydomonas (Fig. 4.54a–d), Chloromonas (Fig. 4.54e, f), Dunaliella (Fig. 4.54g, h), Eudorina, Gonium (Fig. 4.55d), Haematococcus (Fig. 4.54i–k), Lobochlamys, Pandorina (Fig. 4.55e, f), Pleodorina (Fig. 4.55g), Stephanosphaera (Fig. 4.55a–c),Volvox (Fig. 4.55h–j)

Treubarinia

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Bulbochaete, Oedocladium, Oedogonium (Fig. 4.59)

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Chaetopeltis, Floydiella, Hormotilopsis, Koshicola (Fig. 4.62), Pseudulvella

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Aphanochaete, Caespitella, Chaetophora (Fig. 4.61a, b), Draparnaldia (Fig. 4.61d, e), Fritschiella, Schizomeris, Stigeoclonium (Fig. 4.61c), Uronema (Fig. 4.61f–h),

found in the Cladophorales, Blastophysa, and some members of the Ulotrichales (e.g., Urospora and Acrosiphonia). The fourth type is the siphonous thallus organization, which is characterized by thalli consisting of a single giant tubular cell. It is present in the orders Bryopsidales and Dasycladales. In most species, the siphonous cells contain thousands of nuclei, but in several species of Dasycladales,

the siphonous thallus remains uninucleate throughout much of their life cycle with a giant diploid nucleus that only divides at the onset of reproduction (Berger and Kaever 1992). Siphonous cells typically exhibit cytoplasmic streaming, transporting organelles, nutrients, and transcripts across the thallus (Menzel 1994, 1987; Mine et al. 2005). Some siphonous species form large seaweeds with

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thalli differentiated into distinct structures, including rhizoids, stolons, and blades. The giant cells of siphonocladous and siphonous species are characterized by several cytological specializations such as unique mechanisms of cell differentiation, cell division, and wounding response (Menzel 1988; La Claire 1992; Kim et al. 2001; Mine et al. 2008). Molecular phylogenetic studies, initially based on a single gene (mostly 18S) (Watanabe and Nakayama 2007) or several genes (Cocquyt et al. 2010), and more recently based on chloroplast genomic (Leliaert and Lopez-Bautista 2015; Sun et al. 2016; Turmel et al. 2016, 2017) and nuclear transcriptomic data (Del Cortona et al. 2020; Hou et al. 2022), have largely clarified the relationships among the Ulvophyceae, and enabled to elucidate the origin and evolution of the group (Fig. 4.63). The Ulvophyceae likely diversified about 650–750 million years ago, in the late Tonian and Cryogenian periods, an interval marked by two global glaciations in which ice caps reached the equator. The ancestors of green seaweeds may have survived these extreme climatic conditions in isolated refuges at the bottom of shallow seas. Isolation of millions of years may have resulted in the independent evolution of different ulvophyte groups. After these glaciations, temperatures rose in the Ediacaran, releasing and opening up the sea floor, which likely enabled green seaweeds to disperse and diversify. An increased supply of nutrients and biotic

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interactions such as grazing by animals probably triggered the evolution towards macroscopic growth in the different ulvophyte groups through different strategies, such as multicellularity and the evolution of siphonocladous and siphonous forms.

4.2.11.1 Orders Ulvales and Ulotrichales The orders Ulvales and Ulotrichales include a wide diversity of multicellular forms, including branched or unbranched filaments, blades and tubular forms (Fig. 4.64), as well as some unicellular and sarcinoid members. The group is predominantly marine, but several transitions to freshwater or terrestrial habitats have occurred independently. The separation of the orders Ulvales and Ulotrichales (not supported by molecular phylogenetic data) was based on life history features: the Ulvales have a life cycle involving isomorphic alternations of multicellular stages (Fig. 4.40a), while in the Ulotrichales the diploid, spore-producing stage is a small, thick-walled unicell that is attached to the substrate by a stalk (Codiolum stage) (Fig. 4.39b). Under certain conditions, such as high nutrient, temperature and irradiance and low salinity, Ulva species can form free-floating masses, known as green tides, which are a growing worldwide phenomenon (Smetacek and Zingone 2013). Several species of Ulvales (including Ulva and Monostroma) are cultivated and used by humans (Mantri et al.

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Fig. 4.63 Hypothesis for the evolution of multicellularity and macroscopic growth, and transition to benthic marine habitats in the Ulvophyceae. The topology of the tree is based on Del Cortona et al. (2020). The divergence times estimated by Hou et al. (2022) are even older and indicate a Meso-Neoproterozoic origin of the Ulvophyceae

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Fig. 4.64 Diversity in the Ulvales and Ulotrichales: a–d Ulva (Ulvales) forming blade-like (a) or tubular (b, c) thalli; d In coastal environments affected by nutrient pollution Ulva species can form free-floating blooms, known as green tides; e Ulvella tongshanensis (Ulvales), characterized by parenchymatous thalli with filamentous branches, from Zhu et al. (2015); f Ulothrix sp. (Ulotrichales), characterized by unbranched filaments with parietal chloroplast, courtesy of Chris Carter

2020). Ulva is also used as a fertilizer to improve the soil quality, and in pharmacology where several species have been shown to have antimicrobial, antiviral, or other types of bioactive compounds. Several species of Ulva are used as food, especially in Asia where it is known as “aonori”. Ulva is a good source of proteins, essential amino acids, vitamins, and minerals Ca, Mg.

4.2.11.2 Order Cladophorales The order Cladophorales (including Siphonocladales) is a predominantly marine order, but several species also occur in freshwater habitats. Under conditions of high temperature or high nutrient concentrations some species may form blooms in marine or freshwater environments. For example, in shallow lakes, Pithophora species may form thick, free-floating mats causing economic and ecological problems.

The order includes one of the most commonly encountered freshwater and marine genus, the branching filamentous Cladophora (Fig. 4.65a). Species in the order are generally macroscopic, and have a siphonocladous thallus architecture, which means that the multicellular plants are composed of multinucleate cells. Thallus morphology is very diverse, ranging from unbranched or branched filaments to blade-like or giant-celled thalli (Fig. 4.65) with unique cytological traits and modes of cell division (Leliaert et al. 2007; Mine et al. 2008). Cells range in size from a few µm to several cm, and have a large central vacuole surrounded by a thin layer of cytoplasm containing numerous nuclei and chloroplasts. Chloroplasts are often interconnected by delicate strands forming a parietal network or a more or less continuous layer. The multinucleate cells present regularly spaced nuclei in a stationary cytoplasm and arrays of internuclear microtubules which define regular

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Fig. 4.65 Diversity of the Cladophorales: a Cladophora sp., characterized by branched filaments; b Chaetomorpha ligustica forming unbranched filaments; c Boodlea sp. forming net-like thalli; d Anadyomene sp. characterized by blade-like thalli; e Segregative cell division in Struvea sp.; f Chloroplasts, each with a pyrenoid, interconnected forming a parietal network; g Dictyosphaeria cavernosa, characterized by parenchymatous thalli formed through segregative cell division; h Valonia macrophysa, characterized by large vesicular cells, from De Clerck et al. (2005)

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cytoplasmic domains, one for each nucleus (McNaughton and Goff 1990). This is in contrast with cells in the Bryopsidales and Dasycladales where the cytoplasm exhibits vigorous streaming. Nuclear division is synchronous or circumscribed to discrete mitosis patches (Hori and Enomoto 1978; Staves and La Claire 1985; Motomura 1996; Okuda et al. 1997). Siphonocladous thallus organization is also found in the sister clade of Cladophorales, including the marine endophytic alga Blastophysa, where the nuclei divide in regular mitotic waves (Sears 1967). Marine tropical members are morphologically extremely diverse, comprising highly specialized forms and giant cells with unique cytomorphological traits and modes of cell division (e.g., Valonia, Boergesenia, Dictyosphaeria). The life cycle in sexually reproducing taxa is isomorphic diplobiontic. The intensive grazing pressure by fish and sea urchins, which is typical in tropical seashores, has led to the evolution of a special mode of wounding reaction in Cladophorales. After mechanical damage, the giant cells rapidly contract and separate their cytoplasm into numerous spherical protoplasts, which later secrete new cell walls and grow into new mature plants. The whole process takes place in a couple of seconds. Some members of the Cladophorales (e.g., Siphonocladus, Struvea, and Dictyosphaeria) are characterized by a specialized organized mode of cell division, termed segregative cell division, which bears a close resemblance to this wounding reaction: during cell division, the whole protoplasm simultaneously cleaves into spherical portions, which later expand and develop into new cells. In some species, these cells remain into the mother cell, expand, and form new vegetative branches (Fig. 4.65e). In other species, the segregated cells are released from the degenerated mother cell, settle and form new thalli. Segregative cell division is fundamentally different from other cell division types in the green algae, both at the macroscopic and ultrastructural level. The cytoplasmic reorganization of the multinucleate cell during the cell division involves a rearrangement (disassembly and reassembly) of cortical microtubules and the actin cytoskeleton. The protoplasmic contraction itself is mediated by actin-myosin interactions.

4.2.11.3 Order Bryopsidales The order Bryopsidales (also known as Caulerpales, Codiales, or Siphonales) are characterized by siphonous thalli, which means that the entire plant consists of a single giant tubular cell, containing thousands to millions of nuclei. The cytoplasm exhibits vigorous streaming, enabling transportation of transcripts across the plant. Bryopsidales range in morphology from simple, branched siphons (e.g., Bryopsis) to more complex, differentiated thalli (e.g., Codium, Halimeda, Udotea, Caulerpa) (Fig. 4.66). Sexual life cycle

B. Büdel and T. Friedl

is diplohaplontic with isomorphic or heteromorphic generations. Some species (e.g., Caulerpa spp.) are holocarpic, which means that the whole contents of the vegetative thallus divide up into reproductive cells. In other species (e.g., Bryopsis spp.), gametes or zoospores are produced in a part of the siphon separated from the rest of the thallus. The freshwater genus Dichotomosiphon is unique in being oogamous. The order is almost exclusively marine, with only one genus (Dichotomosiphon) occurring in freshwater habitats. Several species form key components of tropical marine coastal ecosystems, where they are among the major primary producers on coral reefs, in lagoons, and seagrass beds. The thallus surface of several species is calcified, and some are important contributors to coral reef structure. Some species in the genera Codium and Caulerpa are notorious for their invasive nature (Verlaque et al. 2003; Matheson et al. 2014). Several siphonous species form large and complex seaweeds that exhibit morphological differentiation into structures that resemble the roots, stems, and leaves of land plants and even have similar functions. The evolution of siphonocladous and siphonous architectures coincided with several cytological and cytoskeletal specializations such as unique mechanisms of wounding response. In addition to facilitating transport of transcripts as mentioned above, the evolution of cytoplasmic streaming in siphonous algae also allowed transport of nutrients and organelles throughout the siphonous algal body. In combination with morphological changes, this allows nutrient uptake from marine sediments and chloroplast migration to optimize photosynthesis and avoid herbivory by micrograzers. Such innovations have most likely had selective advantages and contribute to the ecological dominance of siphonous algae in tropical and warm-temperate coastal ecosystems. Molecular phylogenomic analysis indicates that the Bryopsidales are related to the Chlorophyceae (Del Cortona et al. 2020; Hou et al. 2022), which implies that the siphonous architecture found in the Bryopsidales and Dasycladales has evolved independently.

4.2.11.4 Order Dasycladales The order Dasycladales are marine tropical algae characterized by radially symmetrical thalli that are often encrusted with calcium carbonate (Fig. 4.67). The Dasycladales have a rich fossil record and were much more diverse historically than they are today. Fossil remains date back to the Cambrian Period (540–488 my) and suggests that non-calcified Dasycladales were most diverse during the Ordovician and Silurian periods and declined in favor of calcified representatives after the Early Devonian (±400 my), perhaps as a result of selection for resistance to herbivory. Thallus organization is siphonous; however, unlike Bryopsidales,

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Fig. 4.66 Bryopsidales diversity: a, b Bryopsis spp., relatively small feather-like siphonous thallus; c Chlorodesmis fastigiata. Simple, branched siphonous thallus; d Caulerpa racemosa, siphonous thallus differentiated into horizontal stolon-like runners attached by rhizoids, and erect photosynthetic fronds; e Halimeda distorta, complex, multiaxial siphonous thallus composed of articulated series of calcified segments connected by flexible joints; f Codium sp., complex siphonous thallus of cylindrical branches composed of a medullar region of entangled siphons and a cortical layer of inflated utricles, courtesy of John Turnbull

many Dasycladales remain uninucleate throughout much of their life cycle with a giant diploid nucleus that only divides at the onset of reproduction. The group includes the model organism, mermaid’s wine glass (Acetabularia) which displays an unusual life cycle, the only diploid nucleus being the zygote (Fig. 4.68). Following its formation, the zygote develops an upright axis with whorls of branches. The diploid zygote nucleus increases to form a giant primary nucleus, which lies in the rhizoidal basal part of the plant. When reproductive the plant forms an umbel of laterally fused gametangia. The zygote then drops in size and divides meiotically, followed by numerous mitoses producing up to 20,000 tiny secondary

nuclei, which migrate upwards to the reproductive umbel. In the gametangia, each nucleus surrounds itself by a protoplast and forms a thick rigid wall. At that stage, the plant dies and the gametangia are released. After a period of dormancy, each gametangium releases numerous male or female gametes that fuse and will form a planozygote. Eventually, zygotes settle and will produce new plants.

4.2.11.5 Order Trentepohliales The order Trentepohliales is an entirely terrestrial order growing on humid soil, rocks, or trees. The order was originally omitted from the Ulvophyceae based on atypical characteristics such as a multi-layered structure (MLS) in the

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Fig. 4.67 Dasycladales: a Bornetella oligospora, siphonous thalli composed of a primary axis and whorls of branches with the terminal siphons coalescing to form a closed cortex; b Parvocaulis parvulus, siphonous thalli consisting of a primary axis producing whorls of lateral branches of 2 types: colorless branched hairs (not seen in photograph) and clavate gametophores forming a cap, courtesy of Eric Coppejans

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Isogametes of opposite sexes fertilization Whorls of sterile hairs Zygote (2n) Formation of umbrella-like whorl of gametangia Small haploid nuclei transported to other parts of the thallus by cytoplasmic streaming

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Fig. 4.68 Life cycle of Acetabularia (Dasycladales), based on van den Hoek et al. (1995) and Berger and Liddle (2003)

flagellar root system, phragmoplast-like cytokinesis, the presence of plasmodesmata between vegetative cells, and a unique type of sporangial reproduction implying a relationship with charophyte green algae (López-Bautista et al. 2002). However, molecular data firmly established an alliance with the orders Cladophorales, Bryopsidales and Dasycladales, either pointing towards parallel evolution of

the streptophyte-like ultrastructural features or indicating that some of these characteristics (e.g., a MLS) may represent an ancestral condition in the green lineage; either way it is interesting that a phragmoplast-like cytokinesis has evolved in Trentepohliales and land plants, both found in terrestrial environments (Leliaert et al. 2012).

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Mesostigma, order Mesostigmatales Mesostigma is flagellated unicellular with disk-shaped cells that contains a flattened chloroplast with two pyrenoids in the enlarged portion at the rims of the cell (Fig. 4.70a, b). Motility is conceived by two flagella that are laterally inserted in a deep flagellar pit (Fig. 4.70b). The cell body is

tracheophytes Mosses Liverworts Hornworts

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4.2.12.1 Class Mesostigmatophyceae The class Mesostigmatophyceae comprises three lineages wherein each is represented by a single genus, i.e., the flagellate Mesostigma, the sarcinoid cell packet forming Chlorokybus, and the unicellular coccoid Spirotaenia. In phylogenomic analyses (e.g., Lemieux et al. 2007; Irisarri et al. 2021), the three lineages form a well-supported monophyletic clade which is at the base of all Streptophyta (Fig. 4.28). Therefore, the clade comprises the most ancestral algal relatives plants. Here the three lineages will be treated as a single class, Mesostigmatophyceae, but each lineage may in fact represent a class by its own (e.g., Chlorokybophyceae; Irisarri et al. 2021).

streptophyte algae grade

The streptophyte algae grade is an assemblage of green algal lineages that bears important information on terrestrialization, i.e., the transition from water to land or change from aquatic to a terrestrial lifestyle. It marks a major event in the evolution and diversification of the “new cryptogams” (as treated in this book) and multicellular land plants, the embryophytes (Fig. 4.69). However, genome sequence analyses demonstrate that several of the evolutionary novelties required to make the terrestrialization event of the Streptophyta and its further development so successful have evolved in the streptophyte algae grade which predominantly has an aquatic lifestyle (e.g., Nishiyama et al. 2018; Cheng et al. 2019; Delaux and Schornack 2021). While several distantly related algal lineages, e.g., Trebouxiophyceae in the Chlorophyta and the Xanthophyceae of Stramenopiles, evolved to occupy terrestrial environments, only one represents the land plant ancestor. The grade means a paraphyletic assemblage of green algal lineages of the Streptophyta that precedes the multicellular land plants, the embryophytes (Fig. 4.69). At least seven monophyletic groups or lineages are currently recognized in the streptophyte algae grade: Mesostigma, Chlorokybus, Spirotaenia which may be united into a single class, Mesostigmatophyceae (Cheng et al. 2019), the classes Klebsormidiophyceae, Charophyceae, Coleochaetophyceae, and Zygnematophyceae. Only Chlorokybus, Klebsormidiophyceae, and a few members of the Zygnematophyceae are known from terrestrial habitats. Another independent lineage

of the streptophyte algae grade may represent the terrestrial Streptofilum but its exact phylogenetic position is yet still unresolved (Mikhailyuk et al. 2018). There may be even more lineages of the streptophyte algae grade to be discovered in the future (Irisarri et al. 2021). Within the Streptophyta, the Zygnematophyceae represent the closest relatives to land plants as revealed in robust phylogenies from phylogenomic analyses (e.g., Wickett et al. 2014; Cheng et al. 2019; One Thousand Plant Transcriptomes Initiative 2019).

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4.2.12 Phylum Streptophyta—The Streptophyte Algae Grade

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Fig. 4.69 Phylogenetic lineages and classes of the streptophyte algae grade within the Streptophyta; the transition to a terrestrial environment initial to the evolution and diversification of land plant life (terrestrialization) occurred in the common ancestor of embryophytes; the dotted line marks the boundary between the “new cryptogams” as treated in this book (below the line) and the “plants” (above the line); topology of the phylogeny is a consensus from phylogenomic analyses presented in Puttick et al. (2018), Cheng et al. (2019), and Irisarri et al. (2021)

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Fig. 4.70 a Mesostigma viride a flattened cell of culture strain SAG 50-1; note enlarged portions of the chloroplast (ch); b schematic interpretative drawings of swimming cells at the front (broad), side (edge) and upper surface views; note deep flagellar pit, lateral insertion of the two flagella and characteristic curvature of the cell (from Manton and Ettl 1965); c cross section through the right half of the cell viewed down from the flagellar tip, the pyrenoid (py) is in an enlarged portion of the chloroplast; contractile vacuole (cv), nucleus (nu); note prominent body scales on the cell surface (transmission electron micrograph; Melkonian 1989); a courtesy of Tatyana Darienko, with permission of the SAG Culture Collection of Algae; b with permission from Wiley; c with permission of Elsevier

covered by prominent scales on the surface (Fig. 4.70c). There are thin and small additional pyrenoids (“superficial pyrenoids”) which are located outside of the chloroplast, similar to the pseudopyrenoids in Chlorokybus. Mesostigma reproduces asexually by binary fission at the flagellate stage. Sexual reproduction is not known. M. viride is found in the benthos of small, shallow ponds (Wang et al. 2020a). Ultrastructural analyses already suggested that the earliest divergence within the Streptophyta was a scaly, biflagellate, unicellular green alga like Mesostigma (Rogers et al. 1981). Independent direct evidence for this assumption came then from molecular phylogenetic (Melkonian et al. 1995; Bhattacharya et al. 1998) and phylogenomic analyses (Lemieux et al. 2007; Wickett et al. 2014). Flagellate reproductive cells (zoospores, gametes) in streptophyte green algae have a unilateral broad microtubular flagellar root which is associated anteriorly with a multi-layered structure (MLS) and are covered with a layer of square-shaped scales on their cell surface (Manton and Ettl 1965; Marin and Melkonian 1999). Independent direct evidence for this assumption came then from molecular phylogenetic (Melkonian et al. 1995; Bhattacharya et al. 1998) and phylogenomic analyses (Lemieux et al. 2007; Wickett et al. 2014; Wang et al. 2020a). Chlorokybus, order Chlorokybales The cells of Chlorokybus species have a cup-shaped chloroplast which contains a prominent pyrenoid that is enclosed by parallelly arranged starch grains (Fig. 4.71e, g).

The pyrenoid matrix exhibits a striation from thylakoids that traverse the matrix in a regular pattern (Fig. 4.71g, i; Gärtner and Ingolic 1989). There is a peculiar structure of Chlorokybus cells which is similar to that in Mesostigma but otherwise not found in green algae, i.e., the pseudopyrenoid, which is a kind of “satellite” pyrenoid located outside of the chloroplast at the inner surface of the cell (Fig. 4.71e, f). The vegetative thalli are usually two- to four-celled sarcinoid packets surrounded by a thick mucilage layer (Fig. 4.71). The term sarcinoid describes a thallus that consists of a three-dimensional cluster of cells. At Chlorokybus, the cell packets arise from a cell division that is not followed by deposition of wall material all around each daughter cell (as it is the case in the reproduction by autospores, see Fig. 4.71g), but a septum is formed by an increasing cleavage furrow of the plasma membrane which is independent of the formation of a cell wall (Fig. 4.71g; Rogers et al. 1980; Lokhorst et al. 1988). Therefore, the type of cell division in Chlorokybus is different from that of unicellular members of the Chlorophyta but similar to that of tissue-forming bryophytes and tracheophytes (see Chap. 7). Asexual reproduction is, besides cell division to form cell packets, achieved by zoospores and autospores formed within the parental cell wall (Rieth 1972). A single zoospore per cell is formed. Then gradual disintegration or gelatinization of the cell packets happens. The zoospores are covered by scales like in other streptophyte algae zoospores (Fig. 4.71i). The zoospores have two flagella which emerge laterally from the cell body (Rieth 1972; Fig. 4.71h). The life cycle is haplontic. A summary of the life cycle stages is presented by Irisarri et al. (2021).

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Spirotaenia The cells of Spirotaenia are unicellular coccoid, straight, or slightly curved oblong cylinders, have rounded cell poles and are up to about 250 lm long. They contain a single ribbon-like chloroplast with numerous pyrenoids. The chloroplast ribbon is spirally twisted to the left (Fig. 4.72). Therefore, cells of Spirotaenia superficially resemble vegetative cells of genera of Zygnematophyceae, i.e., the filamentous Spirogyra and the unicellular Spirogloea which, however, has much shorter cells (Zygnematophyceae; Fig. 4.90a). Asexual reproduction is by transverse cell division (Fig. 4.72a). Therefore, in field samples, often two cells remain near each other after division within a common mucilaginous sheath.

At sexual reproduction of Spirotaenia condensata, two individuals that are unequal in length and belong to complementary mating types, lay parallel closely associated with each other in a common mucilaginous sheath. After nuclear and protoplast division, in each cell the two protoplasts round up to form non-motile gametes (Fig. 4.73a). A pair of mated cells forms four gametes which becomes released after the cell wall gelatinizes. The non-sister gametes fuse leaving a pair of zygotes (Fig. 4.73b). The thick-walled zygotes are the dormant resting stages, named zygospores (Hoshaw and Hilton 1966). At zygospore germination following the rupture of the zygospore wall, four small cells emerge (Fig. 4.73c). They remain in a common mucilaginous sheath where each cell contains the typical spiral ribbon-like chloroplast before release (Hoshaw and Hilton 1966).

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Fig. 4.72 Morphology of Spirotaenia condensata; a elongated cell slightly curved, the ribbon-shaped chloroplast filled with starch grains; in the left two cells the nucleus is obvious, the cell at the right is in division; culture strain MZCH SVCK 312; b schematic drawing of two cells at different optical sections illustrating the spirally twisted chloroplast with numerous pyrenoids; scale, 50 lm; courtesy of Tatyana Darienko (a), from Prescott et al. (1972), with permission of NYBG Press (b)

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Fig. 4.73 Sexual reproduction in Spirotaenia condensata (Hoshaw and Hilton 1966); a paired cells of unequal size; in the smaller cell, the protoplast already divided and division products formed two non-motile gametes which become released upon gelatinization of the cell wall; b two gametes fused into a zygote (left), the other two gametes are left still unfused; c a pair of zygospores with peculiar wall ornamentation resulting from the fusion of the four gametes; d germination of a zygospore into four young cells inside a common mucilaginous sheath leaving the zygospore wall empty; young cells with spirally twisted chloroplasts; scale, 50 lm; images from Hoshaw and Hilton (1966); JSTOR, http://www. jstor.com/stable/40022376

In traditional systematics and identification keys, the genus Spirotaenia has been considered a member of saccoderm (= without cell wall constrictions) Desmidiales of the class Zygnematophyceae, next to species of Mesotaenium (Fig. 4.90b). Like desmids, Spirotaenia species live in slightly acidic nutrient-poor aquatic environments such as peat bogs and often occur intermingled with them. Sexual reproduction by conjugation is a characteristic feature of the class Zygnematophyceae (see Sect. 4.2.12.5). However, Spirotaenia condensata and some other species

of the genus studied so far differ from all the conjugating green algae in producing no conjugation tube or vesicle. Instead, there are two gametes per gametangium cell which are released through gelatinizing of gametangial cell walls prior to gamete fusion (Coesel et al. 2017). It is likely that these differences from the conjugation in Zygnematophyceae already mark the deep phylogenetic divergence between Spirotaenia and the Zygnematophyceae (Gontcharov 2008). However, besides the few species of Spirotaenia so far investigated for their phylogenetic position

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(Gontcharov and Melkonian 2004; Wickett et al. 2014), there are about 20 more species names of Spirotaenia taxonomically accepted at present (Guiry and Guiry 2023). Therefore, it cannot be excluded yet that Spirotaenia species are of different origins with some other species being in fact members of the Zygnematophyceae (Coesel et al. 2017).

4.2.12.2 Class Klebsormidiophyceae, Order Klebsormidiales Klebsormidiophyceae is among the early branching lineages (classes) of the Streptophyta and out of those it is the most species rich. For the genus Klebsormidium (Fig. 4.74a– c) >20 species are currently taxonomically accepted which are distributed on several phylogenetic clades within the genus (Rindi et al. 2011; Škaloud and Rindi 2013), but the species phylogeny is still a matter of current research. The Klebsormidiophyceae comprises at present six genera, i.e., Klebsormidium and its closest relative Interfilum (Fig. 4.74 d), Hormidiella (Fig. 4.74f) and Streptosarcina (Fig. 4.74e) which together form a sister group with the Klebsormidium/Interfilum lineage (Mikhailyuk et al. 2018). Finally, there is Entransia which has been included into the class (Sluiman et al. 2008), but recent phylogenetic analyses revealed no significance to support its monophyletic origin together with the other five genera (Mikhailyuk et al. 2018). Most Klebsormidiophyceae are common cosmopolitan species of terrestrial habitats, e.g., in soil or on the surface of artificial hard substrates in urban environments, all over the world from polar to desert regions (Mikhailyuk et al. 2008; Rindi et al. 2008). The worldwide distribution of members of Klebsormidiophyceae can be explained by their ability to cope with high fluctuations in temperature, water availability, pH, and solar radiation (Glaser et al. 2017). Members of Klebsormidiophyceae have developed various physiological, biochemical, and ultrastructural mechanisms to withstand numerous environmental stressors. They include photoprotection (e.g., formation of UV-sunscreen compounds), photochemical quenching, high osmotic values to avoid water loss, and in some groups the flexibility of forming secondary cell walls to maintain turgor pressure in water-limited situations (Karsten and Holzinger 2012; Glaser et al. 2017). Only Entransia and likely a few species of Klebsormidium are known from non-water limited habitats. As an important component of BSC (biological soil crusts) which occur mainly in dry lands or disturbed environments, Klebsormidiophyceae contribute significantly to primary production, carbon and nitrogen cycling, soil stabilization, and water retention (Holzinger and Pichrtová 2016; Pierangelini et al. 2019). Klebsormidiophyceae are mostly filamentous, i.e., form uniseriate unbranched filament without a holdfast (e.g., Klebsormidium, Fig. 4.74a–c) which often can easily

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disintegrate (e.g., Hormidiella, Fig. 4.74f). There are also unicellular forms (e.g., Interfilum, Fig. 4.74d) which tend to form short filaments or cell packages (e.g., Streptosarcina, Fig. 4.74e). Typically, Klebsormidiophyceae exhibit a plate-like chloroplast which is covering just about half of the cell and appressed to the cell wall. It contains a single or several pyrenoids with the pyrenoid matrix encircled by starch (Interfilum) or loosely surrounded by starch grains (Klebsormidium). Often the nucleus, which is within a pocket of cytoplasm, is located directly opposite the pyrenoid which is in the part of the chloroplast that covers the cell wall (Fig. 4.74a, c). Reproduction takes place simply and is most efficient by fragmentation of the thallus (Fig. 4.74b, f). In addition, biflagellated ovoid zoospores devoid of any scales and without an eyespot may be formed singly per cell from which they escape through a pore (Lokhorst 1996). That zoospore release through a pore is regarded as a kind of specialization that distinguishes the Klebsormidiophyceae from other basal lineages of the streptophyte algae grade (Graham et al. 2016).

4.2.12.3 Class Charophyceae, Order Charales The class Charophyceae comprises a single order, Charales. Its members are very distinct from other streptophyte green algae by their morphologically complex and large thallus (plant body) which is several centimeters in size. Fundamentally, Charales form branched filaments. The axis of a thallus is attached to the substrate by branched rhizoids which grow downward (positively geotropic). The thallus is with apical growth and differentiated into nodes and internodes (Figs. 4.75, 4.76 and 4.77). Charales grow in still, clear freshwaters (ditches, pools, lakes) where they can form extensive underwater vegetations (Fig. 4.75). Some species can form extensive meadows even in fairly deep freshwaters. They are particularly abundant in hard water (pH > 7) with a few species occurring in brackish water. For example, some Charales species dominate shallow, sheltered, soft-bottomed areas of the Baltic Sea (Urbaniak and Gąbka 2014). The cell walls often bear surface layers of calcium carbonate in the form of calcite which results from the precipitation of calcium carbonate (CaCO3) in water high in Ca2+, hence the common name of the Charales, stoneworts. Calcification gives some forms a white or pale-green appearance (Fig. 4.75). Vegetative structures At the apex of the thallus main axis, there is a single specialized meristematic cell (Fig. 4.76). New daughter cells appear from its lower surface only, thereby extending the filament in length. Therefore, the Charalean meristematic cell has just one face. In contrast, the apical meristematic cells in bryophytes have three or four faces that cut off new cells and thus generate tissues (Chap. 7). The immediate cell

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Fig. 4.74 General morphology of Klebsormidiales (Klebsormidiophyceae); py, pyrenoid; asterisk, cells in a view that shows the chloroplast covers only about half of the cell; a Klebsormidium sp. from a culture freshly isolated from soil of a plant pot, Göttingen, Germany; cells elongated (about 2 times longer than wide), pyrenoids surrounded by numerous starch grains; arrow, position of the nucleus in a cytoplasm pocket directly opposite the pyrenoid; b K. crenulatum from a three months old culture (strain SAG 37.86); filaments with thick corrugated cell walls and frequent production of H-shaped wall pieces (arrowheads); chloroplasts shaped like an open band; cells about as long as wide; c Klebsormidium sp., filaments readily fragmenting into unicells and short fragments as a means of fast reproduction in culture (strain SAG 2215, isolated from a roof tile in Göttingen, Germany); d Interfilum paradoxum, unicellular form in culture (strain SAG 4.86); arrowheads, daughter cells still connected with each other after division (forming pseudofilaments) by remains of the parental wall; e Streptosarcina arenaria, sarcinoid thallus form (consisting of cell packages) in culture (strain SAG 2562); note plate-shaped chloroplast with a single pyrenoid (py) surrounded by several starch grains; f Hormidiella parvula, short filaments that are fragmenting and empty cell walls (likely remains of sporangial walls after zoospore release), culture strain SAG 2558; courtesy of Vanessa B.A. Spieß (a), Maike Lorenz (c), and Tatyana Darienko (b, e, f), with permission of SAG culture collection of algae; d, original Thomas Friedl

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Fig. 4.75 Species of Chara forming underwater meadows in shallow freshwaters; a Chara sp. at a pond formed by groundwater onto sediment sufficiently rich in organic material to be anoxic close to the surface (sulfur precipitation), at Eschenlohe, Germany; note differentiation into long internodes and nodes with branches; b C. vulgaris close to the spring of the summer-cold creek Fischa, Haschendorf, Austria (underwater photograph); c sterile specimens of C. tomentosa, reddish-colored by c-carotene (Schagerl and Pichler 2000), in an endangered habitat, open water of Lake Neusiedl at Illmitz, Austria, with reed zone in the background; original Friedl (a) and courtesy of Michael Schagerl (b, c)

that cuts off the apical meristematic cell divides mitotically. Then the upper cell divides again laterally which by further divisions will form the complex node (Fig. 4.76). The lower cell develops into a giant several centimeters long internodal cell without further division mediated by a large internal vacuole. From the complex node, the lateral branches arise by a series of asymmetric divisions (Figs. 4.76 and 4.77). The branches have the same kind of alternating nodal and internodal cells as the main axis. The branches in turn produce smaller branchlets (Fig. 4.79). Unlike the apical cell of

the main axis, which can continue to divide indeterminate, the branch apices cease division after a determined number of cells have been produced. In Chara, the nodal cells at the base of the branches generate multiple rows of filaments that grow up and down over the internodal cell surfaces. This forms a complex sheath of cortical cells that envelopes each internode (cortication, Fig. 4.77). The corticating filaments originating from two neighboring nodes meet in the middle of the internodal cell (Fig. 4.79). A cortication is present only in

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Fig. 4.76 Apical meristem of Chara fibrosa (transmission electron micrographs); arrow, apical meristematic cell; nd, nodal cell; in, internodal cell; co, cells of filaments forming the cortication on the surface of the internodal cell; a cross section through a thallus with young lateral branches still not differentiated into nodes and internodes just below the apical meristem; b cross section through a thallus with forming oogonial complexes (asterisks) at the lateral branches below the apical meristem; note internodal cell becoming ensheathed by cortical cells. a, b from Pickett-Heaps (1975), reproduced with permission of Oxford University Press through PLSclear

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Fig. 4.77 Vegetative structures of a thallus of Chara intermedia. Note spirally twisted arrangement of cortical cells (co, cortication) that are with spine cells (sc); a view at a node with lateral branches (upper half) and internode (lower half); at the base of the lateral branches there are stipulodes (st); b details of an internode with cortication. Courtesy of Jacek Urbaniak (Urbaniak and Gąbka 2014)

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a the species-rich genus Chara while it is lacking in other genera, e.g., Nitella, and Tolypella (Fig. 4.78). The internodal cell contains numerous nuclei which resulted from the replication of the single haploid internodal cell’s original nucleus. However, their haploid state was found instable, with most nuclei being polypoid due to endoduplication or gene amplification (Michaux-Ferrière and Soulié-Märsche 1987). In the cytoplasm nearest the central vacuole

b cytoplasmic streaming can be seen which results from actin microfibril activity. Within the internodal cell, there is also a peripheral layer of nonmobile cytoplasm which contains numerous small chloroplasts generated by multiple fission (Fig. 4.78c). They have a discoid shape and are arranged in rows. During cell division, a phragmoplast (see Fig. 4.38) develops, resulting in the formation of a cross wall with plasmodesmata, i.e., a cell plate. It separates internodal

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Fig. 4.78 General morphology of Nitella and Tolypella; a–c Nitella sp. from the Spessart midlands in Germany (creek Aubach, Partenstein); a habitus of thallus; b view at a node with branches and internode cell; c internodial cell at higher magnification exhibiting numerous small chloroplasts, note epiphytic diatoms attached to the cell surface; d Tolypella nidifica, dense branchlets forming whorls. a–c courtesy of Burkhard Büdel; d courtesy of Jacek Urbaniak (Urbaniak and Gąbka 2014)

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from nodal cells and with its large pores it facilitates the passage of materials throughout the vertical axis of the thallus. Sexual reproduction Sexual reproduction is oogamous with motile biflagellate spermatozoids (Fig. 4.82c) released from the antheridium swimming to the non-motile oogonium (Figs. 4.78, 4.79 and 4.82b) to fertilize the ovum (egg cell). The resulting zygote, called the oospore (Fig. 4.83), is the only diploid stage in the life cycle; it functions as a resting stage which sinks onto the sediment of the habitat and germinates only after a period of dormancy with meiosis. Thus, the Charales life cycle is haplontic (Fig. 2.9). There are monoecious species where both the spherical antheridia and flask-shaped oogonia are formed together at the same thallus, and dioecious species with only one type of reproductive organ on separate thalli (Fig. 4.79). In monoecious species, the antheridia and oogonia are paired and lie on the adaxial sides of the lateral branches (branchlets, Fig. 4.79). The male and female reproductive structures are probably the most conspicuous sexual structures of all green algae. The antheridium is a globose (spherical) structure that is composed of eight peripheral shield cells (Figs. 4.79 and 4.80a). The antheridium is bright orange at maturity due to carotenoid droplets generated by the peripheral shield cells within an outer layer of the cells. The shield cells are of a peculiar form and arranged to give rise to distinct patterns which are used for identification of some species (Fig. 4.80a). The development of the globose antheridium starts from haploid nodal cells that occur at lateral branches (Fig. 4.80); it is illustrated in Fig. 4.80. The shield cells are connected to the central part of the spherical antheridium by a manubrium (stalk cell). Attached to the manubrium are two specialized cells, primary and secondary capitulum (Fig. 4.80g). From the latter arise, the spermatogenous filaments (Figs. 4.80g and 4.81b) by repeated division. Each single cell of a spermatogenous filament produces a single asymmetric biflagellated spermatozoid. The mature spermatozoid has a characteristic spiral shape with the flagella inserted subapically, and it has three different regions within the body (Fig. 4.81c). The spermatozoids have a cell covering of scales. The oogonium also develops from a nodal cell at a lateral branch (Fig. 4.82). At maturity, the ovum (egg cell) in Chara is surrounded by five spirally twisted (cortical) sheat cells, each of which ends in a corona cell. The development of the oogonium from a nodal cell and its structures are shown in Fig. 4.82. Narrow fissures between the corona cells allow spermatozoids to penetrate and fertilize the ovum. The resulting zygote is the only diploid part of the life history in Charales. The zygotes develop a thick covering of sporopollenin. Other portions of the zygote wall decay

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leaving the persisting portions of walls of the sheath cells like the threads of a screw (Fig. 4.83). Because the sporopollenin is highly resistant and given favorable depositional conditions, the Charales zygotes are well represented in the fossil record. Fossilized zygote walls are called gyrogonites. Fossils of Charales date back to at least 380 mya. After a period of dormancy, the zygote germinates with meiosis producing a cell with four haploid nuclei, but only one is surviving. The one haploid cell develops into a new haploid thallus by repeated mitotic divisions. Asexual reproduction The Charales can also reproduce vegetatively simply by thallus fragmentation, outgrowth from a node or by multicellular rhizoids which are not differentiated into nodes and internodes. Vegetative propagules that are white spherical or star-shaped structures, called bulbils, develop at the rhizoids or lower nodes. After detachment from the thallus, they germinate with mitosis to develop a new thallus. No asexual zoospores are formed.

4.2.12.4 Class Coleochaetophyceae, Order Coleochaetales The Coleochaetophyceae are small (100) microtubules; note the small R3 root on the left; c, d scaly cell covering; c section perpendicular to the plasma membrane; d face view; from Sluiman (1983) (with permission of Springer Nature publisher)

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Fig. 4.86 Stages of sexual reproduction (development of sexual organs) in Coleochaete pulvinata as shown in the original publication of Friedrich Oltmanns (1898); a filament with a young antheridium (an) at the tip of a branch; b flask-shaped oogonium with long colorless trichogyne (tr) and basal ovum (ov); c zygote (zy) enclosed by outgrowth of neighboring cells forming a protective sheath; e spermocarp after meitotic germinating of the zygote with opening that released the haploid zoospores; modified from Oltmanns (1898)

streptophyte algae grade with an estimated number >3,000 species in some 50 genera (Graham et al. 2016). Zygnematophyceae comprises morphologically simple algae which exhibit basically two types of body forms, i.e., cylindrical cells united permanently into unbranched filaments without holdfast (Fig. 4.88), and non-motile unicells which mostly have a pronounced constriction giving rise to two semicells and often exhibit ornamentations of their cell walls (Fig. 4.91). A very distinctive feature of the Zygnematophyceae is that they completely lack flagellated stages.

During sexual reproduction amoeboid gametes fuse in the characteristic process of conjugation to form zygotes (Figs. 4.89 and 4.94). The amoeboid gametes consist of the protoplasts of the vegetative cells. Union of the two gametes is usually established by a conjugation tube which is continuous from one cell to the other between two cells (Figs. 4.89 and 4.94). The two growth forms gave rise to two orders in the traditional taxonomy, i.e., the Desmidiales and Zygnematales. However, recent transcriptomic analyses revealed five independent origins of the filamentous growth

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Fig. 4.87 Typical habitat of Zygnematales (Spirogyra sp.); green mats of filamentous algae floating in a small slowly running creek; Original Thomas Friedl

within the Zygnematophyceae. It led to the proposal of a new five-order system for the Zygnematophyceae (Hess et al. 2022): Desmidiales, Spirogyrales, Zygnematales, Serritaeniales, and Spirogloeales. Most Zygnematophyceae have an aquatic lifestyle but exhibit a mucilaginous coat (e.g., Fig. 4.90i) that surrounds the cells and may increase desiccation tolerance. Zygnematophyceae is considered the most likely sister group of embryophyte land plants (e.g., Wodniok et al. 2011; Wickett et al. 2014; Cheng et al. 2019; One Thousand Plant Transcriptomes Initiative 2019). Two early diverging Zygnematophyceae, Spirogloea muscicola (Fig. 4.90a) and Mesotaenium endlicherianum (Fig. 4.90b), share the same subaerial habitat with the bryophytes, i.e., earliest-diverging embryophytes. They have genes that increase resistance to biotic and abiotic stresses in land plants (Cheng et al. 2019). Order Zygnematales The Zygnematales are among the most common filamentous freshwater algae. Often, they favor stagnant water bodies where they can form bright-green floating mats (Fig. 4.87) Those mats can even move towards the light source by gliding and curvature of the filaments; they can tolerate high light intensities. Three species-rich genera may be most characteristic for the Zygnematales. Spirogyra features spirally one or two (sometimes even more) twisted chloroplast bands (Fig. 4.88a, b) extending the length of the cell. Mougeotia has a single flat chloroplast plate (Fig. 4.88c, d). In both genera pyrenoids the chloroplasts are with conspicuous and numerous pyrenoids. Mougeotia exhibits a marked chloroplast orientation in response to light, i.e., the chloroplast presents a surface view to the light under low

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intensities whereas it presents an edge view towards high irradiation (Fig. 4.88c, d; Graham et al. 2016). Asexual reproduction is predominant and occurs by fragmentation of the filaments. Zygnema possesses two stellate chloroplasts with a central pyrenoid, oriented along one axis, and with nucleus between them (Fig. 4.88e–g). For sexual reproduction the filaments of the Zygnematales come to lie side by side, and the cells from adjacent filaments push out contiguous conjugation tubes which abut one another and then the two tubes fuse into one (Fig. 4.89). One gamete moves through the conjugation tube and unites with the other gamete inside the parent wall of the other protoplast (gamete). Fusion of both gametes can also happen inside of the conjugation tube. Once the nuclei united, the resulting zygote secretes a tough, resistant cell wall around itself which consists of three layers, called exo-, meso-, and endospore (Fig. 4.89e) The exospore is often sculptured, the mesospore is colored and may contain sporopollenin. The colorless thin endospore contains cellulose and pectin. As dormant stages the zygotes can outlast prolonged periods of environmental stress (e.g., desiccation) before the meiosis and germination takes place. After a resting period of the zygote, meiosis occurs of which three haploid nuclei disintegrate. The life cycle is typically haploid with the zygote being the only diploid cell serving as a resting stage (Fig. 4.89). Order Desmidiales Desmidiales is a unique group of streptophyte green algae, called “desmids, which form unicells that can be solitary, joined end to end in filamentous colonies or united in amorphous colonies (Fig. 4.90). In desmids the cell walls are formed at different times, for which reason their cell wall is said to be composed of two pieces. Desmids commonly have an aquatic lifestyle, but some species which are early diverging in phylogenetic analyses are reported to live in terrestrial habitats (Fig. 4.90a; Cheng et al. 2019). Most desmids live in low-nutrient (oligothrophic) and often slightly acidic freshwaters, i.e., ponds, lakes, peat bogs, and attachedto substrates in streams. Desmidiales cell walls are also perforated with pores, i.e., the cell walls are pierced by many pores (Figs. 4.90b and 4.92), and often highly ornamented (Figs. 4.90f, h and 4.91b). Mucilage is secreted from individual wall pores, often forming a confluent sheath (e.g., Desmidium, Fig. 4.90 i). It may explain the common ability of many desmids to move by mucilage secretion. In addition, there are resistant cell wall polymers in many desmids which may also explain the survival of certain desmids while their habitat falls dry. Placoderm desmids are those members of Desmidiales which have a constriction in the middle of the cell which divides them into symmetrical halves or semicells (Figs. 4.90d–f and 4.91). The

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b Fig. 4.88 General morphology of Zygnematales; a, b Spirogyra sp., filaments of long cylindrical cells, and a single spirally twisted chloroplast

band with many prominent pyrenoids; a culture strain SAG 170.80; b field sample; c, d Mougeotia sp., culture strain SAG 11.96; filaments of long cylindrical cells, one plate-like chloroplast with several conspicuous pyrenoids per cell, in the uppermost second filament the chloroplast presents an edge view (as towards high light intensities); d chloroplast presenting surface view (as under low-light intensities); e–g Zygnema spp., filaments of long cylindrical cells with two stellate chloroplasts, each with a central pyrenoid, and with the nucleus between them (looks like a bridge between chloroplasts), thus, there are two stellate chloroplasts each with a large pyrenoid and the nucleus positioned between both chloroplasts; e Z. cylindricum, culture strain SAG 698-2; f Z. circumcarinatum, culture strain SAG 698–1; g) transmission electron micrograph of a thin section through a young vegetative cell of Zygnema sp.; zentral nucleus (nu) with chloroplasts (ch) on either side with large pyrenoids (py), surrounded by starch grains; large vacuoles (v); cell wall of two distinct layers (w) (from Pickett-Heaps 1975) a–f courtesy of Tatyana Darienko, with permission of the SAG culture collection of algae; g reproduced with permission of Oxford University Press through PLSclear

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Fig. 4.89 The process of conjugation to form zygotes in Spirogyra (Zygnematales), schematic representation; a two filaments of different mating types formed conjugation tubes between each other; b the protoplasts transformed to amoeboid gametes, the gametes from one filament moved to the gamete of the other filament through the conjugation tube; d after fusion of both gametes zygotes are formed in the one filament; d, e zygotes formed after conjugation completed; a–c from Lee (2018), with permission of Cambridge University Press; d, e from Kadlubowska (2009), with permission of Springer Spektrum

constriction may be less (as e.g., in Closterium, Fig. 4.90g) or more pronounced (as in most desmids, e.g., Cosmarium, Micrasterias, Staurastrum, Xanthidium; Figs. 4.90d–f and 4.91b). Thus, the placoderm Desmidiales are unicells composed of two semicells joined by a narrow isthmus (Fig. 4.90d–f). Both semicells are of different age, i.e., the younger cell halve appears less differentiated than the older one (Figs. 4.90d and 4.91a). By contrast, saccoderm desmids have no cell wall constriction (e.g., Mesotaenium, Cylindrocystis; Fig. 4.90a–c). Vegetative cells of desmids display various symmetries, e.g., flattened (biradiate; Micrasterias, Fig. 4.91a) or triradiate when viewed on the cell end (Staurastrum, Fig. 4.91b). Asexual reproduction, cell division. Asexual reproduction is by cell division which follows a pattern unique in all green algae. In highly constricted desmids, cell division starts with mitosis when the nucleus is still centrally located

in the isthmus (Fig. 4.93a–c). During mitosis the semicells move slightly apart. At the end of mitosis, i.e., when the daughter nuclei are re-formed and cytokinesis is almost complete (Fig. 4.93c), the nuclei lie in the forming daughter cells. Semicell expansion starts and at each daughter cell, the chloroplast moves into the new expanding semicell (Fig. 4.93d). Later, the chloroplasts will divide at the isthmus and the nuclei will migrate back to its usual position within the isthmus. Sexual reproduction in Desmidiales is by conjugation like in the Zygnematales (Fig. 4.94). In unicells of Desmidiales, the sexual reproduction commences with pairing of cells. After pairing, the cells move about until they lie flat against one other with one isthmus at right angles to the other (Fig. 4.94b). The paired cells cease moving and secrete mucilage around themselves. Environmental conditions (e.g., low nitrogen supply) and that the cells of both

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Fig. 4.90 Desmidiales morphology, various examples from culture; a–c Saccoderm Desmidiales, without cell wall constrictions; d–i Placoderm Desmidiales, with cell wall constrictions; a Spirogloea muscicola (strain CCAC 0214), note chloroplast band spirally twisted and central nucleus; b Mesotaenium endlicherianum (strain SAG 12.97). Two flat plate-like chloroplasts each with prominent pyrenoid, nucleus centrally between the chloroplasts; c Cylindrocystis crassa (strain SAG 23.97), chloroplast with a somehow stellate structure; d Cosmarium botrytis (strain SAG 136.80), cells with flat ends and deep isthmus, dividing cells with two stages of semicell development in (right), note cell walls ornamented; e Micrasterias radiata (strain SAG 161.80) cell flattened, deeply constricted into two semicells, which possess a deep isthmus and each semicell is divided into three lobes; f Xanthidium cristatum (strain SAG 173.80), semicells ornamented with spines and triradiate symmetry; g Closterium ehrenbergii (strain SAG 134.81); large cell with the wall consisting of two halves of which one is inserted into the other and has no constriction, two chloroplasts with the nucleus in between. Note prominent vacuoles at both ends of the cell; h Pleurotaenium kayei (strain SAG 19.97), long cell with wall highly ornamented with spines; i Desmidium grevileii (strain SAG 637-1a), cylindrical cells are joined end to end in a filamentous colony, the cells excrete mucilage through pores, cell in the lower right is in optical cross section; courtesy of Tatyana Darienko, with permission of SAG culture collection

mating types are present are required for the vegetative cells to become sexual. Chemotaxis seems to be important to attract compatible cells to each other. Both cells form conjugation tubes into which the two protoplasts (gametes) move and fuse to form a zygote (Fig. 4.94). The walls enclosing an escaping gamete define the conjugation tube. A zygote which serves as dormant (resting) stage is formed. The wall consists of an outer exospore with ornamentation and thick pigmented meso- and endospores (Fig. 4.94d, e).

After some resting time, the zygote germinates with meiosis. Of the resulting four haploid daughter nuclei some may degenerate so that the number of haploid cells that are formed is mostly two or even just one. In placoderm desmids, the first cells that arose from zygote germination usually do not resemble the regular vegetative cells (e.g., are less ornamented) and, therefore, are named gones. The desmids’ life cycle is haplontic (Fig. 2.9), like in the Zygnematales. Sexual reproduction has been observed only

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Fig. 4.91 Desmidales morphology, two contrasting types of cell symmetries; a Micrasterias radiata. A recently divided pair of unicells that are strongly flattened. Note the cell walls with pores of the A recently divided pair of unicells that are strongly flattened. Note the cell walls with pores of the outermost left and right (older and more differentiated) semicells, the new semicells (in between) are with smooth wall covering and less differentiated; b Staurastrum vegetative cells with triradiate symmetry; scanning electron micrographs from Pickett-Heaps (1975), reproduced with permission of Oxford University Press through PLSclear

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Fig. 4.92 Pores in the cell wall of Cosmarium botrytis; a immature pore in the secondary wall which is excreted at the end of semicell expansion (see Fig. 4.93d); it does not extend through the primary wall; b Functioning pore in a vegetative cell that extends through the cell wall; a, b transmission electron micrographs from Pickett-Heaps (1975), reproduced with permission of Oxford University Press through PLSclear

sporadically in nature, probably due to sexual incompatibility. Most Desmidiales may survive adverse environmental conditions as vegetative cells rather than as dormant zygotes (Figs. 4.92, 4.93 and 4.94).

4.2.12.6 The Streptophyte Algae and Plant Terrestrialization The lineage of Streptophyta had a profound impact on the face of our planet. All multicellular plants that we see teeming on Earth’s surface can ultimately be traced back to a fateful event referred to as plant terrestrialization. This terrestrialization event has resulted in the birth of the monophylum Embryophyta—the land plants. Land plants encompass non-vascular plants (mosses, liverworts, and

hornworts) and vascular plants (spikemosses, ferns, and seed plants). In their entirety, these organisms are the macroscopic land flora. And this flora is vast. A recent estimate has put a number to the biomass these plants accumulatively represent: 80% (Bar-On et al. 2018). Furthermore, embryophyte diversity encompasses more than 450,000 species— or more. Where did this rich biodiversity come from? The answer is to be found among the streptophyte algae. The monophyletic clade of embryophytes is nested within the clade of Streptophyta. This means that land plants emerged from the grade of streptophyte algae and that the algal progenitor of land plants was a streptophyte alga. We will quickly see that there are many interesting questions to be formulated when bearing this in mind: How did the

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Fig. 4.94 Process of conjugation (formation of zygotes) during the sexual reproduction of the desmid Micrasterias rotata; a vegetative cell of M. rotata. the cells are flattened and each semicell is with four deeply incise lobes; b two paired compatible sexual cells lie flat against one other and are oriented at right angles to one another; c formation of the conjugating tube between the paired cells; the isthmus splits apart at one side and a protuberance of cytoplasm is extended toward a similar one from the other cell; the protoplasts (gametes) migrate quite rapidly out of the semicells and then they fuse to form the zygote; d mature spiky zygote with ornamentation of small spikes on the exospore; e mature sculptured zygote in Cosmarium botrytis between two empty parental cell walls; note orientation of the cells at right angle to one another (scanning electron micrograph); a–d from Lenzenweger R. in Pickett-Heaps (1975); e from Pickett-Heaps (1975) reproduced with permission of Oxford University Press through PLSclear

transition from algae to land plants happen? What are the closest algal relatives to land plants? What were the major challenges that the algae were facing? Which traits are shared by land plants and their closest algal relatives? In this sub-chapter, we will try to provide brief answers to a few of these questions. That said, each of these topics could be a

chapter of its own and many more interesting questions can be formulated. The closest algal sister lineage to land plants. Owing to advancements in phylogenomic analyses in the past decade, we have a good idea of the relationships of organisms within Streptophyta. One of the prime revelations that came from

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the recent phylogenetic frameworks was that the Zygnematophyceae represent the closest algal sister lineage to land plants, followed by the Coleochaetophyceae and the Charophyceae (One Thousand Plant Transcriptomes Initiative 2019). The latter were once considered to be the closest relatives of the land plants (van den Hoek et al. 1995; Karol et al. 2001). This came as a surprise. The Zygnematophyceae have the least elaborate body plan among the other candidate lineages that were closer to land plants. Which specific features unite the Zygnematophyceae and land plants is an active field of research—especially on the molecular level. One feature that is of key interest in the questions of how plant terrestrialization came about is to endure the specific challenges of the terrestrial habitat. These include higher irradiance (both ultraviolet and photosynthetically active light), drought, temperature fluctuations not buffered by a body of water, availability of nutrients in the substrate and other factors. The biology of land plants meets these challenges through various physiological properties. In recent years, comparative genomic analyses offer a window into which genetic properties for key land plant features can be found in streptophyte algae—and thus which genetic properties were already present in the algal progenitor of land plants. Mycorrhizal fungi. Most land plants (likely more than 80%) interact with mycorrhizal fungi, among which arbuscular mycorrhizal symbiosis is the most ancient (for more, see (Delaux and Schornack 2021). Through these symbioses, plants gain an access to enhanced nutrient uptake (e.g., acquisition of phosphorous); the fungus benefits from the photosynthates of the plants. All major lineages of land plants—except for mosses (secondary loss)—engage in these symbioses (Field and Pressel 2018). It is thus fair to say that the last common ancestor of all land plants had the ability to engage in symbiotic interaction with mycorrhizal fungi. The establishment of such symbioses rests on a well-studied chassis of proteins. Homologs for the relevant proteins can also be found across land plant diversity. So, what about streptophyte algae? Comparative genomic investigations have revealed that most of the genes for symbiotic interaction with mycorrhizal fungi have clear homologs in those streptophyte algae more closely related to land plants (Delaux et al. 2015; Nishiyama et al. 2018). Terrestrial stressors and desiccation tolerance. Land plants meet the barrage of terrestrial stressors with the production of protecting specialized metabolites. Such specialized metabolites are to be found across the diversity of land plants, often showing a lineage specific repertoire of compounds. However, there is a common pathway chassis for many of these metabolites. One such chassis has currently been pushed into the limelight: the phenylpropanoid pathway. Homologs for many of the genes of this pathway are present in streptophyte algae (de Vries et al. 2017, 2021).

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Furthermore, there are some recent data on the presence of compounds that emerge from the phenylpropanoid pathway in streptophyte algae, including lignin-like compounds (Sørensen et al. 2011) and flavonoids (Jiao et al. 2020). Many Zygnematophyceae are desiccation-tolerant; further, recent comparative genomic investigations have unveiled that some Zygnematophyceae have a full homologous set of genes that land plants use for mounting a signaling cascade triggered by the phytohormone abscisic acid (de Vries et al. 2018; Cheng et al. 2019). This warrants attention, as abscisic acid is one of the major signaling molecules that land plants employ when they are challenged by drought stress. However, the zygnematophyceaen cascade likely works in a different manner: the algal protein homologous to the abscisic acid receptor of land plants does interact with other relevant components in the signaling cascade—but in an abscisic acid independent manner (Sun et al. 2019). Thus, while the deep roots of the proteins that act in the cascade have been pinpointed, the physiological relevance these proteins have in Zygnematophyceae is currently unknown. That said, such molecular components were likely recruited early during land plant evolution towards the assembly of a drought stress-relevant signaling network. Phytohormones. Land plant growth and physiology is tightly interwoven with phytohormone signaling. The evolutionary roots of phytohormone signaling pathways (not only abscisic acid) in streptophyte algae are hence of major relevance to our understanding of how key traits of land plants evolved. Auxin is the foremost phytohormones regulating plant growth and development. Efforts have been made to pinpoint the role of auxin in streptophyte algae, however resulting in a complicated and inconclusive picture (e.g., Vosolsobě et al. 2020). The picture is more straightforward when it comes to presence of homologs for auxin signaling (Flores-Sandoval et al. 2018; Mutte et al. 2018; Martin-Arevalillo et al. 2019) and its polar transport (Skokan et al. 2019)—which is critical for the action of auxin in growth modulation. In sum, similar to the role of abscisic acid, the role of auxin in streptophyte algae is unclear; the evolutionary roots for components that are of importance for the action of these phytohormones can, however, be traced back to streptophyte algae. Multicellularity, phragmoplast, and plasmodesmata. Multicellularity is the reason why land plants can form specialized cell and tissue types that are particularly apt at performing a certain task: nutrient uptake from the soil, forming the interface to the air, reproduction, and propagation; furthermore, these differentiations paved the path towards upright growth. Land plants’ multicellularity is realized through a specific cell biological chassis, the phragmoplast, which is one defining feature of how plant cells are formed. The phragmoplast consists of cytoskeletal components and vesicles filled with material for the nascent

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cell wall; it is formed during cytokinesis and marks the line along which the new cell wall (cell plate) is drawn that will separate mother and daughter cells. The formation of a phragmoplast is a defining feature that unites Charophyceae, Coleochaetophyceae, Zygnematophyceae, and Embryophyta —all form together a monophyletic entity. For more aspects on streptophyte cell division see (Buschmann and Zachgo 2016). A key feature of multicellular phragmoplastophytes is the formation of plasmodesmata between adjacent cells. Plasmodesmata connect the cytosol of cells, leading to the symplastic continuum found among all land plants. Connections through plasmodesmata can be formed through two mechanisms: during cell division, resulting in primary plasmodesmata, and after cytokinesis (by specifically creating holes in an existing cell wall), resulting in secondary plasmodesmata. Some streptophyte algae have plasmodesmata. However, the occurrence of these occurs in an interesting pattern: Charophyceae have both primary and secondary plasmodesmata, Coleochaetophyceae have only secondary plasmodesmata, and Zygnematophyceae have neither. This nicely illustrates the complexities of tracing the evolutionary roots of land plant traits among streptophyte algae.

4.3

Glaucophyta

Burkhard Büdel Among the monophyletic Archaeplastida (e.g., Jackson et al. 2014), the phylum Glaucophyta is the least species-rich with only four accepted genera and approximately 19 species known so far. Phylogenetically, they are descendants of primary endosymbiosis between a heterotrophic eukaryote and a cyanobacterial endosymbiont roughly 1,200–1,600 million years ago (Yoon et al. 2004, 2010; Rodríguez-Espeleta et al. 2005; Bengtson et al. 2017). Like green plants and red algae, they are primary plastid bearing organisms.

4.3.1 Origin of the Phylum Glaucophyta Eukaryotic photoautotrophic organisms acquired their ability to convert light into chemical energy through endosymbiosis with a cyanobacterium (see Chap. 2). This first endosymbiosis event gave rise to the “primary” plastids, which are present in all green plants (including green algae), the red algae, and the glaucophytes. All their plastids are characterized by being surrounded by two envelope membranes. The glaucophyte plastids are in the form of a cyanelle, an organelle distinct from the chloroplasts of other organisms in that, like cyanobacteria, they have a conspicuous

peptidoglycan wall between its two membranes (Adl et al. 2019). These plastids were originally named “cyanelles”, which was later changed to “muroplasts” when their shared ancestry with other Archaeplastida was recognized (Price et al. 2017). Glaucophytes are supposed to share a common ancestor with the red algae (Rhodophyceae) and a lineage comprising both green algae and land plants (Chloroplastida), (nomenclature used here taken from Adl et al. 2019). As there is no fossil record for the glaucophytes, age determinations are solely based on molecular clock estimates.

4.3.2 Morphology and Cell Structure/Function The glaucophytes are freshwater phototrophs that can be unicellular or colonial and are organized as either monadoid, palmelloid, capsalean, or coccoid. Cell division and reproduction is achieved by binary fission, successive binary fission, or a progressive cleavage into autospores. Sexual reproduction is unknown. The monadoid stages always have a dorsoventral symmetry (Cyanophora, Cyanoptyche, Gloeochaete) and the flagella, when present (Cyanophora, Gloeochaete-zoospores) are pairwise with mastigonemes. The vegetative cells of Gloeochaete are characterized by two pseudocilia and those of the genus Glaucocystis expose two reduced flagella. During the endosymbiotic transition process of the affiliated cyanobacterium to a plastid, the biosynthesis of cytosolic starch also went through a transition process of glycogen to starch storage.

4.3.2.1 Cell Wall and Cell Surface A cell wall is not found in all genera of the glaucophytes. The cell wall of the genus Cyanoptyche is characterized by mucopolysaccharides, in the genus Gloeochaete a non-cellulosic cell wall exists in vegetative cells but no cell wall is found in their zoospores. The cell wall of the genus Glaucocystis is cellulosic and the genus Cyanophora has no cell wall at all. A layer of flat vesicles underneath the plasmalemma occurs in all genera, in the genus Cyanophora, for example, a plate-like structure occurs above the flat vesicles, thus forming a pellicle (Figs. 4.95 and 4.96). A characteristic surface ornamentation formed by the edges of overlapping or attached outermost plate vesicles at the cell periphery (Figs. 4.95 and 4.96) was found in Cyanophora biloba and C. paradoxa, which has been described in detail using various electron microscopic methodologies (Kugrens et al. 1999; Takahashi et al. 2014). The presence of two different colony surface (“mother cell wall”) cellulose fibril types was described for two species groups in the genus Glaucocystis on the basis of modern electron microscopy: the gauze fabric-like fibrils and tightly arranged fibrils (Takahashi et al. 2016).

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Fig. 4.95 Ultrastructure of a glaucophyte cell, Cyanophora paradoxa. Cell surface ornamentation according to Kugrens et al. (1999) and Takahashi et al. (2014). Original drawing Spindler & Büdel

4.3.2.2 The Muroplast The muroplast is characteristically surrounded by an inner and outer plasmalemma layer with the inner membrane originating from the originally endo-cytobiologically affiliated cyanobacterium and the outer membrane from the host cell. Between the two membranes, a periplasmic space is inserted where also the size reduced peptidoglygan wall of cyanobacterium is located (Figs. 4.95 and 4.96; Kies 1976). The thylakoid membranes are concentrically arranged and occur in multiple layers. In the center of each plastid, a large (almost up to 1 µm) central body occurs which was shown to contain the bulk of RuBisCO in

Cyanophora paradoxa (Mangeney and Gibbs 1987). This structure has often been denoted the “carboxysome”, probably because of the comparable function in cyanobacteria. The photosynthetic apparatus, with its phycobilisome antennae, still resembles that of cyanobacteria. An evolutionarily early form of a Toc/Tic translocon is involved in protein import into muroplasts, suggesting that the Toc/Tic system was likely to have been in place in the common ancestor of the Archaeplastida. The muroplast genome itself is unique because of the presence of genes that are not known from other plastid genomes (Price et al. 2017).

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4.3.3 Genome

4.3.4 Classification and Systematic Arrangement of the Glaucophyta

The contribution of cyanobacterial sources to the glaucophyte Cyanophora plastid proteome is, with 12%, considerably lower than that of other Archaeplastida, where 19% occur in the green alga Chlamydomonas and 24% in the vascular plant Arabidopsis (Qiu et al. 2013). The nuclear genome is gene-rich (27,000) compared with the relatively small genomes of unicellular red algae (6,000–10,000 genes; Price et al. 2017). Like in other Archaeplastida, the glaucophyte mitochondrial DNAs (mtDNA) are circular. The mtDNA gene set is quite similar among the different glaucophyte genera and it is comparable to the gene-rich mtDNAs of the green and red algae (Price et al. 2012). Gloeochaete wittrockiana and Glaucocystis nostochinearum mtDNAs lack certain tRNA codons, whereas Cyanophora paradoxa and Cyanoptyche gloeocystis lack trnT genes entirely. As genome data are available from a small number of taxa, only limited conclusions can be drawn at present. However, although substantial evidence suggests that the glaucophytes, red algae, and viridiplantae share a common ancestor, some phylogenetic analyses do not recover these three lineages as a monophyletic clade (Jackson et al. 2015).

The systematic arrangement of the Glaucophyta presented here (Table 4.5) follows Price et al. (2017) and Jackson et al. (2015) based on multigene analyses (see Box 4.2). Species numbers are according to Guiry and Guiry (2023). In the classification system of Adl et al. (2019), the phylum is given a family rank within the Archaeplastida but with uncertain affiliation. Phylogenetic analyses using plastid-encoded genes of Cyanophora paradoxa suggested that the glaucophytes where the first to diverge after the single primary endosymbiotic event, a view that was later supported by the study of nuclear genes of the same genus (Martin et al. 1998; Rodríguez-Espeleta et al. 2005; Reyes-Prieto and Bhattacharya 2007). In a model study of modern taxonomy, based on ultra-highvoltage electron microscopy, field-emission scanning electron microscopy as well as differences in the light microscopic characteristics and molecular phylogenetic results, Glaucocystis strains were exemplary delineated into species. In doing so, the performing scientists were able to establish a new taxonomic arrangement, validating and describing six species of the genus Glaucocystis (Takahashi et al. 2016).

Table 4.5 Overview of the current classification of the Glaucophyta Class Glaucophyceae

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Box 4.2: Phylogeny of the Glaucophytes

Phylogeny of the Glaucophyta and their geographical origin, based on the gene regions of plastidal psbA, psaB, 16S rRNA, mitochondrial cox 1, cob, and nuclear ITS region including ITS 1 and 2, 5.8S, partial SSU, and LSU. Large letters indicate different morphologies in Cyanophora, large letters + numbers indicate different clades according to Chong et al. 2014. Figure modified from Price et al. 2017, morphological characters taken from Kies (1979, 1989).

202 Fig. 4.97 Glaucophyta: a, b Glaucocystis nostochinearum, colonial organization, view of upper cell surface (a) and cell center (b), numerous muroplasts per cell, each cell is surrounded by an own envelope, envelopes of the former mother cell remain and form the multiple envelopes of the colony (e1–e4); c Gloeochaete wittrockiana, colony with pseudocilia (pc), phase-contrast microscopy; d G. wittrockiana, cells with clearly visible nuclei (n). Pictures a and b courtesy of Gerd Günther, using algal strains from the CCAC; figures c and d courtesy of Ludwig Kies

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4.3.4.1 Gloeochaete Only one species, Gloeochaete wittrockiana, is known. The species lives in single cells or forms colonies. It has a capsal organization and lives epiphytically on green algae like Oedogonium, Chara, and Vaucheria (Kies 1976). Cells with one or two pseudocilia per cell. Plastids are many and are arranged closely together in cells. In the species Gloeochaete wittrockiana, each cell exposes two long pseudocilia (Kies 1976; Fig. 4.97c, d). 4.3.4.2 Cyanoptyche Cyanoptyche gloeocystis is a palmelloid colonial alga and so far the only valid species of the genus (Fig. 4.98a, b). Cell groups are not bound by a mother cell wall. Along the periphery of the cells, a peculiar lacunae system occurs. Vegetative cells have two rudimentary flagella, zoospores are dorsiventrally shaped with two heterokont and heterodynamic flagella, originating from a subapical depression. Both flagella possess non-tubular mastigonemes. Main reserve product is starch lying freely in the cytoplasm. Plastids are rounded to slightly elongate (Kies 1989; Jackson et al. 2015). 4.3.4.3 Glaucocystis So far 13 species known. Species of the genus occur as single cells or in a colony and then the cells are embedded in

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a common matrix bound by the original mother cell wall. The autospores remain enwrapped in the mother cell wall (Fig. 4.97a, b) which is very expandable and can harbor up to three daughter cell generations before opening by rupture (Schnepf et al. 1966). Cells are typically oval-shaped and flagella have been reported, however, the evidence for flagella in Glaucocystis is unclear and requires verification. Plastids are in star-shaped groups (Jackson et al. 2015).

4.3.4.4 Cyanophora From this genus 6 species are known so far. Cells are small (9–16 lm  7 lm). Bi-flagellated cells divide by binary fission and they are able to form round cysts. One or more muroplasts are present in each cell and are always even numbered (Fig. 4.98c), (Jackson et al. 2015). The reproduction mode is a longitudinal cleavage.

4.3.5 Ecology The currently known glaucophyte species are all freshwater organisms. Although still limited, we do have some information on the geographical distribution of the glaucophytes. For example, the species Glaucocystis nostochinearum has been most frequently reported and has a more or less cosmopolitan distribution, while other Glaucocystis species are

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Fig. 4.98 Glaucophyta: a Cyanoptyche gloeocystis, colony of large cells with numerous muroplasts; b same species, DAPI staining, and fluorescence microscopy used to show the nucleus (blueish) and muroplasts (red); c Cyanophora paradoxa, monadoid cells with clearly visible muroplast and flagella. All pictures courtesy of Gerd Günther using strains from CCAC

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reported only sporadically. Cyanophora species are also widespread and were reported from China, Germany, Great Britain, Italy, Japan, Netherlands, Russia, Sweden, Ukraine, and USA. Gloeochaete wittrockiana is another quite common species announced from Georgia, Germany, Great Britain, Romania, Scandinavia, Slovakia, and Spain. Cyanoptyche species are less frequently found and are so far known from Spain and the Netherlands only (all distribution data from Guiry and Guiry 2023). In general, glaucophytes are not easy to collect and this is one of the reasons for the rather incomplete knowledge regarding their ecology in general and their characteristic habitats in detail. Glaucophytes were found in small eutrophic ditches near Kharkov, Ukraine, alkaline waters in England, fishponds in Germany, as river plankton in Belarus, in the littoral zone of a lake in Sweden, and in Sphagnum bogs. Also, quite a number of

species are regularly found associated epiphytically with submerged vascular plants, bryophytes, and larger filamentous green algae (Pascher 1929a, b; Pringsheim 1958a, b; Skuja 1956; Geitler 1959; Kies 1979).

4.4

Cercozoa—A Second Primary Endosymbiosis

Burkhard Büdel The phylum Cercozoa circumscribes amoeboids and flagellates feeding by means of filose pseudopods. It harbors the algal lineage of the classes Chlorarachniophyceae (secondary endosymbiosis) and the Filosa with the only genus Paulinella, which is apparently a descendant of a primary

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endosymbiosis (Delaye et al. 2016; Sánchez-Baracaldo et al. 2017; Lhee et al. 2019). The closer taxonomic affiliation within the Cercozoa of the recently described new cercozoan genus and species, Aurantocordis quadriverberis, is still unknown (Chantangsi et al. 2008).

4.4.1 Primary Endosymbiosis? The origin of photosynthesis in eukaryotic organisms certainly was a revolutionary process in the evolution of life. It is now commonly agreed upon that endosymbiosis establishing the chloroplast lineage in eukaryotes can be traced back to a single event. It is suggested that the Archaeplastida, consisting of glaucophytes, red algae, green algae, and land plants, share a common ancestor that lived  1900 Mya (Sánchez-Baracaldo et al. 2017). However, the discovery that the photosynthetic compartment of the thecate, filose amoeba Paulinella chromatophora originated by a similar but more recent process, suggested a second primary endosymbiotic event independent of the primary endosymbiotic process of the Archaeplastida. This new discovery led to the understanding that an endosymbiotic gain of photosynthetic organelles from prokaryotes may be an ongoing process (Marin et al. 2005). The recently described tetraflagellate protist Auranticordis quadriverberis from marine sand in British Columbia, Canada, is phylogenetically clearly related to the cercozoans. However, although displaying numerous orange-colored prokaryotic endosymbionts, their interpretation as cyanobacteria-derived is still unsafe as, for example, the characteristic phycobilisomes are absent (Chantangsi et al. 2008).

4.4.2 Origin Using a lognormal relaxed molecular clock procedure on the 18S rRNA of P. chromatophora, time estimates show that depending on the assumptions made to calibrate the molecular clock, P. chromatophora diverged from heterotrophic Paulinella spp. *90 to 140 Mya (Delaye et al. 2016). This assumption, together with phylogenomic analyses of the plastid, suggested that the ancestor of P. chromatophora established a symbiotic relationship with a cyanobacterium of the Prochloroccocus/Synechococcus guild (e.g., Marin et al. 2005). A comparison of the pseudogene disintegration rate in the heterotrophic bacterial endosymbiont of aphids, Buchnera aphidicola, showing that a pseudogene needs about 40–60 million years to disintegrate completely, led to the speculation of a minimum age of *60 million years for the chromatophore (Delaye et al. 2016).

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Presently Paulinella is the only recent case of primary plastid endosymbiosis. This makes it a valuable model to study the transformation of a “host” from a heterotrophic to a phototrophic lifestyle, as well as the way from a free-living cyanobacterium into a well-integrated photosynthetic cell compartment. Important research questions could be answered using the model system as suggested by Gabr et al. (2020): (1) what is the role horizontal gene transfer in plastid acquisition and integration? (2) which host-derived genes are crucial for the maintenance of the organelle? (3) are there pre-adaptive genes that existed in the genomes of heterotrophic Paulinella, probably derived from horizontal gene transfer and facilitating survival and maintenance of the endocytosed a-cyanobacterium (see Chap. 3 for explanation)? (4) which host-derived genes are not chromatophore targeted but play a role in regulating plastid functions, such as division?

4.4.3 Morphology and Ultrastructure Paulinella chromatophora is a thecate amoeba whose single-celled body is covered by a case composed of silica-scales that form the theca. The rectangular scales are arranged in a regular manner around the cell. The scales are produced singly prior to cell division in large vesicles by the only dictyosom of the cell, closely located to the nucleus (Kies 1974). Paulinella regularly hosts two blue-green, sausage-shaped plastids that have been described previously as endosymbiotic cyanobacteria or cyanobacterial-like organisms by Lauterborn (1895), they function akin to chromatophores for the amoeba. The cytoplasm of the Paulinella-cell is only attached to theca at the apical theca opening. Cell motility is achieved by means of 1–4 thin filamentous pseudopodia (filopodia), originating from a small apical dome-shaped structure (Figs. 4.99b and 4.100a). At the apical pole 2–3 pulsating vacuoles are located (Kies 1974). The chromatophores were later termed “cyanelles” by the prominent phycologist Pascher (1929a, b) to pronounce their chloroplast-like function. The plastids have a 6–13 nm thick wall (Kies 1974), referring to the cyanobacterial peptidoglycan wall (Reyes-Prieto et al. 2007). The plastids are completely surrounded by a host double membrane. Inside the plastid, 15–20 thylakoids are concentrically arranged and plastoglobuli and phycobilisomes are present. In the thylakoid free central part of the chromatophores plasma polyhedral bodies, most probably carboxysomes (authors remark), are located (Kies 1974). The conspicuous, large, sausage-shaped endosymbiont was soon thought to be related to the cyanobacterial genus Synechococcus. It was observed that when squeezed out of the host, they remain

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Fig. 4.99 Paulinella chromatophora with its prominent sausage-shaped photosynthetic symbionts; a thecate cell with pseudopodia (filopodia); b detail of cell body showing the cytoplasm with nucleus and the two sausage-shaped cyanellae/plastids. Cutouts taken from Robert Lauterborn’s original description of this organism in the Zeitschrift für wissenschaftliche Zoologie (1895; Vol. 59, 537–544; Fig. 1)

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10 µm

Fig. 4.100 Cercozoa: a Paulinella chromatophora, monadoid unicellular alga from the thecate amoeba with sausage-shaped plastids; b Auranticordis quadriverberis, image focused on rows of longitudinally arranged orange muciferous bodies (white arrows), ventral depression (vd) and the ventral groove (black arrowheads). Image a courtesy of Gerd Günther using a strain from CCAC; image b courtesy of Heather J Esson and Chitchai Chantangsi; from Chantangsi et al. (2008), BMC Microbiology

alive for some hours (Fritsch 1945). The prominent phycologist Fritsch (1945) reported that: “During cell division of Paulinella, one of the two cyanellae passes through the narrow aperture of the test into the new daughter-individual and, like that left in the parent, soon divides into two”. Phylogenomic studies revealed the alpha-cyanobacterial provenance (see Chap. 3 for a-cyanobacteria) of the Paulinella plastids and that primary endosymbiosis occurred roughly 90–140 Mya. A comparison of the complete

chromatophore genome sequences of the species P. longichromatophora and P. micropora with those from existing chromatophore genomes revealed a basal split among photosynthetic Paulinella species *60 Mya. During the early stages of endosymbiosis major gene losses occurred, but the process slowed down significantly later on, resulting in a conserved gene content across extant taxa. Only 35% of the ancestral gene families from the cyanobacterial endosymbiont remained in chromatophore

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DNA (Lhee et al. 2019). To conclude, Paulinella’s photosynthetic endosymbionts have undergone evolutionary changes that can be interpreted as typical for primary plastids (Marin et al. 2007).

4.4.4 Classification and Systematic Arrangement Presently, two different systematic and taxonomic arrangements of the genus Paulinella exist (Table 4.6): (a) the more sophisticated system based on the results of multiple authors (e.g., Kim and Park 2016) presented in Guiry and Guiry (2023), and (b) the still “in work” system of Adl et al. (2019). Presently, the genus Paulinella comprises five species of freshwater amoeboids: P. chromatophora, P. micropora, P. longichromatophora, P. gracilis, and P. ovalis (Guiry and Guiry 2021). Only very little is known about the ecology of Paulinella species. While P. chromatophora, P. gracilis, and P. micropora are freshwater species, P. longichromatophora originates from a marine habitat (Lhee et al. 2019). Nothing is known about the environmental demands of P. ovalis.

Auranticordis quadriverberis is a new tetraflagellate genus and protist species described by Chantangsi et al. (2008). It was discovered and isolated from marine littoral sand samples in British Columbia, Canada. Morphological features did not allow A. quadriverberis to be assigned to any known eukaryotic guild. However, sequencing the small subunit of rDNA revealed that this lineage evolved from within the Cercozoa. The genus and species is characterized by gliding motility associated with four bundled recurrent flagella, heart-shaped cells about 35–75 lm in diameter, and bright orange coloration caused by linear arrays of muciferous bodies (Fig. 4.100b). Additionally, each cell contains about 2–30 pale orange bodies, enveloped by two membranes and sac-like vesicles (Fig. 4.101). The flagella are arranged in two pairs and are covered by flagellar hairs (mastigomenes). Inside the putative endosymbiont cells, the thylakoids originate from inner membrane invaginations that extend towards a central body, resembling a pyrenoid (Fig. 4.101). The light orange bodies have a size of 4–14 µm in diameter and their ultrastructure is most consistent with photosynthetic endosymbionts of cyanobacterial origin, differing from cyanobacteria by the feature of continuing cell-membrane connectivity of the thylakoids. Three

Table 4.6 The two different systematic and taxonomic arrangements of the genus Paulinella (phylum Cercozoa) Class

Order

Family

Genus

a) In-depth model unclear affiliation yet: Chlorarachniophyceae (treated in Chapter 5)

Aconchulinida

Auranticordis

Paulinellidae

Paulinella

(b) According to Adl et al. (2019) unclear affiliation yet: Silicofilosea

Euglyphida

Auranticordis

Paulinellidae

Paulinella

thylakoids

sc sc sc

putative pyrenoids

sc 10 µm

Fig. 4.101 Auranticordis quadriverberis; transmission electron microscopy of putative primary endosymbionts in the host cell; all endosymbionts are surrounded by sac-like vesicles (sc). From Chantangsi et al. 2008, BMC Microbiology 2008, 8: Fig. 5a

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hypotheses for the identity of the light orange bodies are offered by the researchers: (1) they are ingested photosynthetic prey cells in the earliest stages of degradation; (2) they are transient photosynthetic endosymbionts continuously replenished by kleptoplasty; or (3) they are permanently integrated photosynthetic endosymbionts and thus a special new type of plastid. For a better identification of their nature and function, experiments involving autofluorescence and amplification of plastid molecular markers must be performed. However, considering the rarity of this species in nature and the fact that it cannot yet be kept in culture, does not make further investigations an easy task at all (Chantangsi et al. 2008).

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5

Algae from Secondary Endosymbiosis Thomas Friedl

Contents 5.1 Heterokontophyta—Photosynthetic Stramenopiles.......................................................................... 220 5.1.1 General Ecology and Importance............................................................................................... 222 5.1.2 General Description .................................................................................................................... 223 5.1.3 Evolutionary History .................................................................................................................. 226 5.1.4 Taxonomic Classes ..................................................................................................................... 229 5.1.5 Perspectives................................................................................................................................. 276 5.2 Dinoflagellates ...................................................................................................................................... 280 5.2.1 Organization and Structural Features of Dinoflagellates Cells.................................................. 281 5.2.2 Reproduction............................................................................................................................... 287 5.2.3 Chloroplasts ................................................................................................................................ 287 5.2.4 Kleptoplasty ................................................................................................................................ 291 5.2.5 Non-Photosynthetic Nutrition..................................................................................................... 291 5.2.6 Bioluminescence ......................................................................................................................... 292 5.2.7 Toxins and Harmful Algal Blooms............................................................................................ 292 5.2.8 Phylogeny: Classification ........................................................................................................... 293 5.3 A Cercozoan Secondary Endosymbiosis: Chlorarachniophyta...................................................... 297 5.3.1 General Characters...................................................................................................................... 297 5.3.2 History of Research .................................................................................................................... 297 5.3.3 Morphology and Developmental Stages .................................................................................... 297 5.3.4 Reproduction and Life Cycle ..................................................................................................... 299 5.3.5 Phylogeny and Systematics ........................................................................................................ 300 5.4 Euglenids—(Excavates, Discoba, Euglenozoa, and Euglenida)...................................................... 302 5.4.1 Short Introduction—what Are Euglenids? Why Are They Called Augentierchen? ........................................................................................................................... 302 5.4.2 Taxonomic Classification ........................................................................................................... 303 5.4.3 Origin and Fossil Record ........................................................................................................... 303 5.4.4 History of Research .................................................................................................................... 304 5.4.5 General Information and Diversity of Nutrition Modes............................................................ 305 5.4.6 Characters Uniting Euglenids—An Overview of Morphology and Cell Structure...................................................................................................................................... 309 5.4.7 Phylogenetic Position—Euglenida ............................................................................................. 319

T. Friedl (&) Department of Experimental Phycology and Culture Collection of Algae (EPSAG), Georg August University Göttingen, Nikolausberger Weg 18, 37073 Göttingen, Germany e-mail: [email protected] © Der/die Autor(en), exklusiv lizenziert durch Springer-Verlag GmbH, DE, ein Teil von Springer Nature 2024 B. Büdel et al. (eds.), Biology of Algae, Lichens and Bryophytes, https://doi.org/10.1007/978-3-662-65712-6_5

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220

T. Friedl 5.4.8 Ecology—Where Do We Find Euglenids?................................................................................ 320 5.4.9 Description of Easily Observed Taxa ........................................................................................ 321 5.5 Haptista................................................................................................................................................. 323 5.5.1 General Description .................................................................................................................... 323 5.5.2 Fossil Record .............................................................................................................................. 323 5.5.3 Molecular Clock Record ............................................................................................................ 324 5.5.4 Morphology and Ultrastructure .................................................................................................. 324 5.5.5 Life Cycle ................................................................................................................................... 326 5.5.6 Phylogeny and Classification ..................................................................................................... 328 5.5.7 Eco-Physiology ........................................................................................................................... 330 5.6 Cryptista ............................................................................................................................................... 333 5.6.1 Cell Structure and Function ....................................................................................................... 333 5.6.2 Habitats and Survival Strategies ................................................................................................ 346 5.6.3 Origin and Evolution of the Cryptista ....................................................................................... 350 References ..................................................................................................................................................... 353

5.1

Heterokontophyta—Photosynthetic Stramenopiles

Louis Graf The photosynthetic world is not only dominated by the green color of trees and plants. Large portions of the photosynthetic organisms of the planet Earth dwell in a brownish and yellowish world. These algae are principally members of the large group Stramenopiles. The Stramenopiles is a monophyletic group characterized by the presence of two flagella of unequal length in motile stages (Fig. 5.1). The long anteriorly directed flagellum (immature flagellum) bears two rows of tripartite hairs. The other flagellum (mature flagellum) is short, posteriorly directed, and without hairs. The Stramenopiles forms with the Alveolata (a group comprising notably the microplanktonic Dinoflagellata (Chapter 5.2; Saldarriaga and Taylors 2017), the parasitic Apicomplexa (Votypka et al. 2017), the Ciliophora (Lynn 2017)), and the Rhizaria (a group of mostly heterotrophic amoebae or flagellates like the Foraminifera or the Radiolaria (Boltovskoyet al. 2017)), the supergroup SAR (= Stramenopiles-Alveolates-Rhizaria; see Opuntia tree of life Figs. 1.1 and Fig. 2.8; and Strassert et al. 2019). However, there is a discussion if the Telonemia (= single-celled heterotrophic and photosynthetic species) should not be added to the SAR to form the TSAR-supergroup (Burki et al. 2020). Like the other groups of the SAR, the Stramenopiles combine an immense number of species presenting a wide range of forms, organization, biology, and ecology. The Stramenopiles regroup both photosynthetic and heterotrophic lineages that are monophyletic and sisters to each

other (see Opuntia tree of life Figs. 1.1 and Fig. 2.8). In this chapter, we will only discuss the photosynthetic lineages of the Stramenopiles, hereafter referred to as Heterokontophyta (Box 5.1), and information on the heterotrophic lineages can be found in chapters 13 to 15 of the Handbook of the Protists (Archibald et al. 2017). Most likely, the readers of this chapter might have first encountered members of the Heterokontophyta while walking on the seashores, looking at the brown seaweeds and other kelps. These were actually the first recorded Heterokontophyta in history as there are mentions of them dating back to early China (ca. 3000 BC), Japan (ca. 500 BC), and Greece (ca. 300 BC). Furthermore, the use of brown seaweeds for human and cattle food, medicine, and dyes likely predates these records and potentially played a role in the early dispersal of humans along the seashores (Erlandson et al. 2015; Braje et al. 2017). It is also not surprising that Linneaus (1753), the father of modern taxonomy, formally described the first member of the Heterokontophyta, i.e. the brown alga Fucus. In the following years, more elusive Stramenopiles were described as well as the microscopic Chrysophyceae and diatoms by Müller (1773, 1783, 1786). These descriptions opened a 100 years period of exploration (from 1753 to 1882) of the diversity of the Stramenopiles that saw brown algae described as plants and motile microalgae as animals. Seminal works were published at that time, notably the use of color to classify algae by Lamouroux (1813) that lead to the premise of the Phaeophyceae; or Ehrenberg’s book (1838) regrouping his microscopic observations including organisms now included in the Stramenopiles. However, these taxa remained separated from each other in the mind of scientists until the work of Rostanfinski (1882)

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Algae from Secondary Endosymbiosis

221

d

immature flagellum

m f

e

mature flagellum

n d

thylakoids girdle lamella inner membrane pair outer membrane pair

Fig. 5.1 Flagellate cell of the Heterokontophyta. The cell possesses two flagella of unequal length, a short smooth mature flagellum that is posteriorly directed and a long immature flagellum that is anteriorly directed and bears two rows of hairs. Each flagellar hair (or mastigoneme) is tripartite, i.e. it consists of three parts, a basal attachment region, a tubular shaft, and terminal fibrils. The chloroplast has an envelope of four membranes (two membrane pairs). It encloses the thylakoids, stacked in three, which all are enveloped by a girdle lamella, i.e. a thylakoid that runs around the periphery of the chloroplast beneath the innermost membrane of the chloroplast envelope. The outermost chloroplast membrane is covered by ribosomes. The nucleus (n) is with ribosomes on its outer surface and is connected to the chloroplast. An eyespot (e) is inside the chloroplast associated with the mature flagellum, forming the photoreceptor apparatus. The mitochondria (m) contain tubular cristae. d, chrysolaminarin vacuoles or oil droplets; f, flagellar apparatus. Original drawing Spindler & Friedl

that launched a synthesis period (from 1882 to 1914). He hypothesized an evolutionary link between the diatoms, Chrysophyceae, and Phaeophyceae. This idea was followed in a series of publications (Correns 1892; Klebs 1892a, b; Lemmermann 1899) and culminated in the famous Blackman’s phylogeny (1900). However, these evolutionary relationships were not fully accepted and the final synthesis from Pascher (1914) separated the Chrysophyta (including the Chrysophyceae, Diatomeae, and Xanthophyceae) and the Phaeophyta (Phaeophyceae). Over the next 50 years, evolutionary discussions were left hanging due to the lack of observable homologous traits between these different groups. Indeed, taking for example the various cell coverings of

Heterokontophyta, how does one compare the cell wall of the Phaeophyceae composed of alginate, fucose-containing sulfated polysaccharide and cellulose (see 5.1.4.1), the naked (wallless) cells of the Chrysophyceae (see 5.1.4.2), and the silicified frustule of the Diatomeae (see 5.1.4.4)? Nevertheless, during this 50-year period, many species were described, which increased our understanding of the Stramenopiles and also demonstrated the complexity of their evolutionary history. There was a resurgence of phylogenetic interest in the mid-twentieth century (e.g. Chadefaud 1950; Bourrelly 1957). At the same time, there was a development of new technologies, such as transmission electron microscopy (e.g. Dodge 1973; Hibberd 1976; Taylor 1976; Andersen 1987)

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that led to new morphological (ultrastructural) characters and TLC/HPLC that led to new biochemical (photosynthetic pigmentation) knowledge (e.g. Strain 1951; Quillet 1955; Archibald et al. 1963; Ragan and Chapman 1978), all of which provided new informative characters. Along with the rise of computer-assisted cladistics and molecular systematics, these led to major revisions of Pascher's synthesis. Numerous classes of Heterokontophyta were recognized to be distinct from the established classes (Fig. 5.2). The Eustigmatophyceae (Hibberd and Leedale 1970) were separated from the Xanthophyceae. While out of the Chrysophyceae, the Dictyochophyceae (Silva 1980), the Chrysomerophyceae (Cavalier-Smith et al. 1995), the Pelagophyceae (Andersen et al. 1993), and the Phaeothamniophyceae (Bailey et al. 1998) were established. This period also saw the discovery of entirely new lineages, the Bolidophyceae (Guillou et al. 1999), the Pinguiophyceae (Kawachi et al. 2002a, b), the Schizocladiophyceae (Kawai et al. 2003), the Synchromophyceae (Horn et al. 2007), and the Aurearenophyceae (Kai et al. 2008) and continues to this day with the Chrysoparadoxophyceae (Wetherbee et al. 2019), the Phaeosacciophyceae (Graf et al. 2020b), and Olisthodiscophyceae (Barcytè et al. 2021). With the sequencing of diatom genomes (e.g. Thalassiosira pseudonana, Armbrust et al. 2004), brown alga genome (Ectocarpus siliculosus, Cock et al. 2010), Pelagophyceae (Aureococcus anophagefferens, Gobler et al. 2011), and Eustigmatophyceae (e.g. Nannochloropsis gaditana, Radakovits et al. 2012), the research on Heterokontophyta entered another era. Genomic data (i.e. genome sequences, transcriptomes, and proteomes) allowed unprecedented exploration of the biology, ecology, and evolution of the Heterokontophyta. The relatively complex history of our knowledge of the Heterokontophyta is summarized in Fig. 5.2. A thorough discussion on the history of the Heterokontophyta (Andersen 2004) will provide more interesting details on this subject. Currently, 17 classes are recognized and these classes are divided into three major clades (Fig. 5.2). For the global phylogenetic position of Heterokontophyta, see the “(Opuntia-) Tree of Life, see Figs. 1.1 and 2.8.

5.1.1 General Ecology and Importance From open oceans to coastal estuaries, from mountain streams to lakes, from soil to snow, Stramenopiles are found virtually in every environment supporting life on Earth. Extremely diverse and highly successful ecologically, they are abundant in aquatic environments where they can dominate the eukaryotic community. The recent exploration of the eukaryotic diversity using metagenomics methods

T. Friedl

confirmed the cosmopolitic nature of this group and their importance (e.g. Moon-van der Staay et al. 2001; Díez et al. 2001; Seeleuthner et al. 2018; Fawley et al. 2021). As photosynthetic organisms, they are the principal primary producers in many ecosystems. The Diatomeae alone are thought to be responsible for 20% of the total global C-fixation (Mann 1999). The algae of the Phaeophyceae, and notably the kelps, are also important species in the coastal ecosystems where they not only act as primary producers (with carbon uptake of up to 1 kg C m−2 yr−1) (Mann 1973; Pfister et al. 2019) but also transform and shape the ecosystem to support other species, as so-called engineer species (Teagle et al. 2017). It appears that early on during their evolution, some lineages showed clear ecological preferences toward either a marine or a freshwater environment (Table 5.1). Algae of the Aurearenophyceae, Bolidophyceae, Chrysoparadoxophyceae, Pelagophyceae, Pinguiophyceae, Schizocladiophyceae, and Synchromophyceae have only been reported from marine environments (Andersen et al. 1993; Guillou et al. 1999; Andersen and Preisig 2002a; Kawachi et al. 2002a, b; Kawai et al. 2003; Horn et al. 2007; Kai et al. 2008; Han et al. 2018; Wetherbee et al. 2019). The Phaeophyceae is principally marine with only five genera thriving in freshwater (Bold and Wynne 1985; Kawai and Henry 2017); similarly, the Phaeosacciophyceae has a single freshwater genus (Graf et al. 2020b). Conversely, the Chrysophyceae, Phaeothamniophyceae, and Xanthophyceae are almost exclusively found in freshwater or soil environments (Ettl 1978; Reith 1980; Billard 1984; Starmach 1985; Hibberd 1990a; Ettl and Gärtner 1995; Bailey et al. 1998; Kristiansen and Preisig 2001; Andersen and Preisig 2002b; Kristiansen and Škaloud 2017; Maistro et al. 2017; Graf et al. 2020a). Finally, algae of the Diatomeae, Dictyochophyceae, Eustigmatophyceae, Olisthodiscophyceae, and Raphidophyceae inhabit equally freshwater or marine environments (Heywood 1990; Hibberd 1990b; Round et al. 1990; Moestrup 1995; Potter et al. 1997; Heywood and Leedale 2002; Moestrup and O’Kelly 2002; Eliáš et al. 2017; Horiguchi 2017; Mann et al. 2017; Barcytè et al. 2021). Despite being predominantly a group of photosynthetic organisms (i.e. autotrophy), an important number of Heterokontophyta are capable of phagocytosis and of uptaking organic molecules. Such ability to adapt their mode of nutrition is named mixotrophy and presents a wide spectrum (Mitra et al. 2016). Within the Heterokontophyta, the Chrysophyceae notably could well be entirely mixotrophs at some level. Furthermore, secondary loss of the chloroplasts is not unusual, especially in the mixotrophic lineages such as the Chrysophyceae (Rothhaupt 1996; Olrik 1998), Dictyochophyceae (Havskum and Riemann 1996;

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Table 5.1 Summary table of the general ecology and cell coverings of the Heterokontophyta. Fossils for which affiliation to the Heterokontophyta remains discussed are indicated in parentheses. The ecological niche in which algae are rarely found are put in parentheses Systematic group

Oldest accepted fossil

Ecological preferences

Cell covering

Diatomeae

190 Mya1 (or 570 Mya and older2)

Marine, freshwater, soil, snow

Siliceous frustules

Bolidophyceae

72 Mya2 (or 80 Mya2−3)

Marine

Naked, siliceous plates

Dictyochophyceae

113 Mya4

Marine, freshwater

Naked, organic scales, siliceous skeletons

Pelagophyceae

None

Marine

Naked, cell walls, thecae, gelatinous coverings

Chrysophyceae

228 Mya6 (or 811 Mya7)

Freshwater, soil (marine)

Naked, cell walls, organic lorica, organic or siliceous scales, gelatinous coverings

Synchromophyceae

None

Marine, freshwater

Naked, lorica

Eustigmatophyceae

None

Marine, freshwater

Cell walls

None

Marine

Naked, mineralized lorica, gelatinous coverings

SIII

SII

Incertae sedis Pinguiophyceae Olisthodiscophyceae

None

Marine, brackish

Scale covering

Chrysomeridophyceae

None

Brackish

?

Raphidophyceae

None

Marine, freshwater

Naked

Aurearenophyceae

None

Marine

Cell wall

Phaeothamniophyceae

None

Freshwater

Cell walls

SI

Phaeosacciophyceae

None

Freshwater (marine)

Cell walls

Chrysoparadoxophyceae

None

Marine soil

Cell walls

Xanthophyceae

(1000 Mya8)

Freshwater, brackish, soil

Naked, cell walls

Schizocladiophyceae

None

Marine

Cell walls with alginates

Phaeophyceae

99.6 Mya9 (or 550 Mya10)

Marine (freshwater)

Cell walls with cellulose and alginates

1

Rothpletz 1900. 2Siemińska 2015. 3Abe and Jordan 2021. 4Hajós and Stradner 1975. 5McCartney et al. 2014a. 6Zhang et al. 2016b. 7Allison and Hilgert 1986. 8Butterfield 2004. 9 Rajanikanth 1989. 10Xiao et al. 1998

Sekiguchi et al. 2003; Gerea et al. 2016), and Raphidophyceae (Jeong 2011). With the loss of plastids, the organisms exist by heterotrophy. Heterokontophyta has been extensively used by humans throughout history, not only for food consumption but also for their extracts and chemicals (e.g. Cosenza et al. 2017; Senthilkumar et al. 2017; Archer et al. 2019; Levasseur et al. 2020). A number of species of the Heterokontophyta (notably of the Pelagophyceae and Raphidophyceae) are known to produce blooms that can sometimes be harmful to fishes and humans (Gobler et al. 2005; Anderson et al. 2021). All these aspects of the Heterokontophyta ecology and its importance will be discussed for each class.

5.1.2 General Description The tremendous diversity of the Heterokontophyta makes it difficult to define characters shared across the entire group. These algae occur in a variety of shapes, forms, and sizes that make it impossible to prepare a brief summary. Furthermore, morphological variability occurs at the ultrastructural level within cells, and exceptions can be found for common trends. For example, most heterokonts have a girdle lamella within the chloroplast, but the girdle lamella is absent for members of the Eustigmatophyceae. Despite this apparent absence of unity, the Heterokontophyta are known to form, based upon molecular phylogenetic analyses, a

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monophyletic group (Leipe et al. 1994; Riisberg et al. 2009; Derrelle et al. 2016; Thakur et al. 2019), and some traits can be considered to be generally shared across the group.

5.1.2.1 Flagella The most distinctive trait and name-giving feature of the Heterokontophyta is probably swimming cells with two flagella of different lengths, i.e., a shorter mature flagellum and a longer immature flagellum. Furthermore, the long immature flagellum bears two rows of tripartite tubular hairs (= mastigonemes) (Fig. 5.1; Box 5.1). These hairs are formed of three parts (hence tripartite): a basal attachment to the flagellum region, a tubular shaft, and terminal fibrils (Fig. 5.3a–c) all composed of glycoproteins (Bouck 1971). The orientation of the flagella can vary from two forward-directed flagella (e.g. Synura) to one forward-directed flagellum and one trailing flagellum (e.g. Ectocarpus sperm). The long immature flagellum is mobile and beats following a sinusoidal mode. Its hairs, acting like oars, reverse the flagellum's thrust and therefore the flagellum pulls the cell forward (Sleigh 1974, 1989; Goldstein 1992). Such typical biflagellate cells are observed (at least during some life stage) in the Bolidophyceae, Chrysomerophyceae, Chrysoparadoxophyceae, Chrysophyceae, Eustigmatophyceae, Phaeophyceae, Phaeosacciophyceae, Phaeothamniophyceae, Raphidophyceae, Schizocladiophyceae, and Xanthophyceae, in some Pelagophyceae and in some Pinguiophyceae (Table 5.2). In the swimming sperm cells of centric diatoms and motile cells of the Dictyochophyceae, there is only a single immature flagellum; only the basal body of the mature flagellum is present. In the Pelagophyceae, Pelagomonas has an immature flagellum with only bipartite hairs, and the basal body of the mature flagellum is completely absent after cell division (see 5.1.4.8). Other peculiar cases, such as the hairless immature flagellum observed in the Pinguiophyceae Glossomastix and Polydochrysis (Fig. 5.11), demonstrate the vast diversity within the Stramenopiles. The flagella are fixed to the cell via the flagellar root apparatus, which may be composed of microtubular roots, striated roots, and transitional fibers (Andersen 1991, 2004). The organization and composition of the flagellar apparatus vary greatly among Heterokontophyta taxa, and flagellar structures were at the center of the taxonomic revisions during the second half of the twentieth century. 5.1.2.2 Chloroplast Another unifying trait of the Heterokontophyta is their chloroplast that originated from the endosymbiosis of a red algal derived endosymbiont (Sect. 4.2; Yoon et al. 2002; Le Corguillé et al. 2009; Dorrell et al. 2017; Dorrell and Bowler

T. Friedl

2017). With the exception of Chrysoparadoxa (Chrysoparadoxophyceae; 2.16), the chloroplast is encompassed by four membranes separating the stroma from the cytosol. From the innermost to the outermost, they correspond to the cyanobacterial membrane (1st and 2nd), the primary host plasma cell membrane (3rd), and finally, the secondary host phagocytic membrane, which is contiguous with the endoplasmic reticulum (Fig. 5.3a–e). The outer two membranes are likely the result of the secondary/tertiary endosymbiosis (see Chap. 2 and Fig. 2.6). Within the stroma of each chloroplast, the lamellae are formed by three closely stacked thylakoids. A specific lamella, named the girdle lamella (Fig. 5.3d–e), encloses all the other lamellae in the stroma as a sac in all classes except the Eustigmatophyceae (see Sect. 5.1.4.6 Eustigmatophyceae). • The primary function of the chloroplast is photosynthesis, and the Heterokontophyta possesses a wide range of light-harvesting pigments, the majority being photosynthetically active. These include the chlorophylls a, c1, c2, and c3 and a vast diversity of carotenoids (Table 5.2) (e.g. diatoxanthin, fucoxanthin) that give many Heterokontophyta their golden-brown color. • The photosynthesis product is a ß-1,3-linked glucan (20– 50 glucose residues) that is stored outside the chloroplast. For osmotic reasons, the storage molecules are contained within a vacuole. • For swimming cells, an eyespot may be present inside the chloroplast; in the Eustigmatophyceae, the eyespot is outside the chloroplast. The eyespot is associated with the mature flagellum to form the photoreceptor apparatus (Fig. 5.1), a cellular structure that is used to change the direction of swimming (i.e. phototaxis) by detecting the direction of the light source (Foster and Smyth 1980; Kawai and Kreimer 2000). Recent proteomics and transcriptomics analysis unveiled the role of a phototaxis protein named helmchrome (Fu et al. 2016). With the rapidly increasing number of fully sequenced plastid genomes, the gene content, genome structure, and organization, and their evolution in the Heterokontophyta is now under study (reviewed in Dorrell and Bowler 2017; see also Ševčíková et al. 2019; Kim et al. 2019, 2020; Starko et al. 2021).

5.1.2.3 Cell Coverings Cells of the Heterokontophyta often have distinctive and diverse coverings (Table 5.1). Roughly, these cell coverings can be divided into three large types: cell walls, silicified coverings, and gelatinous layers; the types are sometimes combined and there are important variations. Cell walls are common in the SI clade (except Raphidophyceae) as well as

+

+

+

Dictyochophyceae

Pelagophyceae

– +

–

–

Synchromophyceae

Eustigmatophyceae

+

+

?

Olisthodiscophyceae

Chrysomeridophyceae

+

Phaeophyceae

+

+

+

Chrysoparadoxophyceae

+

+

Phaeosacciophyceae

Xanthophyceae

+

+

Phaeothamniophyceae

Schizocladiophyceae

+

–

Aurearenophyceae

+

+

?

+

–

±

Raphidophyceae

SI ±

?

+

Pinguiophyceae

±

Incertae sedis-

±

+

+

+

+

±

Plastid-nucleus membrane connected

Chrysophyceae

SII-

+

Bolidophyceae

Girdle lamella

Diatomeae

SIII

Taxon

+ ±

± ±

±

±

2 (meiospores, sperm)

2, 0 2 (zoospores)

± –

±

2

+

+

2

2 (zoospores)

+

2

–

2

1

2

2, 1, 0

2, 0

0

2, 1

2, 1, 0

1

2, 0

0, 1

# flagella

+

+

+

+

+

+

+ –

– – –

+

±

±

–

+ (outside)

–

±

±

±

+

–

Pyrenoid

–

Eyespot

a, c1, c2

a, c (type ?)

a, c1, c2

a, c1, c2

?

a, c1, c2

a

a, c1, c2

?

a, c1, c2

a, c1, c2

a

a, c2

a, c1, c2

a, c1, c2

a, c1, c2

a, c1, c2, c3

a, c1, c2, c3

Chlorophylls

–

+

–

+

?

+

+

±

?

+

+

–

+

+

+

+

+

+

Fucoxanthin

–

?

–

+

?

+

? +

?

–

+

±

?

+

+

+

+

+

–

–

–

–

Violaxanthin, Zeaxanthin

–

?

+

±

–

?

–

–

–

–

–

+

+

+

Diatoxanthin, diadinoxanthin

Table 5.2 Summary table of the morphological and ultrastructural characters and pigments of the Heterokontophyta. The eyespot of the Eustigmatophyceae is placed outside the chloroplast. Life stages during which the flagella are present are indicated in parenthesis, if not the flagella are present during the entire life span. The type of chlorophyll c present in the Schizocladiophyceae is unknown. + = present; - = absent; ± = present; ? = unknown

5 Algae from Secondary Endosymbiosis 225

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the Chrysophyceae, Eustigmatophyceae, and Pelagophyceae. The wall composition likely varies greatly between lineages, but in many lineages their composition is still poorly studied (i.e. Aurearenophyceae, Chrysoparadoxophyceae, Phaeosacciophyceae, Phaeothamniophyceae, Pelagophyceae, and Xanthophyceae). One recent analysis suggests that they could be predominantly composed of cellulose (Okuda et al. 2004). The most fully studied is the cell wall of the Phaeophyceae, impregnated by alginates and cellulose and with a connection through plasmodesmata (see 5.1.4.1; Deniaud-Bouët et al. 2014, 2017; Kawai and Henry 2017). In the Schizocladiophyceae, the cell wall is comparable to that of the Phaeophyceae but lacks cellulose and plasmodesmata (Kawai et al. 2003). The cell wall of the Eustigmatophyceae appears to be single-layered and principally composed of cellulose (Okuda et al. 2004), but some taxa present bilayer cell wall with a more complex composition (Scholz et al. 2014). While cell walls completely surround the protoplasm and provide mechanical strength so that osmotic pressure may form within the cell, there are other cell coverings that do not completely surround the protoplasm. For example, loricas are common among the Chrysophyceae. The lorica consists largely of a wall-like covering but with one or more openings within the covering. As a consequence, the lorica provides protection but does not allow osmotic pressure; freshwater loricate cells typically have contractile vacuoles to maintain cellular osmolarity. The lorica of Dinobryon has a vase-like morphology and the flagella extend out through the open lorica mouth (Fig. 5.10). The lorica is composed of fibrils of cellulose and/or chitin interwoven together (Herth et al. 1977). The lorica of Epipyxis also has a vase-like morphology, but the lorica is formed from overlapping organic scales. The lorica of Lagynion is flask-shaped, and like Dinobryon, consists of a continuous cellulose-based layer; however, Lagynion has a very narrow pore opening and a pseudopod extends out through the pore. As in other eukaryotic groups (Knoll 2003; Finkel 2016), silica (silicon oxide SiO2) is incorporated in many Heterokontophyta, i.e. the silica frustule of diatoms, the silica scale coverings in the Bolidophyceae and some Chrysophyceae (notably the order Synurales), and silica skeletons in the Dictyochophyceae (Table 5.1). The silicified frustules (see 5.1.4.4) of the diatoms are probably the most famous and were even the center of an art form in the late 1800s. In the Bolidophyceae (see 5.1.4.11), the silica scales are large interlocking plates that cover the cell. Within the Chrysophyceae (see 5.1.4.2), the order Synurales are all covered by finely sculptured, bilaterally symmetrical, and overlapping silica scales positioned in what appear to be spiral rows. Other Chrysophyceae have radially symmetrical scales that do not form into tight scale cases. The

T. Friedl

Chrysophyceae also produce silica-resting cysts. Chrysophyceae scales and cysts are formed by the precipitation of silica into deposition vesicles shaped into the form of the final structure (Preisig 1994; Hildebrand et al. 2018). Finally, the silicoflagellates of the Dictyochophyceae (see 5.1.4.7) produce an external star-shaped skeleton, and the cell protoplasm is held in this basket-like skeleton. In summary, a typical swimming cell of the Heterokontophyta is characterized by two flagella of unequal length: a long immature flagellum with two rows of tripartite hairs and a short smooth mature flagellum. The chloroplast is enclosed in four membranes, with the outermost membrane continuous with the endoplasmic reticulum/nuclear envelope. Inside the chloroplast, the lamellae are enclosed by a sac-like girdle lamella (Table 5.2). Other typical features of the Heterokontophyta include mitochondria with tubular cristae (Taylor 1976; Stewart and Mattox 1980), Golgi bodies anterior to the nucleus with cis-cisternae adjacent to the nuclear envelope (e.g. Hibberd 1976).

5.1.3 Evolutionary History The Stramenopiles are now widely understood to form a supergroup with the Alveolata and Rhizaria (SAR; see Opuntia of life Fig. 2.8), and now the Telonemia [Burki et al. 2020, forming the TSAR]). Within the Stramenopiles, the Heterokontophyta form a monophyletic group sister to non-photosynthetic lineages (Leipe et al. 1994; Derrelle et al. 2016; Thakur et al. 2019). The divergence of the Heterokontophyta within the Stramenopiles was estimated by molecular clock analysis to have happened between c. 898 Mya (Parfrey et al. 2011) and c.384 Mya (Berney and Pawlowski 2006). The latest phylogenomic analysis of the eukaryotes placed the divergence of the Heterokontophyta between 1298 and 622 Mya, considerably pushing back the previous estimates (Strassert et al. 2021). The mineralized cell coverings of some Heterokontophyta provide a fossil record for a limited number of lineages. The silicified cysts and scales of the Chrysophyceae are common in geological deposits (e.g. Tappan 1980; Allison and Hilgert 1986; Riaux-Gobin and Stumm 2006; Siver et al. 2015; Zhang et al. 2016b). Similarly, diatom frustules (Sims et al. 2006; Harwood et al. 2007), plates of Bolidophyceae (Stradner and Allram 1982; Konno et al. 2007; Abe and Jordan 2021), and external skeletons of some Dictyochophyceae (Deflandre 1950; Perch-Nielsen 1985; McCartney 2013, 2014; McCartney and Witkowski 2016) have been recorded, dating back to the Cretaceous Period (145–66 Mya). Therefore, the fossil record for silica structures is essentially much younger than the molecular clock estimations (Brown and Sorhannus 2010). This incongruity could be explained by the instability

5

Algae from Secondary Endosymbiosis

over time of the mineral composing the coverings once in sediment. Notably, silicified coverings (e.g. diatom frustules) can convert to porcelanite and then to chert (Calvert 1977) in a process that ultimately destroys the fossil signal of the original frustule. Furthermore, it remains entirely possible that older fossils exist, notably in the diatoms for which fossils older than 570 Mya have been reported from Poland and are currently being discussed (Siemińska 2015). Fossils of the Phaeophyceae and Xanthophyceae are very rare and scarce, while fossils of other classes have not been reported at all (Table 5.1). Despite their rarity, these fossils are of particular importance. The earliest fossil assigned to the Heterokontophyta is Paleovaucheria, a Xanthophyceae found in a geologic formation c.1000 Mya (Hermann 1981; Butterfield 2004). Because of the lack of fossils, molecular phylogenetic relationships have supplied the principal insights into the evolutionary history of the Heterokontophyta. After attempts at cladistic analyses (e.g. Hibberd 1979; Williams 1991), beginning in the 1990s taxonomists turned toward the promises of molecular phylogenetics. Importantly, they revealed the monophyly of the photosynthetic lineages within the Stramenopiles (Blackwell and Powell 2000; Moriya et al. 2002; Riisberg et al. 2009; Beakes et al. 2014). However, even if these early analyses revealed consistent and clear relationships between the photosynthetic lineages (reviewed in Andersen 2004), an overall phylogeny of the Heterokontophyta remained unresolved. Recently, this situation started to clear with the analysis of a large set of taxa covering most of the variety of the Heterokontophyta (Yang et al. 2012) and/or phylogenomic analyses (Derelle et al. 2016; Kim et al. 2019; Thakur et al. 2019). Through these analyses, a consensus phylogeny emerged that divides the Heterokontophyta into three large clades, named SI, SII, and SIII (Fig. 5.1; Yang et al. 2012). Despite this advancement, relationships between and within those clades are not completely resolved and will surely be at the center of future phylogenomic analyses. The SI clade (Fig. 5.2 green) regroups lineages with (a) unicellular species (Aurearenophyceae, Chrysoparadoxophyceae, and Raphidophyceae), (b) multicellular species (Phaeosacciophyceae, Phaeothamniophyceae, Schizocladiophyceae, and Xanthophyceae), and (c) complex multicellularity as found in the Phaeophyceae. The SII clade (Fig. 5.2 red) regroups principally colonial and unicellular species (Chrysophyceae, Eustigmatophyceae, and Synchromophyceae). In this clade, mixotrophy is common notably within the Chrysophyceae, and non-photosynthetic species are heterotrophic. The SIII clade (Fig. 5.2 blue) regroups lineages (Diatomeae, Bolidophyceae, Dictyochophyceae, and Pelagophyceae) with reduced flagellar apparatuses or without flagella completely, with the notable and puzzling exception of the Pelagophyte order Sarcinochrysidiales.

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Despite the lack of strong support in the deep nodes of the phylogenetic tree of the Heterokontophyta (i.e. branching order between the three clades), a general evolutionary history of various traits can be suggested. Both the molecular phylogenies and the fossil record support an oceanic origin of the Heterokontophyta, with multiple independent transitions to freshwater and/or land. The loss of one flagellum, or even both flagella, in the SIII clade is a derived state; i.e. the biflagellate condition is widespread among both photosynthetic and non-photosynthetic Stramenopiles as well as protists in general. Similarly, it is now well accepted that a high endosymbiotic event (secondary maybe tertiary) gave rise to the chloroplast of the Heterokontophyta. That is, analysis of the genomes of oomycetes and labyrinthulomycetes (heterotrophic Stramenopiles lineages) did not reveal the enrichment of genes as remnants of an ancient endosymbiosis (Stiller et al. 2009, 2014; Dorrell et al. 2017), but studies have identified a handful of genes potentially inherited horizontally (Andersson and Roger 2002; Robertson and Tartar 2006). These analyses support a chloroplast origin in the common ancestor of the photosynthetic Stramenopiles (= common ancestor of the Heterokontophyta) rather than in the ancestor of all Stramenopiles (i.e. not the Chromalveolate hypothesis; Cavalier-Smith 1999). Phylogenetic analysis of chloroplast encoded genes generally places the Heterokontophyta sister to the red algae (Le Corguillé et al. 2009; Dorrell et al. 2017), suggesting that the endosymbiont was a red alga. Therefore, the chloroplast of the Heterokontophyta has been largely viewed to have originated from a single secondary endosymbiosis with a red algal endosymbiont that took place in the common ancestor of the photosynthetic lineages (see Chap. 2, Endosymbioses). However, recent analyses have started to challenge this view and a more complex evolutionary scenario may be emerging (Bodył et al. 2009; Stiller et al. 2009; Strassert et al. 2021). These studies suggest that the chloroplast of the Heterokontophyta was the result of a tertiary endosymbiosis with a Cryptophyte endosymbiont. Furthermore, genomics analysis revealed that both red algae and green algae contributed importantly to the genome composition of the Heterokontophyta, either through lateral gene transfer or through a cryptic endosymbiosis (Moustafa et al. 2009; Dorrell et al. 2017; Morozov and Galachyants 2019; Sibbald and Archibald 2020). The clear chain of events leading to the establishment of the chloroplast and the genomic mosaic of Heterokontophyta remains the subject of current debate and ongoing research. In various lineages, the chloroplast was secondarily lost, further demonstrating the dynamic evolution of the Heterokontophyta. In some cases (e.g. Ciliophrys,

BROWN 1945 1st electron microscopy of Heterokontophyta

1914 - 1950 FLORISTIC PERIOD

PASCHER 1914 first synthesis stops phylogenetic studies

1882 - 1914 FIRST SYNTHESIS

BLACKMAN 1900 important phylogeny of Heterokontophyta CHADEFAUT 1950 return of phylogenetic aspects

GUNDERSEN et al. 1987 1st molecular phylogeny of Heterokontophyta

BAILEY et al. 1998 Phaeothamniophyceae

CAVALIER-SMITH et al. 1995 Chrysomeridophyceae

separation of Synurophyceae

ANDERSEN et al. 1993 Pelagophyceae

1950 - 2004 PHYLOGENETIC PERIOD

HIBBERT & LEEDALE 1970 Eustigmatophyceae

ANDERSEN 1987

SILVA 1980 Dictyochophyceae

ARMBRUST et al. 2004 1st genome sequence of Heterokontophyta

COCK et al. 2010 1st genome sequence of brown alga

KEI et al. 2008 Aurearenophyceae

YANG et al. 2012 introduction of the three subclades of Heterokontophyta

WETHERBEE et al. 2019 Chrysoparadoxophyceae

GRAF et al. 2020 Phaeosacciophyceae

BARCYTÈ et al. 2021 Olisthodiscophyceae

2004 - present day GENOMIC PERIOD

KAWAI et al. 2003 Schizocladiophyceae

KAWACHI et al. 2002 Pinguiophyceae

HORN et al. 2007 Synchromophyceae

(including Synurophyceae)

Chrysophyceae: ~1,180 spp. (~216 gen.)

Pelagophyceae: 30 spp. (14 gen.)

Dictyochophyceae: ~160 spp. (19 gen.)

Bolidophyceae: 18 spp. (3 gen.)

Diatomeae: ~16,700 species (in ~1.050 genera)

Phaeosacciophyceae: 10 spp. (4 gen.)

Phaeothamniophyceae: 35 spp. (15 gen.)

Aurearenophyceae: 1 sp. (1 genus)

Raphidophyceae: 40 spp. (16 gen.)

Chrysomeridophyceae: 1 sp. (1 genus)

Olisthodiscophyceae: 2 spp. (1 genus)

Pinguiophyceae: 5 spp. (5 gen.)

Eustigmatophyceae: ~100 spp. (18 gen.)

Synchromophyceae: 7 spp. (2 gen.)

Phaeophyceae: ~2.060 spp. (~300 gen.)

Schizocladiophyceae: 1 sp. (1 genus)

Xanthophyceae: ~700 spp. (97 gen.)

Chrysoparadoxophyceae: 1 sp. (1 genus)

Fig. 5.2 Chronological history of knowledge on the Heterokontophyta. The time scale is shown in log scale. Dashed arrows represent separation and re-assignment of taxa to a newly described class. The classes are colored based on the generally accepted phylogeny of the Heterokontophyta shown on the right side. Green: clade SI; red: clade SII; blue: clade SIII; gray: classes for which the position in the phylogeny remains undetermined (dashed branches in the phylogenetic tree)

1753 - 1882 DISCOVERY PERIOD

ROSTAFINSKI 1882 1st phylogeny of the Heterokontophyta

LINNAEUS 1753 description of Fucus 1st Heterokontophyta

de CANDOLLE 1801 Xanthophyceae

DIESING 1865 Raphidophyceae

MÜLLER 1773 1st description of Chrysophyceae

MÜLLER 1783 1st description of Diatomeae

Heterokontophyta S III Heterokontophyta S II Heterokontophyta S I

Heterokontophyta: S III

Heterokontophyta: S II

Heterokontophyta: S I

GUILLOU et al. 1999 Bolidophyceae

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Cornospumella, Nitzschia, Pedospumella, Poteriospumella, and Pteridomonas), colorless vestigial chloroplasts, named leucoplasts, are present (Sekiguchi et al. 2002; Kamikawa et al. 2015; Grossmann et al. 2016), but these species are free-living heterotrophs that feed through osmotrophy or phagotrophy. These secondary losses appear to be especially frequent in the Chrysophyceae (see Bütschli 1883–87), a

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class in which mixotrophy is also frequent and in which many species lack carbon anhydrase (Raven 1995, 2010; Maberly et al. 2009; Raven and Giordano 2017). Recently, genomic investigations revealed important and converging nuclear genome reduction (Dorrell et al. 2019; Majda et al. 2019) as well as photosynthetic-related genes lost from the plastid genome (Kim et al. 2020).

Box 5.1 Names Throughout history, numerous names were used to describe lineage of the photosynthetic stramenopiles, and therefore it can be confusing while browsing the large bibliography concerning these algae. Furthermore, these names were used to describe photosynthetic organisms before there was a need to accommodate the non-photosynthetic Stramenopiles. Currently, the two most commonly used names are stramenopiles and heterokonts. The name Heterokontae (Greek hetero—different + kontos—pole) refers to the two flagella of unequal length (Fig. 5.1). The name was first used by Luther (1899) to classify yellow-green freshwater algae that are now placed in the classes Xanthophyceae and Raphidophyceae. The name evolved and extended to include more algal classes and became the division Heterokonta (Cavalier-Smith 1978) or Heterokontophyta (van den Hoek 1978). The taxonomic division, regardless of spelling, included only photosynthetic organisms and the colorless species resulting from the loss of photosynthesis, i.e. it did not include the heterotrophic Stramenopiles such as the Oomycetes and thraustochytrids. However, it was already apparent that scientists considered the photosynthetic Stramenopiles to be a group containing several taxonomic divisions, e.g. Bacillariophyta, Phaeophyta, and Xanthophyta. Therefore, during the 1980s, the nontaxonomic name “heterokont” progressively replaced the division names and the non-photosynthetic lineages were sometimes included. Despite the existence of other groups of algae presenting flagella of unequal length, like the Dinoflagellata (Sect. 5.2), the term heterokont is specifically applied to those algae with predominantly one short smooth flagellum and one long hairy flagellum, i.e. mature and immature flagellum, respectively (Fig. 5.1). The name stramenopiles was proposed by Patterson (1989), and it was based on the presence of tripartite tubular hairs on a flagellum (Latin, stramin—straw and pila—hairs); see Fig. 5.1. This name has become widely used, and although Patterson originally intended the name to be without taxonomic rank, today the name often has taxonomic rank significance and is spelled with a capital S (Stramenopiles). The names Chromophycés (Chadefaud 1950), Chromophycota (Chadefaud 1960), Chromophyta (Christensen 1962), and Chromophyta (Bourrelly 1968) were proposed to describe (with variations and in different combinations) groups of algae that included the heterokonts/stramenopiles. However, they also included algal groups that are distinctly different, e.g. the Dinoflagellata (Sect. 5.2), Cryptophyta (Sect. 5.5), and Haptophyta (Sect. 5.6). These names eventually evolved into the name Chromista (Cavalier-Smith 1981) to regroup the chlorophyll c containing algae only (i.e. Cryptophyta, Haptophyta, and Heterokontophyta). Even more recently, the name Chromalveolata appeared in the literature (e.g. Adl et al. 2005). Today, these names are not widely used because the chlorophyll c containing algae have been shown to be polyphyletic. Finally, within the Chromista and Chromalveolata, the photosynthetic Stramenopiles lineage was given the name Ochrista (Cavalier-Smith 1986) that, in turn, evolved into Ochrophyta (Cavalier-Smith and Chao 1996) or the stramenochromes (Leipe et al. 1994). Here, we use the less confusing names Heterokontophyta and photosynthetic Stramenopiles.

5.1.4 Taxonomic Classes This section contains detailed information that is specific to each taxonomic class. The classes are arranged in chronological order of their descriptions.

5.1.4.1 Phaeophyceae The macroalgae of the class Phaeophyceae form a clear and intuitively distinct lineage of the Heterokontophyta. Named after their typical coloration (Greek phaeo - gray, dusky), they are generally referred to as brown algae, in opposition to the red and green algae.

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Brief history and current taxonomy Brown marine seaweeds have long been used by human populations for various purposes, largely predating any scientific work on them. Logically, they were some of the very first described Heterokontophyta species (Fucus by Linnaeus 1753), and the first formally defined class of Heterokontophyta was the Fucophyceae (Warming 1884) later renamed Phaeophyceae (Kjellman 1891). Since then, the distinction of this class was never seriously questioned; most of the discussions revolved around their affiliation with other Heterokontophyta like the Xanthophyceae, Diatomeae, and Chrysophyceae (Blackman 1900; Pascher 1914; Papenfuss 1955). Many important studies on their morphology, development, and reproduction were conducted during this period (reviewed by Fritsch 1935). Recently, reports of the genome sequences of multiple Phaeophyceae started to bring new insights into their biology and physiology (Cock et al. 2010; Ye et al. 2015; Nishitsuji et al. 2016; Nishitsuji et al. 2019; Wang et al. 2020; Graf et al. 2021). Furthermore, these sequences paved the way for genomics and transcriptomics studies involving sexual life cycle evolution (e.g. Lipinska et al. 2017, 2019), gene expression in response to stresses (e.g. Dittami et al. 2009; Machado Monteiro et al. 2019; Rugiu et al. 2020), biogeography (Starko et al. 2019; Bringloe et al. 2020b), and more generally on the evolution of the Phaeophyceae (see Bringloe et al. 2020a for a comprehensive review of the recent progress). There are currently 19 orders and over 2,000 species recognized in the class Phaeophyceae, with the majority of the diversity contained in four orders: the Dictyotales, Ectocarpales, Fucales, and Laminariales (Fig. 5.4). General description Phaeophyceae are typical members of the Heterokontophyta in terms of cell structure and biology but possess other distinguishing traits. The class regroups strictly and truly multicellular species (i.e. no colonial species), which range from filamentous to pseudoparenchymatous (aggregated filaments) to parenchymatous organisms. The thalli have very diverse morphologies (Fig. 5.4). Some species can be found as simple filamentous tufts, but most species have simple and branched thalli with a differentiated holdfast, a stem-like stipe, and a foliose blade, itself presenting various morphologies. Holdfasts (also called haptera) serve to anchor the algae to their substrates (often rocky seashores) by the mean of cellular growth (directed by negative phototropism; Buggeln 1974), completely occupying the substrate space (Tovey and Moss 1978). The sexual reproductive structures are mostly differentiated from the blade tissue, but they can

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also be found on the stipe (e.g. Undaria) or the holdfast (e.g. Aureophycus). Because of their multicellular thalli, an important aspect of the cellular structure is their cell walls. Walls are mostly composed of cellulose, fucose-containing sulfated polysaccharides, and alginates (Deniaud-Bouët et al. 2014, 2017). Other wall components include proteins, glycoproteins, halogenated and/or sulfated phenolics known as phlorotannins (Meslet-Cladière et al. 2013), halides such as iodide (Verhaeghe et al. 2008), and various ions (Deniaud-Bouët et al. 2014, 2017; Terauchi et al. 2016). Fucose-containing sulfated polysaccharides can be a different form of homopolymers (i.e. fucans) or heteropolymers (i.e. fucoidans), and species-dependent structural variations exist. Alginates are linear polymers of mannuronate and guluronate molecules. The relative composition of mannuronate and guluronate in the chains can affect the chemical properties of the alginates, which likely affect cell wall rigidity and texture (Draget et al. 2005; Deniaud-Bouët et al. 2014). The halide metabolism of the brown algae (especially the Laminariales) was shown to play an important ecologically defensive role through the accumulation of halides in the cell wall (Küpper et al. 2008; Leblanc et al. 2006; La Barre et al. 2010). In Phaeophyceae, the protoplasm of the cells is interconnected by plasmodesmata (e.g. Terauchi et al. 2015, Fig. 5.5). Chloroplasts are typical, with four membranes and the outermost being the chloroplastic endoplasmic reticulum continuous with the nuclear envelope. The thylakoids are three stacked and a girdle lamella encloses them. A pyrenoid is commonly present, but it may be absent during certain life stages or totally absent in some species. Photosynthetic pigments include chlorophyll a, c1, and c2, as well as the accessory pigments ß-carotene, violaxanthin, and important amounts of fucoxanthin. The photosynthetic reserve product is a ß-1,3-glucan named laminarin, which is similar to chrysolaminarin, and cells also contain large quantities (up to 25% of the dry weight) of D-mannitol, a 6-carbon sugar alcohol (Schmitz 1981). The metabolism of the Phaeophyceae (minus the Fucales) is unique for their mechanism that increases the uptake of inorganic carbon when cells are illuminated with blue light (Forster and Dring 1994). This allows for tuning between the increased inorganic carbon uptake and photosynthetic carbon fixation (i.e. only when the cells are illuminated), when the energy necessary for the inorganic carbon uptake is saved when the cells are not able to photosynthesize (i.e. while in the dark). Plasmodesmata are straight, plasma membrane-lined channels of 10–20 nm diameter that pass through the cell walls of adjacent cells (Fig. 5.5a–b). Plasmodesmata are established during cytokinesis and in many species with a complex thallus; they can be observed in groups that are

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b Fig. 5.3 Transmission electron micrographs of the Heterokontophyta. a Immunolabeled whole cell with the immature hairy flagellum (AF) and

the mature smooth flagellum (PF) of the zoospore of Scytosiphon lomentaria. b Detail of the immature flagellum of the zoospore of S. lomentaria showing the mastigonemes. c Negatively stained mastigoneme of S. lomentaria showing the terminal filament (T), the tubular shaft (S), and the base (B). d Plastid of Phaeodactylum tricornutum showing the girdle lamella (G), thylakoids (T), pyrenoid (P) and oil droplets (OB), the nucleus (N), and mitochondrion (Mito). e Detail of a plastid of P. tricornutum showing the four extraplastidial membranes (Mem). a–c modified from Honda et al. (2007); d–e modified from Dorrell and Bowler (2017)

named a “pit field” (Terauchi et al. 2015). Another distinctive feature of the Phaeophyceae is the cytoplasmic organelle called the physode (Fig. 5.5c). Noticeable in light microscopy due to their refraction, they contain phlorotannins (polymers of phloroglucinol) produced in the Golgi apparatus and endoplasmic reticulum in the periphery of the cell (Schoenwaelder 2002; Pavia et al. 2003). Oxidation of the phlorotannins turns them into phycophaein, a dark pigment that gives to dried Phaeophyceae their characteristic black color. Various functions have been proposed for physodes (Shibata et al. 2002a, b; Henry and Van Alstyne 2004; Halm et al. 2011; Nakajima et al. 2015), but generally they are thought to be a constitutive (i.e. always present) defense (Flöthe et al. 2014). Also, the physode may secrete phlorotannins by exocytosis, which in turn blocks polyspermy after fertilization (Clayton and Ashburner 1994). Laminariales are remarkable within the Phaeophyceae for their specialized cell types. The level of differentiation varies depending on the localization and the age of the cell (Ducreux 1984; Gall et al. 1996). Generally, the blades contain three types of cells that are differentiated from the outside to the inside. The specialized cells include mucilage-secretion cells, physode accumulating cells, and trumpet hyphal cells (Fig. 5.5d–e). The last cell type forms vascular tissue in the center of the stipe, which allows for long-distance transport and response (Raven 2003; Drobnitch et al. 2015) as well as potentially innate immunity (Flöthe et al. 2014; Thomas et al. 2014). Growth localization and axes vary between the different lineages of the Phaeophyceae. Some species do not have a meristem and have a diffuse growth, but in a majority of species, a meristem is present. Different types of meristems can be found, from a simple single apical dividing cell to more complex intercalary meristems to the complex meristoderm found in kelps (reviewed in Charrier et al. 2012). Sexual reproduction is widespread and obligatory in most species. The life cycle is characterized by the alternation of generations between a macroscopic diploid sporophyte and microscopic halploid gametophytes. The different species of Phaeophyceae present specificities in their life cycle that have been reviewed in detail (Lee 2018), but generally two types of life cycles can be recognized. The kelps of the order Laminariales are good examples of the oogamous diplo-haplontic life cycle (Fig. 5.6). The large kelp thalli, which are easily observed on the temperate coastlines, are the diploid sporophyte form of this cycle. Upon

maturation, a specific region (e.g. on the blade in Laminaria and Saccharina, the stipe in Undaria or the holdfast in Aureophycus) forms a specialized tissue named the sorus (plural = sori). It is formed by a dense concentration of unilocular sporangia in which meiospores are formed by meiosis. In each unilocular sporangium, large number of meiospores are formed (e.g. 32 in Laminaria, Motomura et al. 1997) and released through the apex of the sporangium. The meiospores are typical biflagellate cells with a single chloroplast and an eyespot is present in some species. Swimming ability varies depending on the species. Dispersal can be local or up to a few kilometers (Santelices 1990; Gaylord et al. 2004; Bobadilla and Santelices 2005). The meiospores settle on a substrate and by mitosis develop into a gametophyte. The gametophytes are dioecious with separate male and female morphologies. The male gametophytes are generally more branched and have smaller cells when compared to the female. The male gametophyte produces antheridia at the extremities of its branches, and cells within each antheridium differentiate into spermatozoids. The spermatozoids are biflagellate, unicellular cells typical of the Heterokontophyta. The female gametophyte produces elongated oogonia that differentiate, with each forming a single egg. This difference in the morphology of the gametes (i.e. spermatozoid and egg) defines this cycle as oogamous. Once fully formed, the eggs secrete a sexual hormone named lamoxirene that triggers the release of the spermatozoids and by positive chemotropism attracts them to the egg (Motomura 1991). Fertilization of the egg takes place on the female gametophyte and results in a zygote that germinates to first form a flat embryo and then subsequently to form the sporophyte. In the order Fucales, the sexual reproduction does not necessitate the formation of separate gametophyte thalli (Fig. 5.7). The gametes are formed in specialized conceptacles that are localized on the blade of the sporophyte. The conceptacles are multicellular structures with cells forming the floor and walls and an opening to the outside (ostiole). Depending on the species, the thalli of the Fucales are monoecious or dioecious; for monoecious species, the conceptacles enclose both antheridia and oogonia. In the antheridia, gametes are formed by meiosis followed by multiple mitotic divisions (e.g., four in Fucus) before their differentiation into spermatozoids (e.g., 64 per antheridium in Fucus). When the antheridia mature, their cell wall breaks and releases the spermatozoids (sperms) in a mucilage that dissolves in the seawater. The oogonia are supported by a

Fig. 5.4 Diversity of the Phaeophyceae. a Fucus vesiculosus (Fucales). b Hormosira banksii (Fucales). c Cystoseira tamariscifolia (Fucales). d Bifurcaria bifurcata (Fucales). e) Colpomenia sp. (Fucales). f Undaria pinnatifida (Laminariales). g Nereocystis luetkeana (Laminariales). h Yellowish thalli of Durvillea antarctica (Fucales) and Sargassum sp. (Fucales). i Himanthalia sp. (Fucales), young thalli. j Ascophyllum nodusum (Fucales). k Dictyota dichotoma (Dictyotales). l Saccharina latissima (Laminariales). All pictures were kindly provided by Burkhard Büdel

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100 nm

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Fig. 5.5 Multicellularity of the Phaeophyceae. a Longitudinal section of Saccharina japonica showing plasmodesmata between cortex cells. Arrowheads = membranous structures near the plasmodesmata. cw = cell wall. b Transverse view of a pit field in S. japonica. c Ultrastructural organization of the cells in Pylaiella littoralis showing physodes (black vacuoles in the periphery of the cell). d–e Longitudinal section of Nereocystis luetkeana showing conducting cells. H = Hyphal cell, S = sieve element, arrows = chloroplasts, empty arrow = callosed cell. Enlargement
1050. a–b from Terauchi et al. 2015; c from Markey and Wilce 1975; d–e from Schmitz and Srivastava (1976)

stacked cell of the conceptacle’s wall. They undergo three nuclear divisions before the cytoplasm cleaves, resulting in eight gametes. The cell wall of the oogonia is formed of three layers, i.e. the exochite, mesochite, and the endochite (from the outermost to the innermost). When mature, the exochite breaks and releases the eight eggs that are still contained in the mesochite and endochites, which will dissolve in seawater. Spermatozoids are attracted to the egg by a pheromone, fucoserratene, secreted by the eggs. Fertilization of the egg by the spermatozoid forms a zygote that will develop into a diploid sporophyte. Occurrence The Phaeophyceae are commonly found in coastal regions throughout the world, and they perform fundamental ecological roles in the intertidal and shallow subtidal ecosystems (Steneck et al. 2002; Schiel and Foster 2006; Mineur et al. 2015; Teagle et al. 2017). Kelp forests are so dense and important in temperate oceans that they impressed Darwin during his travel around the world aboard the Beagle

(Darwin 1909). Kelp forests are ecologically important because they provide nursery grounds for other organisms (Holbrook et al. 1990; Kitada et al. 2019); they form habitats for many organisms (Steneck et al. 2002; Graham 2004; Markel et al. 2017), and they increase ecosystem complexity (e.g. Gattuso et al. 2006; Stephens and Hepbrun 2014; Estes et al. 2016). In addition, kelps are excellent primary producers that are able to attain productivity rates of 1 kg C m−2 yr−1, comparable to that of tropical rain forests (Mann 1973; Pfister et al. 2019). Furthermore, kelps support secondary productivity through herbivory and the production of detritus (Duggins et al. 1989; Krumhansl and Scheibling 2012). The highest species diversity of Phaeophyceae is observed in the cold temperate regions. In these regions, the rocky intertidal zone is dominated by the Fucales, while the Laminariales dominate the shallow subtidal zone. One of the most emblematic ecosystems formed by the Phaeophyceae are the kelp forests (Fig. 5.8). Composed principally of the large (up to 50 m) Laminariales genera (e.g. Ecklonia, Macrocystis, Laminaria, Saccharina, Undaria), kelp forests can also be formed by large members of the Ascoseirales

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unilocular sporangium m

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Fig. 5.6 The life cycle of the order Laminariales with heteromorphic alternation of generations. Meiosis in a unilocular sporangium on the sporophyte leads to haploid meiospores. The sporangium segregates sex-determining genes, such that meiospores develop into unisexual (dioecious) gametophytes. The gametophytes are microscopic, branched filaments (male and female gametophytes are different by themselves in morphology) and, therefore, rather different from the dominant and large sporophyte stage. There are male gametophytes that produce antheridial cells each giving rise to a single sperm. On the female gametophytes, some (egg-producing) cells release wallless content which function as eggs; the egg-producing cells may be regarded as oogonia. The eggs often remain attached to their former cell wall on the female gametophyte. A pheromone, secreted by the eggs, attracts the sperms. Subsequently, fertilization and development of the zygote into the sporophytes occur (see also Fig. 2.9c). Original drawing Spindler & Friedl

(e.g. Ascoseira), Desmarestiales (e.g. Himmantothallus), Fucales (e.g. Durvillaea), or Tilopteridales (e.g. Sacchoriza and Phyllariopsis). Kelps are restricted to cold waters (generally with an upper temperature limit of 18–19 °C), but the composition of their ecosystems varies in different

regions. In the polar regions, the Laminariales Chorda is abundant in the Arctic but absent in the Antarctic, which is dominated by Desmarestiales. In the Atlantic Ocean, kelp forests are composed of the Laminariales Alaria, Laminaria and Saccharina (Northern Atlantic), and Ecklonia (South

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Fig. 5.7 The diplontic life cycle of the order Fucales. Gametes are generated by gametic meiosis within gametangia produced in branch termini of the sporophyte. There is no alternation of generations. In the example shown here, both types of gametangia, the antheridium (male) and oogonium (female), are produced on the same sporophyte body (monoecious species) within chambers, known as conceptacles. Meiosis directly leads to the formation of gametes, i.e. sperms and egg cells. Gamete discharge into seawater is through the open pore (ostiole) of the conceptacle. The sperms are attracted to eggs by a pheromone that is released by the eggs. Fertilization occurs in the water; the zygotes then attach to substrates where they further develop into sporophytes (see also Fig. 2.9b). Original drawing Spindler & Friedl

Africa) or Macrocystis (South America). Laminariales communities are also dominant in the Pacific Ocean, as illustrated by the Macrocystis forests along the west coast of North America (Fig. 5.8) or the Saccharina and Undaria

forests along the Japanese and Korean coasts. Finally, forests of Ecklonia dominate the south coast of Australia. Most of the species forming the canopy of these kelp forests are perennials having a life span of several years (2 to

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Fig. 5.8 Kelp forest with Macrocystis pyrifera at Monterey Aquarium, California. Courtesy of Burkhard Büdel

15 years), but some species are annuals (e.g. Undaria). Despite this, the growth is seasonal and occurs principally during the cold season and the early spring (Chapman 1974; Dayton 1985). The Fucales are also highly productive, but because they principally inhabit the intertidal zone, they are exposed to important stresses, which induce them to excrete a large amount of organic matter that therefore limits their biomass (Mann 1982). The growth of the Fucales occurs in the summer; during the winter, previously stored carbohydrates compensate the lower rate of photosynthesis (Chapman 1974). In tropical regions, the most commonly found and dominant brown algae belong to the genus Sargassum (Fucales). Famous for producing immense rafts of unattached and floating plants such as occur in the so-called Sargasso Sea. The floating Sargassum rafts occur in warm waters that are too deep for benthic attachment; however, these floating rafts create ecosystems that are biologically analogous to kelp forests (Fulton et al. 2019). In recent years, Sargassum blooms have started to become a major issue. The “great Atlantic Sargassum belt” forms every year since 2011, and is visible from space, extending from the Sargasso Sea to the West coast of Africa. Pollution and global climate change are likely playing a role in the formation of these blooms, and the blooms are now having an ecological and economical impact along the Atlantic intertropical coastlines (de Széchy et al.

2012; Addico and deGraft-Johnson 2016; Louime et al. 2017; Mendez-Tejeda and Rosado Jiménez 2019). Other Phaeophyceae species (e.g. Dictyota, Lobophora, and Padina) also play important ecological roles in tropical coastal areas despite their small size when compared to kelps or Sargassum (Briones-Fourzán and Lozano-Álvarez 2001; Vieira 2020). Only a handful of brown algal species are known from freshwater. All freshwater browns are relatively small and limited to filamentous or crustose forms (Bold and Wynne 1985; McCauley and Wehr 2007; Dittami et al. 2017). Brown algae are economically important as well. Kelp forests cover around 25% of the world’s coastlines (Wernberg et al. 2019), and in addition to the ecological benefices listed above, they also provide substantial indirect services to humans. These services include notably carbon sequestration (Krause-Jensen and Duarte 2016; Krause-Jensen et al. 2018; Hoegh-Guldberg et al. 2019; Hwang et al. 2022), also supporting fish stocks and tourism. Overall, these indirect services are estimated to be worth $500,000–1,000,000 per year per km2 (Filbee-Dexter and Wernberg 2018) for a global value probably reaching hundreds of billions of USD per year. Phaeophyceae also have direct economic value through food consumption and commercial extracts (Mautner 1954; Vásquez et al. 2014; Bennett et al. 2016; Milledge et al. 2016). Brown algae have been harvested from the wild for

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centuries (Bell 1981; Ainis et al. 2019), and since the 1950s industrial-scale mariculture has been established in Eastern Northern Asia (Hwang et al. 2019) and elsewhere (Araújo et al. 2021; Brakel et al. 2021). The most widely cultivated brown algae are Saccharina, Undaria, and Sargassum. The global value of this mariculture is estimated to be 6.8 billion US$ for 14.9 million wet tons (FAO 2020). Furthermore, farming of seaweed is now seen as a way to locally offset carbon emissions and reduce ocean acidity (Froehlich et al. 2019). The unique metabolism of the Phaeophyceae produces many bioactive chemicals with interesting applications, like the anti-inflammatory, anti-viral, anti-biotic, anti-oxidant, and anti-coagulant properties of the fucose-containing sulfated polysaccharides (Li et al. 2008; Fitton 2011; Morya et al. 2012; Van Weelden et al. 2019; Jin et al. 2021). Particularly, the unique ability of kelps to concentrate iodide in their cell wall has been linked to reduced breast cancer incidence in populations consuming brown algae (Teas et al. 2013). Evolutionary history The long academic interest in brown algae and their general ecological and economic importance explains the dynamism of the research on the evolution of the Phaeophyceae. In general, reading the extensive and comprehensive review by Bringloe et al. (2020a) is highly recommended to gain insights into the evolution of the Phaeophyceae. Only a handful of Phaeophyceae species produce hardened bodies (genera Newhousia and Padina), and this has greatly limited their fossilization. Furthermore, convergent evolution within the green and red algae makes morphological identification of brown algal fossils a complicated and debatable task (Fry and Banks 1955; Krings et al. 2007). For long, the oldest fossil assigned with certitude to the Phaeophyceae dated from the Cretaceous Period (145–66 Mya), and was morphologically consistent with the modern genus Padina (Rajanikanth 1989). A much older fossil from the Neoproterozoic (600–550 Mya) was assigned to the brown algae based on the presence of conceptacles typical of the contemporary Fucales (Xiao et al. 1998). Other controversial ancient brown algal fossils have been reported from the Silurian Period (443–419 Mya), but their nature remains controversial (Taggart and Parker 1976). Laminariales and Sargassum-like fossils from Miocene deposits (13–17 Mya) have been recovered and described (Parker and Dawson 1965). In the past decade, the use of time-calibrated multigene phylogenies has provided a timeline and phylogenetic frame to investigate the evolution of the Phaeophyceae (Silberfeld et al. 2010; Kawai et al. 2015; Starko et al. 2019; Yip et al. 2020). Briefly, it is now thought that the lineages of the Phaeophyceae diversified in the Mesozoic Era (252–66 Mya) in roughly three phases: early

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divergence of the Discosporangiales and Ishigeales; the mid-Mesozoic diversification of the Sphacelariales, Syringodermatales, Dictyotales, and Onslowiales; and the radiation of the remaining orders (the brown algal crown radiation; BACR) during the Cretaceous (reviewed in Bringloe et al. 2020a). This timeline, along with multiple phylogeographic and genetic studies (e.g. Starko et al. 2019; Zhang et al. 2019; Bringloe et al. 2020b), also brings insights into the contemporary distribution of the Phaeophyceae and the events linked to it. For example, the Oligocene Epoch (34–23 Mya) was characterized by global cooling that allowed the Laminariales to become dominant (reviewed in Bringloe et al. 2020a). However, these time-calibrated multigene phylogenies did not include the oldest potential fossil from the Neoproterozoic (600–550 Mya), and therefore the timeline of the diversification of the brown algae could be much older than it is currently thought. In the future, the discovery of more fossils and more phylogenetical analysis should bring new insights on this matter. Evolutionary studies of the Phaeophyceae have gained pace with the sequencing of a species of Ectocarpus (Cock et al. 2010; Cormier et al. 2017) and the development of genomic resources (i.e. transcriptomics, high-density genetic map) for the Phaeophyceae (reviewed in Bringloe et al. 2020a). It is hypothesized that ancestral sexual reproduction was isomorphic with isogamous gametes like that of Discosporangiales and Ishigeales (Cho et al. 2004; Bringloe et al. 2020a, b), with later transitions to heteromorphic cycle like that of Laminariales or Ectocarpales and multiple transitions to anisogamous and oogamous gametes (Silberfeld et al. 2010; Heesch et al. 2021; Bringloe et al. 2020a, b). Furthermore, sex determination has been recently investigated with the analysis of transcriptomes in multiple Phaeophyceae species (Martins et al. 2013; Lipinska et al. 2015; Coelho et al. 2019). The genetic basis of the development of the complex morphologies of the Phaeophyceae is also beginning to be unrevealed (Peters et al. 2008; Charrier et al. 2012; Macaisne et al. 2017).

5.1.4.2 Chrysophyceae The Chrysophyceae regroups a vast diversity of organisms named after their golden-brown coloration (Greek chrysós— gold). Brief history and current taxonomy Some of the first described Heterokontophyta were members of the Chrysophyceae, i.e. Anthophysa and Monas (=Oikomonas in Müller 1773, 1786). Probably due to their abundance in freshwater bodies of water across Europe, many Chrysophyceae species were reported, among other microorganisms, in the beautifully exemplified work by Ehrenberg (1838). Typical and iconic colonial genera such

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as Dinobryon, Synura, and Uroglena (Fig. 5.9) have been known since that period, with the first concept of the class being indicated a few decades later (Stein 1878). The Chrysophyceae progressively became a group into which virtually all golden-brown flagellate, amoeboid, capsoid, and coccoid microorganisms were placed (Pascher 1913). This large class was fixed by Pascher (1914, 1921) and represented for a time all the diversity of the Heterokontophyta outside diatoms, Phaeophyceae, and Xanthophyceae (Bourrelly 1957, 1965). Within the Chrysophyceae, taxonomy was based on the number of flagella, distinguishing the uniflagellate and biflagellate lineages (e.g. Pascher 1913; Bourrelly 1957 who also further separated those taxa with no known flagella). The technological revolution of the electron microscopy revealed a second extremely short flagellum in uniflagellate species prompting discussion on the taxonomy of Chrysophyceae (e.g. Hibberd 1976; Andersen 2007). Furthermore, electron microscopy and molecular biology-generated data led to distinguishing new classes of algae from the polyphyletic Chrysophyceae sensu Pascher or Bourrelly (Fig. 5.1). Among these classes were the Synurophyceae (Andersen 1987), which was differentiated from the Chrysophyceae based on pigment composition, flagellar root system, photoreceptor, posterior contractile vacuoles, the absence of an eyespot, and the absence of a chloroplast endoplasmic reticulum. More recently, molecular phylogenetic analyses showed that the “Synurophyceae” were nested within the Chrysophyceae (e.g. Ben Ali et al. 2002; Del Campo and Massana 2011; Yang et al. 2012; Škaloud et al. 2013; Dorrell et al. 2019) and prompted the combination of the Synurophyceae within the Chrysophyceae as the order Synurales. Including around 1,200 species (Guiry and Guiry 2021), the Chrysophyceae exhibit a wide range of forms and shapes, colonies and solitary species, autotrophic, mixotrophic, or heterotrophic species (Fig. 5.9). Therefore, they present interesting models for the study of algal evolution. General description The Chrysophyceae are a diverse group presenting many morphologies and cellular organizations. The Chrysophyceae include unicellular, colonial, amoeboid, capsoid, and coccoid taxa. Many specific cases can be described in the Chrysophyceae, and presenting an exhaustive description would depass the scope of this chapter. We will limit to the description of characters most commonly found in the Chrysophyceae and point to further readings for the interested reader (e.g. Pascher 1925; Bourelly 1957; Starmach 1985; Kristiansen and Preisig 2001; Andersen 2007; Nicholls and Wujek 2015; Kristiansen and Škaloud 2017).

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As in other lineages of the Heterokontophyta, motile and nonmotile forms of cells are commonly observed in the Chrysophyceae. Vegetative cells have one or two (rarely more) chloroplasts that dominate the protoplasm. Chloroplast shape varies, but frequently a chloroplast will be bilobed and located parietally within the cell. Pyrenoids, when present, are usually located in the center of the chloroplast. The cell posterior usually contains storage products, i.e. a single large chrysolaminarean vacuole and several to many spherical lipid bodies. The periphery of the cell may have specialized vesicles that eject material (e.g. discobolocysts, Hibberd 1970; mucocysts, Andersen 1982). Normally, there is a single Golgi body that lies adjacent to the nucleus on the anterior end of the cell. Freshwater organisms lacking a cell wall typically have one or two contractile vacuoles. Swimming cells have two flagella (Fig. 5.9), although in some taxa the mature flagellum is not visible by light microscopy (and these formed the so-called “uniflagellate” Chrysophyceae). The length of the mature flagellum defines two general morphologies of the Chrysophyceae swimming cells: the Ochromonas-like type with two easily visible flagella and the Chromulina-like type with a single visible flagellum (Fig. 5.9). The orientation of the two flagella may be parallel as found in Synura (e.g. Schnepf and Deichgräber 1969), approximately perpendicular as found in Chrysosphaerella (e.g. Andersen 1990), or extending in opposite directions as found in Hibberdia or Kremastochrysopsis (Andersen 1989; Remias et al. 2020). At the base of the mature flagellum, a photoreceptor is generally found in a swelling. An eyespot can be observed at the anterior end of the chloroplast of most species, but it is absent in the Synurales. Maybe the characters most clearly unifying the Chrysophyceae into a single distinct class are biochemical and ultrastructural. The flagellar system is typical and complex (reviewed in Kristiansen and Škaloud 2017). The flagella are anchored with varying arrangements of microtubular and striated roots (e.g. Andersen 1991), but all species examined have transitional fibers that connect the basal body/flagellum transition to the plasma membrane. Chloroplasts are typical of the Heterokontophyta, surrounded by four membranes with a chloroplast endoplasmic reticulum, three-fold stacked lamellae, and a girdle lamella (except in the order Synurales). Photosynthetic pigments are primarily chlorophyll a, c1 and c2 (the Synurales lack chlorophyll c2). Accessory pigments give the Chrysophyceae their golden-brownish color and are mainly fucoxanthin, and ßcarotene of the antheraxanthin-violaxanthin cycle. One or more mitochondria are found in the cell, and they have typical tubular cristae.

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Fig. 5.9 Diversity of the Chrysophyceae. a Living colony of Uroglenopsis turfosa showing the typical circular arrangement with dense peripheral cells and an opening in the center of the colony; b Living colony of Dinobryon sertularia; c Cells of D. sertularia within the colony; the immature flagellum escapes from the opening in the lorica and the eyespot (arrow) near the base of the flagella; d Cells of Chrysamoeba radians showing the multiple pseudopodia; e Cells of Derepyxis amphora growing epiphytically on an unidentified chlorophyte; f Living colony of Synura spinosa showing the pyriform cell morphology and the organization of the colony, radiating from a central point; g Close-up of a short filamentous branch on the thallus of Hydrurus c.f. foetidus; h Macroscopic view of the branching thallus like of H. c.f. foetidus; i Cells of Mallomonas caudata showing the radiating bristles; j Close-up of a cell of M. caudata showing the overlapping scales and the attachment of bristles; k Ochromonas triangulata; upper cell exhibiting the typical triangulate cell morphology with two flagella and a red eyespot, lower cells demonstrating the cell morphological variability; l Chrysopyxis bipes growing attached to an unidentified chlorophyte (bottom). a–e, l adapted from Nicholls and Wujek (2015); f, i, j courtesy of Robert A. Andersen; g, h courtesy of Michael Schagerl; k adapted from Andersen et al. (2017)

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Fig. 5.10 Scales and stomatocysts of the Chrysophyceae. a Scales and bristles covering the cell of Mallomonas sp. b Scales covering the cells of a colonial Synura lanceolata. c Cell of S. soroconopea showing the overlapping scales and their arrangement. d, e Components and diversity of the scales of Mallomonas spp. f, g Components and diversity of the scales of Synura spp. a, d, e adapted from Siver et al. (2015); b, c adapted from Jo et al. (2016)

In addition to unicellular flagellates, the Chrysophyceae include colonial flagellates (e.g. Synura, Uroglena, and Uroglenopsis), loricate colonial flagellates (Dinobryon, Epipyxis), macroscopic colonies (e.g. Hydrurus, Celloniella), and amoeboid species (e.g. Chrysamoeba) (Fig. 5.9). The colonies of Uroglena consist of numerous naked biflagellate cells attached to bifurcating cytoplasmic strands, and each cell resembles the classical Ochromonaslike cell. The amoeboid Chrysamoeba has vegetative cells with pseudopodia and frequently a short visible immature flagellum, and at times it produces zoospores that are indistinguishable from a “typical” Chromulina-like swimming cell. If many species of the Chrysophyceae occur as naked cells, with their cell membrane directly in contact with

water, some are covered by a cell wall, lorica, mineralized scales, spines, and bristles (Fig. 5.10). The loricas are cell walls presenting one or multiple openings and can take various shapes and composition (e.g. Belcher 1969; Herth et al. 1977). The lorica of Poterioochromonas resembles a champagne glass, with the cell residing inside the “bowl” and the “stem” attached to the substrate. The lorica of Dinobryon is often described as “vase-shaped”, with the cell attached inside the vase and the two flagella extending out the mouth of the vase. Dinobryon cells attach together in a tree-branching fashion to produce a colony (Fig. 5.12). Unlike the sessile Poterioochromonas, Dinobryon actively swims in the plankton. Both Poterioochromonas and Dinobryon produce cellulose loricas. The

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colonial Epipyxis is sessile, but the loricas are joined together similar to Dinobryon; however, Epipyxis loricas are formed from overlapping organic scales. Scales are also found surrounding the cells of some genera, and like loricas, the scales are presumed to provide protection for the otherwise naked cell (Fig. 5.10). Scales may be either organic or mineralized. Organic scales may have elaborate shapes, such as those found around the cell body of Chrysolepidomonas (Peters and Andersen 1993a, b), or they may be small disk-like structures such as those found along the flagella of Synura (Hibberd 1973). Silica scales are typical but not restricted to the order Synurales. Similar to diatoms, silicification occurs in deposition vesicles produced in the chloroplast endoplasmic reticulum (Schnepf and Deichgräber 1969), molded by vesicles of Golgi body origin and finally extruded outside the cell and placed correctly in relation to previously formed scales. In Mallomonas, bristles are formed similarly (Wujek and Kristiansen 1978; Mignot and Brugerolle 1982) and fixed on each scale through a complex mechanism (Beech et al. 1990). In the genus Paraphysomonas, scale production involves two different vesicles both from endoplasmic reticulum origin (Preisig and Hibberd 1983). All these ornaments can be extremely complex and elaborate, so they are frequently used for species-level taxonomic identifications (e.g. Škaloud et al. 2013). The life history of the Chrysophyceae is also extremely diverse. Sexuality is known but remains poorly understood for many species. When observed, gametes are morphologically similar to swimming cells (i.e. isogamete); they fuse and form a globular zygote (e.g. Fott 1959). Dinobryon is particularly well studied and representative of both autogamic (i.e. asexual reproduction) and gametic reproduction in Chrysophyceae (Fig. 5.11; Sandgren 1981). In species of the order Synurales, reproduction is similar (Wawrick 1972; Sandgren and Flanagin 1986). Many Chrysophyceae can form a silica cyst, especially under unfavorable conditions (Fig. 5.12). These unique cysts are also known as statospores or stomatocysts. The formation of stomatocysts can be sexual or asexual and appears to be uncorrelated with a particular environmental trigger but rather with cell density. In culture, encystment is observed during exponential population growth or stationary phase. In the first case, mostly sexual cysts are formed whereas asexual spores are produced in later growth. This fine-tuning between the population and its environment suggests the capacity of the algae to produce or detect chemical clues. All stomatocysts, whether sexual or asexual, are formed in a peripheral silica deposition vesicle resulting from the fusion of Golgi vesicles. This process is reminiscent of silica scale formation and of frustule formation in Diatomeae. A region at the apex of the cell escapes the silification and forms an

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Fig. 5.11 Life cycle of Dinobryon sp. During sexual reproduction, the zygote fusion occurs within one lorica between a stationary gamete (termed female) and a motile gamete (termed male) entering the lorica (from Lee 2018)

opening called a pore through which the cytoplasm exterior to the deposited silica wall may retract. Once the cell is inside the cyst, the pore is closed by a pore plug, generally surrounded by a silicified collar. Stomatocysts are mostly spherical, 2 to 30 µm in diameter, smooth or ornamented by protuberances or spines in different arrangements, and generally species-specific (reviewed in Duff et al. 1995 but see Findenig et al. 2010). Like encystment, excystment (germination) is also poorly understood but appeared to be most frequent in spring or when exposed to light (Sheath et al. 1975; Sandgren 1983). Occurrence Chrysophyceae are primarily members of the freshwater plankton and may constitute most of the planktonic biomass in ponds or bogs. Chrysophyceae thrive in neutral or slightly acidic water with a moderate supply of nutrients but are also found in more acidic or alkaline waters. Interestingly, many silica-scaled species occur within a narrow well-defined pH

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Fig. 5.12 a–l Various stomatocysts’ morphotypes of Chrysophyceae. Scale bars = 2 µm. All figures adapted from Firsova et al. (2008)

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range or trophic gradient (Smol et al. 1984; Siver and Hamer 1990; Siver and Marsicano 1996) and can be used as bioindicators. Some species are found in running water (e.g. Hydrurus), as neuston attached to the surface layer (e.g. Chromophyton), or even on snow (Tanabe et al. 2011). Many species occur primarily in cold water and therefore occur in open waters during spring and autumn or under ice during the winter; nevertheless, some common chrysophytes are also found in warmer waters during the summer. Only a relatively few marine species are known. The colorless Paraphysomonas includes a considerable number of marine species that play an important bacterio-grazing role in the marine food web (Lim et al. 1999; Ikävalko 2001). A few Ochromonas-like flagellates, including Melkoniania moestrupii, inhabit the coastal and open oceans (Andersen 2011), while Ochromonas triangulata is found in saline lakes and can be grown in seawater (Andersen et al. 2017). Freshwater plankton such as Dinobryon, Mallomonas, and Synura are often collected in seawater after they have washed into the ocean, and they remain alive for several hours but eventually die. Recent environmental DNA studies have revealed important and unsuspected cryptic marine diversity (del Campo and Massana 2011; Delmont et al. 2022). Chrysophyceae often express more than one mode of nutrition (autotrophy, mixotrophy, and heterotrophy), and as a consequence, they may fill more than one role in the

community (Sanders and Porter 1988). Furthermore, the degree to which mixotrophic species rely on phototrophy, osmotrophy, or phagotrophy varies a lot. The Synurales, capsoid forms (e.g. Chrysocapsa, Hydrurus) and those with true cell walls (e.g. Phaeoplaca) appear to be phototrophs, the colorless flagellates (e.g. Paraphysomonas, Spumella) are heterotrophic phagotrophs, many of the flagellates with chloroplasts (e.g. Dinobryon, Epipyxis, and Ochromonas) are mixotrophic by photosynthesis and phagotrophy, and amoeboid forms with pseudopods (e.g. Lagynion) may be mixotrophic by photosynthesis and osmotrophy. These fascinating physiologies and their ecological implications have been extensively studied (e.g. Bird and Kalff 1986; Caron et al. 1990; Wetherbee and Andersen 1992; Holen and Borass 1995; Zhang and Watanabe 2001; Maberly et al. 2009). The prey of the phagotrophic species includes in certain cases other eukaryotes such as diatoms or green algae (Caron et al. 1990), toxic cyanobacteria (Cole and Wynne 1974), cannibalism (Caron et al. 1990), and feeding on non-living particles (Wetherbee and Andersen 1992). However, in most cases, the prey is bacteria with up to 190 bacteria consumed per cell per hour (Holen and Borass 1995). Flagella play an important role during predation, both by beating to create a strong current toward the apex of the cell and by seizing the prey (Wetherbee and Andersen 1992). Blooms of Chrysophyceae species can produce undesirable effects, notably through the organic compounds they excrete.

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Among these, aldehydes and ketons can give a fishy smell and taste to water and foul drinking water reservoirs (Nicholls and Wujek 2015). Evolutionary history Fossil record of the Chrysophyceae is abundant due to the silicified scales and stomatocysts that form sediment layers preserved through geological time. Stomatocysts are more resistant to dissolution processes than scales and bristles because the silica layer is thicker; however, cyst morphologies are rarely linked to species and therefore species identifications are usually not possible (Duff et al. 1995). The oldest cysts reported thus far date back from the Late Triassic (235–228 Mya) from a freshwater paleoenvironment (Zhang et al. 2016b) and from the Upper Cretaceous (112 Mya) from marine deposits (Riaux-Gobin and Stumm 2006). Silica scales have been reported from the Cambrian (541–485 Mya), but these are probably better assigned to protists other than the Chrysophyceae (Allison and Hilgert 1986). Fossils of scales of the genus Synura are known from the Paleocene epoch (60 Mya; Siver et al. 2013) and other silica genera (e.g. Mallomonas) are known from the younger Middle Eocene epoch (47 Mya; Siver and Wolfe 2005; Siver et al. 2009; Siver and Lott 2012). Based upon molecular clock analysis, the origin of Chrysophyceae was estimated in the Permian period (279– 250 Mya) in two independent studies (Brown and Sorhannus 2010; Jo et al. 2013) but these did not include the Late Triassic (235–228 Mya) old stomatocysts. Similarly, diversification within the Chrysophyceae was estimated at earlier ages than suggested by the fossil record (Jo et al. 2013; Siver et al. 2013, 2015). Finally, both the ancient silica fossils and the phylogenetic position of a silica-scaled genus Paraphysomonas as the first diverging lineage of the Chrysophyceae (Andersen et al. 2017; Dorrell et al. 2019) suggest that silification is an ancestral trait in the Chrysophyceae. It is noteworthy that no silica cysts have been reported for Paraphysomonas. Historically, most Chrysophyceae species were described based on morphology. However, the higher level classifications based on morphology have often proved to be incongruent with molecular phylogenetic analyses. In recent years, it has become clear that the Ochromonas-like taxa (over 100 taxa) and their “colorless” counterparts the Spumella-like taxa (over 30 taxa) both represented polyphyletic assemblages due to convergent evolution (e.g. Grossmann et al. 2016; Andersen et al. 2017). That is, the genera Ochromonas and Spumella are now very small genera and the remaining species are assigned to several other genera. Similarly, it seems likely that the Chromulina-like taxa (over 200 classical species) will be recognized as a polyphyletic assemblage once the type species; Chromulina nebulosi is

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anchored in a molecular phylogeny. Finally, the colonial genera Uroglena and Uroglenopsis were once combined into a single genus (Bourrelly 1957), but a recent study shows that they are not only distinct genera but that a third genus, Urostipulosphaera, has a similar morphology (Pusztai and Skaloud 2019). Furthermore, these polyphyletic lineages are all interspread together with multiple transitions between Ochromonas-like and “colorless” Spumella-like, unicellular Ochromonas-like, and Uroglena-like colonies and so forth. In summary, the higher level classifications and phylogenies, which were based upon light microscopical morphological characters (e.g. Pascher 1914; Bourrelly 1957; Starmach 1985), are not supported by molecular phylogenetic analyses. A majority of Chrysophyceae genera have not been examined using molecular sequence data, and therefore the current classification of the Chrysophyceae is in flux. Furthermore, the recent progress reported above revealed the dynamic evolutionary processes occurring within the Chrysophyceae. The diversity of nutrition modes observed in the Chrysophyceae is also of evolutionary interest. Whether the common ancestor of the Chrysophyceae was an obligate phototroph or a mixotroph remains debatable. However, because the sister lineage of the Chrysophyceae (i.e. Synchromophyceae) has a mixotrophic mode of nutrition, the latter appears more parsimonious. Because mixotrophy can have a higher metabolic cost than obligate phagotrophy or phototrophy, why is this nutritional variability maintained in the Chrysophyceae? An ancestral loss of the carbon anhydrase likely occurred in the Chrysophyceae, and could help explain the importance of phagocytosis to complement photosynthesis (Maberly et al. 2009; Raven 1995, 2010). In addition to providing carbon, phagocytosis provides important B vitamins and biotin (Moestrup and Andersen 1991; Holen and Boras 1995). Finally, Chrysophyceae need important amounts of iron to synthesize one of their cytochromes, and phagocytosis is thought to provide this iron (Raven 1995). Obligate heterotrophy is widespread in the Chrysophyceae and photosynthesis was lost multiple times (Dorrell et al. 2019). Interestingly, genomic exploration of these independent secondary losses of photosynthesis revealed converging loss of plastid functions (Dorrell et al. 2019; Kim et al. 2020). Plastid genomes were greatly reduced (Kim et al. 2020) and even completely absent in some cases (Dorrell et al. 2019). Interestingly, proteins that were ancestrally targeted to the plastid now support the mitochondria metabolism (Dorrel et al. 2019).

5.1.4.3 Xanthophyceae The Xanthophyceae are a diverse and large assemblage of algae that differ from most other Heterokontophyta groups because of their yellow-green coloration (Greek xantho = yellow).

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Brief history and current taxonomy Macroscopic forms, such as Vaucheria (de Candolle 1801; Fig. 5.16), were first classified as a distinct order, the Confervales, within the green lineage Chlorophyceae (Braun 1855; Borzi 1889, 1895; Bohlin 1897). Interestingly, they were characterized not only by their yellow-green chloroplasts but also by motile cells possessing a single flagellum. However, with the description of Chlorosaccus, a new member of the Confervales led to the discovery that its motile cells and those of Tribonema (Fig. 5.13) and Botrydiopsis possessed two flagella of unequal length (Luther 1899). Progressively, studies of the Confervales revealed the incompatibility of their inclusion within the Chlorophyceae and following Luther’s work, Pascher (1914) described the Heterokontae with the orders Heterochloridales, Heterocapsales, Heterotrichales, and Heterosiphonales for the flagellate, capsoid, coccoid, filamentous, and siphonous forms, respectively. The name Xanthophyceae (Allorge 1930; Fritsch 1935) was proposed about 20 years later, and this class name became widely accepted. The first half of the twentieth century saw the description of many species of the Xanthophyceae, principally by Adolf Pascher who authored two-thirds of the generic names. Pascher's work culminated in his famous Heterokontae (Pascher 1939). The development of electron microscopy in the second half of the twentieth century helped to define the morphological characteristics of the Xanthophyceae (Manton et al. 1952; Greenwood et al. 1957; Greenwood 1959; Hibberd and Leedale 1971; Ott 1982). Hibberd and Leedale (1970, 1971) discovered that some yellow-green algae belonged in a new class, the Eustigmatophyceae, and taxa continue to be transferred from the Xanthophyceae to the Eustigmatophyceae (see Sect. 5.1.4.6 Eustigmatophyceae, below). Many Xanthophyceae still have not been investigated by EM, and molecular analysis have revealed that many orders, families, and genera are paraphyletic or polyphyletic (Negrisolo et al. 2004; Maistro et al. 2007, 2009). Currently, 690 species in 129 genera (Guiry and Guiry 2021) are recognized in the Xanthophyceae, but the boundaries of this class are still under discussion and many species await re-investigation and taxonomic revision. General description The Xanthophyceae are principally represented by coccoid unicellular species but filamentous and siphonous (coenocytic, with multinucleated cells) species are also well known (Figs. 5.13, 5.16). Cells usually contain several chloroplasts that give the bright green to yellow-green coloration. This coloration is unusual within the Heterokontophyta.

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Chloroplasts have chlorophyll a, c1, and c2, but the chlorophyll c pigments are only found in extremely low concentrations (Table 5.2). The accessory pigments include diadinoxanthin, diatoxanthin, heteroxanthin, vaucheriaxanthin esters as well as ß-carotene (Whittle and Casselton 1975a, b; Rowan 1989; Jeffrey et al. 2011). The notable absence of fucoxanthin in the Xanthophyceae is responsible for the green to yellow-green coloration, as it usually confers their brownish nuances to the other Heterokontophyta. The vegetative cell is covered by a cellulose cell wall in most species, and the wall may be sculpted by fine ornamentations, and it may be composed of two overlapping halves (bipartite cell wall) in the filamentous species and the large coccoid species (Fig. 5.13). The two halves are not easily observable and are most apparent when they separate (e.g. during zoospore release). When the cell wall is bipartite, the two halves can be more or less equal in size but always overlap widely. In the filamentous species, the bipartite cell wall is formed of two interlocking H-shaped pieces (Fig. 5.13). Many Xanthophyceae species produce zoospores or spermatozoids. These cells usually have an ovoid or pyriform shape and range from 5 to 20 µm. The motile cells possess the typical Heterokontophyta flagella inserted laterally near the apex of the cell on a small raised cytoplasmic protuberance (Hibberd and Leedale 1971). The long immature flagellum bears tripartite hairs and the short mature flagellum is smooth. The motile cells usually have two chloroplasts, and an eyespot is present directly under the mature flagellum swelling (Fig. 5.14). Ultrastructural analysis show the typical Heterokontophyta organization of the chloroplasts, with three stacked thylakoids, a girdle lamella, and four membranes, the outermost being continuous with the nuclear envelope. Pyrenoids can be found in the chloroplasts but are not as easily observed as in other Heterokontophyta lineages. Most Xanthophyceae species reproduce asexually either by simple division or by producing biflagellate zoospores or aplanospores (Box 5.1). Filamentous species can also reproduce by fragmentation of the filament. The formation of the zoospores greatly modifies the cell structure (Falk 1967; Deason 1971; Hibberd 1980). Briefly, the chloroplasts move to a more internal position within the cell, and each is associated with a nucleus. After the formation of eyespots in the chloroplasts, the nucleus-chloroplast pairs are separated by vacuoles to each form a zoospore. Interestingly, Vaucheria forms a multinucleate multiflagellate synzoospore at the tip of siphonous filaments (e.g. Ott and Brown 1974). In some filamentous genera (e.g. Vaucheria), asexual reproduction can also occur by aplanospore formation at the apex of the filament or within the filament by akinete

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Fig. 5.13 Various morphologies of Xanthophyceae species in culture; a–d Morphology of three strains of Heterococcus viridis showing morphological plasticity of sarcinoid pseudofilaments; a prostrate colonies produced by branched filaments on the surface of an agarized culture, 16 weeks old (strain B10); b Enlarged view of a young (4 weeks old) colony from a liquid culture (strain SAG 835–7); c Enlarged filament, 6 weeks old from an agarized culture (strain MZ3-7); d Coccoid cells in a 4 weeks old liquid culture (strain SAG 835–7); e Coccoid unicells of Pleurochloris meiringensis (strain SAG 2574); note thin envelope of mucilage autosporangia (packages of cells); f Short and thin filaments of Heterothrix mucicola (strain A16-5); g Tribonema vulgare (strain SAG 24.95) with prominent H-pieces of the cell walls (arrow heads) at one end of a filament fragment and in the middle of a filament; h Bumilleria sp. (strain SAG 2159), with cell wall composed of overlapping H-pieces (arrow heads); i Bumilleriopsis sp., cell wall composed of two interlocking parts, vegetative cells (left half of the image), dividing cells, and a large autosporangium (right half of the image); j Ophiocytium maius (strain SAG 855–1), elongated slightly curved cells, cell wall with a basal small stalk with which the cells are attached to the substrate, and composed of two interlocking parts of unequal size; a–d from Rybalka et al. (2013), BMC Evolutionary Biology 13: 39; e–h, j courtesy of T. Darienko with permission of the SAG culture collection; f courtesy of T. Friedl; i courtesy of Nataliya Rybalka, from FAO, ITPS, GSBI, SCBD, and EC. 2020. State of knowledge of soil biodiversity—Status, challenges and potentialities, Report 2020. Rome, FAO. https://doi.org/10.4060/cb1928en, www.fao.org

formation (Maistro et al. 2017). Sexual reproduction has been studied in the siphonous genera Botrydium and Vaucheria. Sexual reproduction is isogamous in Botrydium (Rostafinski and Woronin 1877; Kolkwitz 1926; Rosenberg 1930). In the genus Vaucheria, sexual reproduction occurs

through the differentiation of specialized sexual organs (gametangia) on the filament, the sperm-producing antheridium, and the egg-producing oogonium (Figs. 5.15, 5.16e, f). These sexual organs can either be found adjacent to each other on a single filament (i.e. monoecious species;

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Fig. 5.14 Zoospore of Mischococcus sphaerocephalus. Drawings adapted from light and transmission electron micrographs from Hibberd and Leedale (1971). C = chloroplast; CV = contractile vacuole; E = eyespot; FS = flagellar swelling; LF = immature flagellum; N = nucleus; SF = mature flagellum; V = vacuole (from Lee 2018)

Fig. 5.16e, f) or separated on distinct filaments (i.e. dioecious species). The fertilization occurs with the spermatozoids entering the oogonium through pores at its apex. The fertilized oospore (= zygote) will germinate and form a new filament (Fig. 5.15). Occurrence Xanthophyceae are distributed from the tropics to the polar regions. Most of their diversity is found in freshwater habitats, but they also compose a large part of the terrestrial microflora (Vischer 1945; Ettl and Gärtner 1995). A number of Vaucheria species are found in marine coastal habitats (e.g. Ott and Hammersand 1974; Fig. 5.16a–c). Planktonic species are found in still waters, with a preference for low pH, iron-enriched lakes, and bogs. The soils of Antarctica have been shown to contain coccoid and filamentous Xanthophyceae (Rybalka et al. 2009) as do the alpine soils (Ettl and Gärtner 1995). Despite their relatively large diversity, most Xanthophyceae occur in low density and are rarely observed. Many species have never been observed again since their

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Fig. 5.15 Life cycle of Vaucheria sp. On top, the three modes of asexual reproduction are presented by means of aplanospores, zoospores, or akinetes (from outside to inside). On the bottom, the sexual reproduction of a monoecious (sexual organs present on a single filament) species is presented (modified from Sahoo and Kumar 2015 and modified from Lee 2018)

description. The most easily observed Xanthophyceae are the genera Botrydium and Vaucheria (Fig. 5.16). Botrydium can form microscopic green thalli visible to the naked eye, and these can cover large patches of drying mud (Fig. 5.16d). Species of the genus Vaucheria are found in freshwater in running clear water or on the shoreline of lakes. They are also found in brackish and marine coastal waters. Vaucheria grows attached to various substrates where they can form dense mats easily observable (Fig. 5.16a–d). Evolutionary history Only a handful of Xanthophyceae fossils have been discovered despite their modern diversity and cellulosic cell walls, and sometimes doubt on their identity persists (Butterfield 2004). The Xanthophyceae were first associated with the chrysophytes and diatoms in Pascher's (1914) Chrysophyta, but electron microscopic observations of swimming cells suggested a relationship with brown algae (Moestrup 1970)

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Fig. 5.16 Siphonous (coenocytic) Xanthophyceae. a–g Vaucheria; h, i Botrydium; a muddy hummocks during low tide exposure with dense turfs of green filaments of Vaucheria; the alga is spreading at the transition between intertidal and subtidal zones in the Wadden Sea (European Atlantic), near the island of Sylt (Germany); b unicellular filaments of Vaucheria from mudflat with colorless rhizoids (washed free from sediment) between which residential tubes of mudflat animals are interwoven; c unicellular filaments of Vaucheria at the tip of a turf in backlight; d Vaucheria forming dark-green mats on moist soil; e a young gametophore of a monoecious Vaucheria bearing two developing oogonia (dark) and terminated by an circinate antheridium; f oogonium and circinate antheridium of V. sessillis at different levels of focus; g organization of tubular cells of Vaucheria, lower magnification level (left) showing numerous plastids that are located in a thin layer of cytoplasm surrounding a large vacuole, higher-magnification view (right), some of the nuclei are indicated by arrows; h thallus of coenocytic vesicles of Botrydium granulatum on the surface of garden soil; i coenocytic vesicle of B. granulatum with rhizoidal extensions; inset, schematic drawing of a Botrydium vesicle with rhizoidal extensions; a–c courtesy of Karsten Reise; d, e courtesy of Choi Seokwan; f, i Ott et al. (2015); g Graham et al. (2016); h courtesy of Thomas Friedl; i inset, original Frederik Spindler

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Fig. 5.17 The lidded box principle of the diatom cell wall; a scanning electron micrograph of the two larger components (valves) of the cell wall of the pennate diatom Caloneis amphisbaena with bilaterally symmetrical cells; the two valves lie separated from each other; the upper valve (lid portion) is the older and slightly larger epivalve (epitheca), it is shown from the outer side, the lower valve (bottom portion) is the younger and smaller hypovalve (hypotheca), it is shown from the inner side; note central raphe and systems of striae (stripes) at the periphery of the cell wall; b valve view of a living cell of C. amphisbaena; note the two brownish chloroplasts appressed to the plasmalemma and location of the nucleus at the center in a bridge separating the two vacuoles; c schematic drawings of a hypothetical pennate diatom cell at two different angles illustrating the box-like construction of two overlapping valves; the left drawing is at valve view with view on the outer surface of the hypotheca (bottom); middle and right drawings are at girdle view at surface view (middle cell), and optical section (right cell); a, b Courtesy of Research Group Diatoms, Bo Berlin, Freie Universität Berlin, modified from Zimmermann et al. (2021), www.biuz.de, VBiO e.V. under the CC-BY-SA 4.0 license

that was then confirmed with molecular phylogenetic analysis (Ariztia et al. 1991). Further studies pointed to the close relation between the Xanthophyceae and the Phaeophyceae (forming the so-called PX clade [Kai et al. 2008], which now includes the Aurearenophyceae, the Chrysoparadoxophyceae, Phaeosacciophyceae, Phaeothamniophyceae, and Schizocladiophyceae [Graf et al. 2020a, b]). The different forms found in the Xanthophyceae (i.e. coccoid, filamentous, and siphonous) appeared independently multiple times with no clear order (Maistro et al. 2009). The modes of sexual reproduction within the genus Vaucheria have been another focus of evolutionary studies in the Xanthophyceae. Molecular phylogenies revealed a gradual evolution of the sexual reproduction with transitions from an asexual ancestor to hermaphrodism, the monoecy, and finally a single transition to dioecy (Andersen and Bailey 2002).

5.1.4.4 Diatomeae—The Diatoms Diatoms are a monophyletic group that is gradually divided into several phylogenetic lineages. Here, we treat the diatoms as a single group, the Diatomeae sensu Adl et al. (2019). The name “diatom” refers to the two overlapping silica frustules covering the cell in this group (Greek diatomos—split in half). The diatoms are the most diverse and species-rich group of algae, and ecologically they contribute

up to 20% of the global photosynthetic carbon fixation (Mann 1999). Brief history and current classification Diatoms have been known to science since the eighteenth century (Round et al. 1990), but the term “diatom” was established in the early nineteenth century (genus Diatoma described by de Lamarck and De Candolle 1805 and Agardh 1824) and by the year 1850 more than 800 species had been described both from living material and from fossils. Because many express motility, diatoms were first interpreted as animals (e.g. Ehrenberg 1838). Kützing (1844) showed that diatoms were autotrophs, and he established their classification as algae. In the second half of the nineteenth century, the focus changed from the description of genera and species to studies of cell structures and life cycles. Particularly, the cell size reduction with each generation was explained independently (MacDonald 1869; Pfitzer 1869), complimenting the earlier discovery of the size restoration (Thwaites 1847). This work was further extended in the twentieth (e.g. Geitler 1932; Roshchin 1994), and the demonstration that meiosis was an important contribution to diatom life cycles (Karsten 1912; von Stosch 1950). Historically, the taxonomical knowledge increased greatly

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during scientific expeditions on lands and seas, to Arctic and Antarctic regions, etc. (e.g. H.M.S Challenger’s voyage 1873–1876; Cleve and Grunow 1880; Heiden and Kolbe 1928). The dedication of single researchers also greatly advanced diatom knowledge, e.g. Friedrich Hustedt who described nearly 2,000 new species in the first half of the twentieth century. In the second half of the twentieth, the transmission electron microscopy provided unprecedented observations of cell and organelle ultrastructure that helped systematics and taxonomy to progress significantly. Scanning electron microscopy was also very important, providing detailed observations of the frustule and girdle bands (e.g. Round et al. 1990). The tremendous diversity of the cell wall structures in diatoms represents an important challenge for their classification and the correct identification of species (Williams 2020). Diatom classification is still under discussion. The classification of Round et al. (1990) regarded the diatoms as a phylum (Bacillariophyta) which they split into three classes, Coscinodiscophyceae, Fragilariophyceae, and Bacillariophyceae. These classes are readily identifiable by their cellular and valve organization (Mann et al. 2017). The Fragilariophyceae (araphide pennates) and Bacillariophyceae (raphid pennates) together comprised the pennate diatoms, the Coscinodiscophyceae comprised more or less all the centric diatoms. Several modern textbooks still follow that classification, e.g. Graham et al. (2016). Unfortunately, the system of Round et al. (1990) cannot be kept anymore as the two classes Fragilariophyceae and Coscinodiscophyceae were found paraphyletic in molecular systematics, and, therefore, the three-class system apparently does not capture the essential features of diatom evolution (Mann et al. 2017). Despite subsequent suggestions for diatom classification being based on molecular phylogenetics (e.g. Medlin 2016), no consensus has been reached so far as to what should replace the Round et al. classification and which systematic reconstructions accurately reflect diatom evolution (for review, see Mann et al. 2017). We therefore treat the diatoms here as a single group, Diatomeae, following Adl et al. (2019). It is a major and very diverse species-rich monophyletic lineage of the Heterokontophyta. In molecular phylogenetics, it forms together with the Bolidophyceae, Dictyochophyceae, and Pelagophyceae, the SIII clade (Yang et al. 2012; Figs. 2.8 and 5.2). As based on the Adl et al. classification system of eukaryotes, the Diatomeae is split into 9 lineages (subphyla), and 5 classes of diatoms are recognized (Adl et al. 2019). Diatoms have been good model organisms for exploring the physiology and biochemistry of the Heterokontophyta. This trend was accentuated in the early 2000s when Thalassiosira pseudonana became the first Heterokontophyta to have its genome fully sequenced (Armbrust et al. 2004). This achievement, along with the development of high

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throughput sequencing methods, allowed unprecedented analysis of the genetics, genomics, metabolism, ecology, biogeography, and evolution of the Diatomeae (e.g. Evans et al. 2009; Allen et al. 2011; Godhe et al. 2013; Alexander et al. 2015; Aumeier et al. 2015; Patil et al. 2015; Vanormelingen et al. 2015; Malviya et al. 2016; Moeys et al. 2016; Lewitus et al. 2018; Dorrell et al. 2021). Because of their long scientific history, their diversity, prevalence in the ecosystems, and utility for humans, the diatoms have been the primary subject of a large body of literature, e.g. The diatoms, Applications for the environmental and Earth sciences (Smol and Stoermer 2010), The Diatom World (Seckbach and Kociolek 2011), or Diatoms: fundamentals and applications (Seckbach 2019). General description The diatoms are essentially unicellular, but some lineages form chains of cells attached by polysaccharide bridges (e.g. Amphitetras; Tabellaria, Fig. 5.19a; Grammatophora, and Skeletonema) or silica spines and other interlocking cell wall extensions (e.g. Biddulphia, Chaetoceros, and Stephanopyxis). Their cells vary greatly in shape and size, from a few µm in diameter (e.g. Thalassiosira 3–5 µm; Fig. 5.19d) up to the mm scale (e.g. Ethmodiscus 2 mm; Thalassiothrix 5 mm), but most diatoms are in the 10–600 µm size range (Fig. 5.19). The diatoms are lacking the typical flagella of the Heterokontophyta, but in certain lineages, flagellated cell can be observed during the reproduction (i.e. in centric oogamous diatoms). A single flagellum is present, corresponding to the immature flagellum of the Heterokontophyta as it bears tripartite hairs. The second mature flagellum appears to have been completely lost, as did all the structure associated with it in other Heterokontophyta lineages. Despite the lack of flagella, certain diatoms (particularly the pennate forms with a central raphe in their wall) are capable of motility by means of gliding on a substrate. These movements are highly irregular in speed with short accelerations, and they successive forward and backward gliding movements; for details, see Graham et al. (2016). Diatoms are easily identified by their silicified cell wall, termed a frustule, that is only found in these algae. The wall is formed of two overlapping halves (thecae or valves), in a way reminiscent of a petri dish that are held together by silicious girdle bands (Fig. 5.17). The thecae are ornamented by pores of various sizes arranged in patterns (Figs. 5.17–5.19). The shape and size of the thecae and the arrangements of the pores are incredibly diverse and species specific. Two main valve shapes are recognized: the centric and pennate valves (Figs. 5.19, 5.20). Contrary to what their name suggests, shape of the centric valves can be oval, triangular, quadrate in addition to simply circular, but their pores are always

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arranged in radiating rows from a single ring (Fig. 5.17). The pennate diatoms were named after the feather-like organization of the pores (Latin penna—feather) that are arranged in rows on both sides of the rib that divides the generally elongated shape of the valve (Fig. 5.17). In fact, the diatoms' cell wall is multipartite. That is, besides the two main components (epivalve and hypovalve; Fig. 5.17) there is a series of smaller, linking components (girdle bands; Fig. 5.19d); for details, see Cox (2015) and Graham et al. (2016). Other complex structures and delicate arrangements of the valves, girdle bands exist (see Mann et al. 2017). The coloration of diatom cells varies from greenish, yellowish to brown. The majority of the cell volume appears colored because of the size and number of the chloroplasts; the shape and position is constrained by the cell wall structure. Despite the variety of shape of the chloroplasts depending on the species (e.g. discoid, lobed, and ribbonlike); they are always enclosed by four membranes and the thylakoids are surrounded by a girdle lamella. The photosynthetic pigments are chlorophylls a and c as well as, depending on the species, accessory pigments commonly found in the Heterokontophyta like fucoxanthin, diatoxanthin, and diadinoxanthin or other carotenoids (Egeland 2016). A pyrenoid is commonly observed in the chloroplast, and it can be very large and of various shapes (Fig. 5.3). The diatom life cycle is relatively well understood. Vegetative reproduction of the Diatomeae happens through mitosis during which the two valves separate during cytokinesis. As a result, each daughter cell carries only one of the valves and forms a new one, the hypotheca (bottom valve), to complete the mitosis cycle (Fig. 5.18). However, due to the size difference between the two valves forming the frustule, the deposition of the daughter frustule within the mother cell (inside of the plasmalemma), and once formed the diatom valves cannot increase in size; successive mitosis of the cell will mechanically result in a diminution of the valve dimension (Fig. 5.18). This continuous reduction could ultimately lead to the extinction of the population due to excessive dwarfism. The restoration of cells’ maximum size occurs through the sexual reproduction that is initiated when a critically small size is reached (Chepurnov et al. 2004). In the centric diatoms, reproduction is generally oogamous and homothallic whereas in the pennate diatoms, it is most commonly isogamous and heterothallic (the case illustrated in Fig. 5.18). Two cells enter the sexual reproduction and, when they do so, often secrete a protective mucilage surrounding them, within which the gametogenesis and fertilization occurs. Diatoms are diploid during their vegetative stages. Thus, their life cycle is diplontic (see Fig. 2.9c). Once in contact, formation of gametes occurs in both cells where they are contained in the silicified cell wall of the mother cell but they themselves are not enclosed in a silicified cell wall. In

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isogamous species, gametes are identical and non-flagellated but despite this, the pair of gametes from one cell (i.e. male gametes) will by amoeboid movement reach the other pair of gametes (i.e. female gametes), and they will fuse (Fig. 5.18). After the fertilization, the zygotes develop into a specialized cell type named the auxospore. During the differentiation of the auxospore, an organic cell wall is produced and as the auxospore grows, it can take different shapes. During its development, the auxospore will grow considerably in size and therefore restore the maximum size of the species. The cycle is completed with the formation of an initial cell within the auxospore through two successive mitoses each followed by the formation of a large valve, consequently reforming the typical frustule of the species. Occurrence As their important species diversity would suggest, the diatoms are commonly found in all marine and freshwater systems. In fact, the Diatomeae colonize all aquatic ecosystems where photosynthesis is possible, but they are found in greater number in productive zones such as the ocean’s upwelling regions. Outside of the oceans or lakes, the diatoms are also capable of growing on wet soil and even on the moist areas of Bryophytes or on the surface of leaves in tropical forests (Round 1981; Knapp and Lowe 2009). They can either live attached to the substrate, mobile on the substrate (benthic species), or suspended in the water column (planktonic). The planktonic species are not distributed evenly in the photic zone of oceans and lakes and are mostly found at a depth of 30–40 m in temperate and cold oceans and deeper in tropical oceans. Because the Diatomeae are nonmotile, they are subjected to currents and wave action to stay in the water column. When isolated from those forces, the silicified cell wall of the diatoms act as a weight and the cells sink. Some large species can maintain afloat in the water column by maintaining a large vacuole providing some buoyancy (Raven and Waite 2004). A major limiting factor for the growth of the diatom populations is the availability of silica, and blooms sometimes occurring in spring can be stopped by the total consumption of the silica resulting in the death of the majority of the diatoms (Lund 1949, 1950). The diversity and species number of benthic Diatomeae appear to be far larger than that of the planktonic, maybe driven by the diversity of the substrate they can grow on and/or the greater concentration of nutrients in the benthic habitats. Diatomeae species are highly adapted to their substrate and have specificities for each of these habitats. Briefly, epipelitic species have been shown to have circadian movements in and out of the sediment (Palmer and Round 1967; Round 1981); epipsammic species can fully cover sand grains; epiphytic species have developed various strategies to attach to the plant surfaces; metaphytic species

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Fig. 5.18 Life cycle of a pennate diatom. Upper right; division of the diploid vegetative cells by successive mitoses and diminution of the valve dimension. The slightly larger epitheca (left valve) overlaps the slightly smaller hypotheca (right valve). During cell division, only a smaller valve (hypotheca) is deposited within the protoplast so that successive divisions result in the size reduction of a part of the population (exemplified by right cell). Once a minimal size has been reached, the sexual reproduction (lower left) is initiated. After meiosis, non-flagellated gametes being capable of amoeboid movement are formed, they fuse, and form zygotes (auxospores) with the fusion of the nuclei and, finally the auxospore expands to reach the maximum cell size within mucilage produced by the two compatible cells. The various stages of sexual reproduction follow those observed for Pinnularia (Poulíčková et al. 2007). Original of Spindler & Friedl

have been reported in the mucilage produced by other algae; epizoic species have been found on the skin and feathers of various animals (e.g. cetaceans Denys and De Smet 2010; turtles Majewska et al. 2015); epilithic species and species

growing on ice have also been reported (Thomas and Dieckmann 2003). A famous result demonstrating the importance of the Diatomeae is the estimation that they contribute to

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Fig. 5.19 Diversity of freshwater diatoms. a examples of diverse shapes and sizes of genera of mostly pennate benthic diatoms from the Berlin-Brandenburg area, Germany; 1, Amphora; 2, Neidium; 3, Navicula; 4, Caloneis; 5, Cymbella; 6, Gyrosigma; 7, Gomphonema; 8, Hantzschia; 9, Epithemia; 10, Navicula; 11, Nitzschia; 12, Cymatopleura; 13, Tabellaria (zig-zag shaped colony of four cells); 14, Diatoma; 15, Anomoeoneis; 16, Craticula; 17, Aneumastus; 18, Cocconeis; 20, Karayevia; 19, Nitzschia; 21, Cymatopleura; 22, Cymbopleura; 23, Hydrosera; b scanning electron micrograph of a valve of the pennate diatom Cocconeis placentula; it is viewed from the inner side; note central raphe, and the girdle band attached to the valve; c, d examples of centric diatoms; c Stephanodiscus sp. in valve view; d scanning electron micrograph of Thalassiosira sp. in girdle view; note girdle bands that link both valves. a–d Courtesy of Research Group Diatoms, Bo Berlin, Freie Universität Berlin, modified from Zimmermann et al. (2021), www.biuz.de, VBiO e.V. under the CC-BY-SA 4.0 license

approximately 20% of the total global C-fixation (Mann 1999). Furthermore, they are crucial primary producers and had a considerable role in the evolution of life on Earth (Falkowski and Knoll 2007; Renaudie 2016). They also have practical importance for human societies, and they are widely used for biomonitoring of water quality and ecology (e.g. Szczepocka and Żelazna-Wieczorek 2018). Deposits

resulting from the sedimentation of a large number of Diatomeae, named diatomite (Fig. 5.21), can be used in industries (Smol and Stoermer 2010). The ability of forming complex and refined silicified frustules at conditions compatible with life is at the center of interest of many engineers hoping to develop new methods for synthesizing silica-based materials (De Tommasi et al. 2017). In counterpart,

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Fig. 5.20 Morphological diversity of centric diatoms from marine phytoplankton representing four different phylogenetic lineages (classes), i.e. Corethrophyceae (a, b), Mediophyceae (c–f), Leptocylindrophyceae (g), and Biddulphiophyceae (h); a filamentous colony of Melosira octogona, optical section (left) and surface (right) of girdle view of the same colony, culture strain CCMP 483; b cleaned frustule of Coscinodiscus wailesii, culture strain CCMP 2513, scanning electron micrograph; c Thalassiosira nordenskioeldii, culture strain CCMP 997, scanning electron micrograph; d Odontella aurita, culture strain NIES-582, the upper two left cells are connected to a short chain; note valve with horns, nucleus (nu), girdle view; e Ditylum brightwellii, culture strain NIES-350; note rectangular cells with numerous small chloroplasts and central nucleus (nu) in girdle view, a central long tubular process connects two cells; f Chaetoceros socialis, culture strain NIES-377, rectangular cells with long setae, connected to form curved chains, cells with a single chloroplast, girdle view; g Leptocylindrus danicus, cylindrical cells connected to a chain, numerous elongated chloroplasts, girdle view; h Biddulphia mobiliensis, single cell with long and narrow horns and spine-like processes that diverge, numerous chloroplasts, girdle view (dark field microscopy); a, b courtesy of Robert A. Andersen, with permission of the CCMP culture collection; c courtesy of I. Kaczmarska, with permission of CCMP culture collection with permission of the CCMP culture collection; d–f courtesy of Microbial Culture Collection, National Institute for Environmental Studies, Tsukuba, Japan; g https://www.flickr.com/photos/noaaphotolib/ 16087258835/in/photostream/lightbox/; h courtesy of John R. Dolan, CNRS, France with permission of Observatoire Océanologique de Villefranche-sur-Mer, France

Diatomeae have relatively few undesirable effects. Their overgrowth can cause trouble as they can obstruct filtration systems, and species of the genera Nitzschia and Pseudo-nitzschia are known for producing the neurotoxin domoic acid that can cause “amnesic shellfish poisoning” (Trainer et al. 2012). Evolutionary history Due to the formation of their silicified frustule and their diversity, the fossil record of the Diatomeae is one of the most abundant among the Heterokontophyta (Sims et al.

2006; Harwood et al. 2007). The earliest fossil dates from the Early Jurassic (ca. 190 Mya; Rothpletz 1900) and the extensive fossil record of the Paleocene and Neogene (ca. 66 to 2.6 Mya) was used for calibration of molecular clock that helped estimate the divergence between the important lineages of the Diatomeae and placed their origin somewhere in the Mesozoic or late Paleozoic (Sorhannus 2007; Medlin 2015). The origins and the long evolutionary history of the Diatomeae through geological time have been the source of many publications (e.g. Sims et al. 2006; Kooistra et al. 2007; Medlin 2011, 2016). Recently, it was shown that the Diatomeae experienced a burst of diversification in the late

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Fig. 5.21 a deposit of diatomite (diatomaceous earth, kieselguhr, white arrow) at the volcanic Vogelsberg region, Germany, near the worldwide first mining location in 1855. The deposit was formed from a freshwater lake under subtropical/tropical conditions in a volcanic maar, roughly 18 Million years ago. b Scanning electron micrograph of kieselguhr (not from the Vogelsberg deposit, source unknown) showing the fragmented empty frustules of diatom; a courtesy of Hartmut Kraus, Homberg/Ohm; b courtesy of Dawid Siodłak, Opole University, from Wikipedia

Eocene, with different rates among the different lineages (Lewitus et al. 2018). However, re-investigation of marbles from the Upper Proterozoic (625–550 Mya) revealed that they contain fossils very closely resembling diatom frustules (reviewed in Siemińska 2015). If confirmed, these fossils will greatly reshape our understanding of the evolution of the diatoms.

5.1.4.5 Raphidophyceae The class Raphidophyceae currently regroups ten genera that are found in either freshwater or marine habitats. All known species are considered unicellular flagellates, sometimes producing cysts when conditions are unfavorable. The name refers to the numerous trichocysts present in some Raphidophyceae species (Greek raphid—needle).

refers to Raphidomonas (Stein 1878), which is an illegitimate name for Gonyostomum and is not in current use. During the twentieth century and until recently, several brown-colored marine genera were described, and the class now contains 10 genera. The marine taxa include Chattonella (Biecheler 1936), Fibrocapsa (Toriumi and Takano 1973), Heterosigma (Hara and Chihara 1987), Haramonas (Horiguchi 1996), Chlorinimonas (Yamaguchi et al. 2010), Viridilobus (Demir-Hilton et al. 2012), and Psammamonas (Grant et al. 2013). The genus Chattonella proved to be polyphyletic (e.g. Bowers et al. 2006; Kamikawa et al. 2007; Demura et al. 2009; Klöpper et al. 2013), and one species was transferred to the Dictyochophyceae (Hosoi-Tanabe et al. 2007; Chang et al. 2012). General description

Brief history and current taxonomy Three yellow-green freshwater taxa, Gonyostomum (Diesing 1865), Vacuolaria (Cienkowsky 1870), and Merotricha (Mereschkosky 1879), were described in the second half of the nineteenth century. They were first combined as the Chloromonadina (Klebs 1892a, b), then as the order Chloromonadales (Engler 1898), which then became a component of the class Heterokontae by Luther (1899, Box 5.1). When Pascher (1914) limited the Heterokontae to what today we call the Xanthophyceae, this left the Chloromonadineae as an uncertain group. It later became clear that this group should be recognized as a class (e.g. Chadefaud 1950) and after some invalid names, the Raphidophyceae (Silva 1980) was validly published. Interestingly, the name

Raphidophyceae cells are relatively large in size, ranging from 10 to 80 µm, and they are ovoid, pyriform, or spherical in shape (Fig. 5.22). The cells are naked; no scales or cell walls have been observed in the Raphidophyceae. In some species, mucilage can be observed around the cells, produced by mucocysts and trichocysts (Horiguchi and Hoppenrath 2003; Yamaguchi et al. 2008; Klöpper et al. 2013). There is a clear coloration difference between the freshwater and marine species (Fig. 5.22). The freshwater species are usually bright green and the marine species light brown-yellowish. This coloration is due to the numerous and large (up to 3
5 µm) chloroplasts found in the periphery of the cell and variation in the photosynthetic pigments

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Fig. 5.22 Diversity of the Raphidophyceae. a–b freshwater Raphidophyceae; c–f marine Raphidophyceae. a Merotricha bacillata (strain NIES-1809); b Gonyostumum semen (strain NIES-1380); c Chattonella marina (strain NIES-113); d Haramonas dimorpha (strain NIES-716); e Heterosigma akashiwo (strain NIES-6); f Fibrocaspa sp. (strain NIES-1378); a–f courtesy of Microbial Culture Collection, National Institute for Environmental Studies, Tsukuba, Japan

between freshwater and marine species. The chlorophylls a, c1, and c2 are found in all species, but the accessory pigments vary (Table 5.2). Freshwater species lack fucoxanthin; diadinoxanthin is the major xanthophyll and heteroxanthin, vaucheriaxanthin, and ß-carotene are also present (Bjørnland and Liaaen-Jensen 1989). Conversely, most marine species contain fucoxanthin as well as violaxanthin, zeaxanthin, and 19' butanoyloxyfucoxanthin (Jeffrey et al. 2011). An exception is the yellow-green, marine Chlorinimonas sublosa that lacks fucoxanthin and contains diadinoxanthin (Yamaguchi et al. 2010). No eyespots were observed in the Raphidophyceae, and there was no flagellar swelling (Kawai and Inouye 1989). Flagella are inserted at the apex of the cell or in a small depression near it, and are typical of the Heterokontophyta. The long immature flagellum bears tripartite hairs and beats rapidly while the short mature flagellum is naked and trails, beating infrequently (Fig. 5.23). Ultrastructural analysis of the Raphidophyceae revealed characteristic Heterokontophyta traits such as mitochondria with tubular cristae, lamellae with three stacked thylakoids and a girdle lamella (in most taxa), and the chloroplast outermost membrane continuous with the endoplasmic reticulum. A pyrenoid was observed in the chloroplast of marine species (including Chlorinimonas) but not in those of the freshwater species.

Fig. 5.23 Drawing of Fibrocapsa japonica. Note the typical chloroplast arrangement. AF = immature flagellum, BB = basal body, Ch = chloroplast, G = Golgi body, M = mitochondrion, MC = mucocyst, Nu = nucleus, PF = mature flagellum, PL = plasmalemma, Py = pyrenoid, and V = vacuole (from Hara and Chihara 1985)

Raphidophyceae usually reproduce asexually by mitosis but under stressful conditions sexual reproduction has been reported (Yamaguchi and Imai 1994; Cronberg 2005;

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Figueroa and Rengefors 2006). The life cycle of Raphidophyceae consists of an alternation of generations, when vegetative cells undergo environmentally triggered meiosis. Haploid cells will differentiate into cysts that sink and can remain dormant in the sediment for several months. When the conditions become favorable, the cysts germinate into motile haploid cells that will fuse to re-establish the diploidy. This cycle usually covers a year with meiosis and encystment occurring in late summer and the germination and fusion of gametes in the spring (Imai and Itoh 1987). Occurrence The Raphidophyceae are found in both, marine and freshwater habitats and are cosmopolitan, being found on all continents and their coastlines except Antarctica (reviewed in Horiguchi 2017). The different genera and species of the Raphidophyceae have various and specific ecological preferences that range from cold water (4 °C) to hot water (32 °C), acidic (pH 3.2) to alkaline (pH 8.3), low salinity to hypersalinity (41‰), and many other ecological conditions (Horiguchi 2017). Most species are planktonic, but some are sand-dwelling species (Demir-Hilton et al. 2012; Grant et al. 2013). Despite this wide ecological tolerance, species of the Raphidophyceae are capable of daily vertical migration demonstrating their ability to respond and adapt rapidly to unfavorable conditions (Wada et al. 1987). All the Raphidophyceae possess functional chloroplasts, but phagocystis of bacteria (i.e. mixotrophy) has been reported for several genera, Chattonella, Fibrocaspa, and Heterosigma (Jeong 2011). The Raphidophyceae are infamous for their ability to kill fish (i.e. ichthyotoxicity) through mechanisms that are still poorly understood. Toxicity was proposed to be caused by brevetoxin compounds (Khan et al. 1997), by reactive oxygen species (ROS) causing damage to the gills (Ishimatsu et al. 1996; Hiroishi et al. 2005), by toxic polyunsaturated fatty acids (Marshall et al. 2002) or by a combination of these (Marshall et al. 2003; Dorantes-Aranda et al. 2015). Blooms of Raphidophyceae cause significant economic losses (Taylor 1992). Evolutionary history There is no fossil record of the Raphidophyceae, probably due to their naked single cells. Therefore, the evolutionary history of the Raphidophyceae was inferred from phylogenetic analysis (Yamaguchi et al. 2010; Demir-Hilton et al. 2012; Grant et al. 2013). It appears that the common ancestor of the Raphidophyceae was a marine alga and that the transition to freshwater habitats occurred once in the common ancestor of the clade formed by Merotricha, Gonyostomum, and Vacuolaria. The evolution of the

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photosynthetic pigments is more complex. The ancestor of the Raphidophyceae likely contained the violaxanthinzeaxanthin cycle; it was replaced in the common ancestor of the freshwater lineages by the diatoxanthindiadinoxanthin cycle. However, the marine genus Chlorinimonas also presents the same replacement of pigments but is not related to the freshwater lineages. This suggests that two independent replacements occurred during the evolution of the Raphidophyceae.

5.1.4.6 Eustigmatophyceae The Eustigmatophyceae was shown to be distinct from the Xanthophyceae (Hibberd and Leedale 1970). The large characteristic eyespot observed in the zoospores was used to name the class (Latin stigma – mark, brand). Brief history and current taxonomy During the early twentieth century, numerous new species and genera were described and classified, including Chlorobotrys, Ellipsoidion, Pleurochloris, and Vischeria. During his thesis, Dr. Hibberd studied the morphology and ultrastructure of the zoospores of 15 of these coccoid algae species. While some presented zoospores typical of the Xanthophyceae (e.g. Botrydiopsis, Bumilleriopsis), others were clearly different (e.g. Chlorobotrys, Ellipsoidion, Pleurochloris, or Vischeria). That is, the large orange-red eyespot was outside the chloroplast, and there was a swelling near the base of the flagellum (Hibberd and Leedale 1970, 1972). Although originally classified in the Xanthophyceae (e.g. Pascher 1937–1939), these algae were placed into a distinct class, the Eustigmatophyceae (Fig. 5.2; Hibberd and Leedale 1970). The pigment composition of these algae further supported their distinction from the Xanthophyceae (Whittle and Casselton 1969, 1975a, b). In the years following the establishment of the class, important taxonomic work (Hibberd 1981) was accompanied with the transfer of more taxa from other classes to the Eustigmatophyceae, such as Trachydiscus (Přibyl et al. 2012) or Tetraëdriella (Fawley and Fawley 2017) and the description of new genera such as Pseudellipsoidion (Neustupa and Němcová 2001) or Vacuoliviride (Nakayama et al. 2015). The class now comprises 18 genera, but recent studies using environmental DNA have unrevealed an unexpected diversity that will need to be explored in the future (Fawley et al. 2014). General description Vegetative cells range in size from 2 to 25 lm and present a variety of shapes from spherical, ovoid, and fusiform to discoid (Fig. 5.24). Their color is typically green to yellow-green. The cells are usually free floating and rarely attached (e.g., Pseudocharaciopsis). The easiest and most

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20 µm Fig. 5.24 Diversity of the Eustigmatophyceae (light microscopy). a Vischeria vischeri (strain SAG 860–1). Upper half, round vegetative cells; lower half, 4-celled autosporangia and elliptic vegetative cell; b V. polyphem (strain SAG 38.84), vegetative cells with a 2-celled autosporangium in the upper-left corner; c Nannochloropsis australis (strain CS-416), upper cells are mature and appear round while bottom cells are immature and angular; d N. salina (strain SAG 40.85), vegetative cells; e Monodopsis subterranea (strain SAG 848–1), vegetative cells; note empty cell walls; f Characiopsis acuta (strain SAG 14.95), vegetative cells with stalk; g Pseudostaurastrum limneticum vegetative cell; h Vischeria stellata (strain SAG 887–2). Note the cell wall projections on the cells in the right and bottom half of the figure, a 2-celled autosporangium on the lower-left corner; i Chlorobotrys regularis, vegetative cells; j Monodopsis subterranea vegetative cells. a, b, h from Kryvenda et al. (2018); c from Fawley et al. (2015); d, e courtesy of Anastasiia Kryvenda, with permission of the SAG culture collection, Göttingen University, Germany; f courtesy of Tatyana Darienko, with permission of the SAG culture collection, Göttingen University, Germany; g, i, j from Ott et al. (2015)

definitive trait to recognize algae of the class Eustigmatophyceae is the presence of a circular reddish globule in the cytoplasm (Fig. 5.24a–h). This body can occupy an important part of the cytoplasm in older cells, and its color varies from yellow–brown to red-brown. It can be easily observed under ultraviolet light where it has a yellow fluorescence (Přibyl et al. 2012). The reddish globule is an irregular aggregation of droplets not enclosed in a single membrane. The cytoplasm also contains a vacuole with granular contents presenting Brownian movements. A single chloroplast is observed in most cells but some genera present multiple chloroplasts as in Pseudoellipsoidion (Neustupa and Němcová 2001) or Pseudocharaciopsis (Hibberd 1981).

Vegetative cells in Eustigmatophyceae form a cell wall that is generally smooth but can present ornamentations as in Vischeria stellata (Fig. 5.24h, Přibyl et al. 2012; Fawley and Fawley 2017). The zoospores of the Eustigmatophyceae are naked cells of elongated oval shape with generally one visible flagellum but two unequal flagella inserted near the apex of the oval (Fig. 5.25). They possess one chloroplast and, as in vegetative cells, a distinctive red–orange eyespot that can be really large and fill the anterior part of the zoospore. Ultrastructural analysis of Eustigmatophyceae algae revealed both characters unifying them with the Heterokontophyta and some specific characters. As in other

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Fig. 5.25 Drawing of the zoospore of Pseudocharaciopsis texensis. C = chloroplast; CE = chloroplast envelope; CER = chloroplast endoplasmic reticulum; E = eyespot; F1 = immature flagellum; F2 = mature flagellum; F1B = basal body of the immature flagellum; F2B = basal body of the mature flagellum; FS = flagellar swelling; G = Golgi body; H = tripartite hair; L = layered material; LV = laminate vesicle; M = mitochondrion; MT = microtubules; N = nucleus; NE = nuclear envelope; OV = osmiophilic vesicles (from Lee and Bold 1973)

Heterokontophyta, the mitochondria contain tubular cristae. The anterior flagellum possesses tripartite tubular hairs; a second flagellum, if present, is smooth and stiff. Uniflagellate zoospores have a second basal body that remains after flagellar transformation. Unlike other Heterokontophyta, the chloroplast of the Eustigmatophyceae does not form a girdle lamella. Also, the plastid endoplasmic reticulum is not connected to the nucleus except in Microchloropsis, Monodopsis, and Nannochloropsis. Photosynthetic pigments are also uncommon for the Heterokontophyta because chlorophyll c is absent and only chlorophyll a is present (Table 5.2). The principal accessory pigments are ß-carotene, violaxanthin, and vaucheriaxanthin. Other minor pigments are sometimes present such as zeaxanthin, canthaxanthin, and astaxanthin, but fucoxanthin, diadinoxanthin/diatoxanthin, or heteroxanthin are not present. Sexual reproduction has yet to be reported for the Eustigmatophyceae. Vegetative reproduction occurs by the formation of autospores or zoospores.

The Eustigmatophyceae are cosmopolitan. Most are found in freshwater lakes and ponds, but an important part of their diversity has also been reported from soils. The genera Eustigmatos, Monodopsis, Pseudocharaciopsis, and Vischeria have been isolated both from freshwater and soils, rocks, and even desert crusts (e.g. Neustupa and Němcová 2001; Büdel et al. 2009; Czerwik-Marcinkowska and Mrozinska 2009; Neustupa and Škaloud 2010; Ott et al. 2015). The genera Nannochloropsis and Microchloropsis are the only ones found in marine or brackish habitats, where they can form blooms (Andreoli et al. 1999; Fawley et al. 2015; Zhang et al. 2015). Freshwater species frequently occur in mesotrophic and eutrophic bodies of water with neutral to slightly alkaline pH (Fawley et al. 2014). But many species have been reported from acidic Sphagnum bogs (Hibberd 1974; Lara et al. 2011; Ott et al. 2015; Fawley and Fawley 2017). Association with vegetation is not uncommon, and an unidentified Eustigmatophyceae species have been reported to be the endosymbiont of a freshwater sponge (Frost et al. 1997). Other unusual reports of Eustigmatophyceae species include the eutrophic cooling pond of a nuclear plant (Přibyl et al. 2012), heavy metal-polluted calamine mine spoils (Trzcińska et al. 2014), Antarctic ice-covered lakes (Bielewicz et al. 2011), or even in a bottle of glue (Nakayama et al. 2015). While the Eustigmatophyceae have received great interest for academic reasons, their commercial utilization has led to a surprisingly large literature and genomic resource (reviewed in Eliáš et al. 2017). The various forms of triaglycerol and long-chain polysaturated fatty acids synthesized by species of the genus Nannochloropsis (and Microchloropsis) have been of great interest for the biofuel and biotechnological industries (reviewed in Ma et al. 2016). The production of other bioproducts of interest has been investigated too, like sterols (Patterson et al. 1994); vitamin E (Durmaz 2007); or more generally as a food source in aquaculture (Pfeiffer and Ludwig 2007; Patil et al. 2007; Ferreira et al. 2009). Finally, the ability of the Eustigmatophyceae to survive in polluted water (see above) has led to their use in bioremediation of heavy metals (Moreno-Garrido et al. 2002), Caesium radioactivity (Fukuda et al. 2014), and arsenic toxicity (Upadhyay et al. 2016). Evolutionary history Because of the absence of Eustigmatophyceae fossil record, the evolutionary history of this class was only inferred from molecular phylogenies and comparative analyses. Recent

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phylogenetic analyses have resolved most of the relationships within the Eustigmatophyceae (Přibyl et al. 2012; Fawley et al. 2014) despite the absence of some lineages and still some uncertainty in the early diverging lineages. Based on this phylogenetic framework, it is now thought that the characteristic large eyespot placed outside the chloroplast was absent from the common ancestor of the Eustigmatophyceae. Other unusual features such as the non-continuity chloroplastic endoplasmic reticulum and the nuclear envelope or the reduction of the posterior flagellum appear to have been derived multiple independent times (reviewed in Eliáš et al. 2017).

5.1.4.7 Dictyochophyceae The Dictyochophyceae are a distinct group of the Heterokontophyta. The name of this class refers to the highly reticulated cytoplasm (Greek dictyon—net) of the first alga described in this class, Dictyocha. Brief history and current taxonomy Interestingly, the majority of the diversity is only known through fossils, and the first Dictyochophyceae alga was described based on fossil material (Ehrenberg 1839). Before the first living Dictyochophyceae was isolated, much fossil species were described based on their silicified structures and they formed the Silicoflagellates (= Dictyochales), an order placed in the Chrysophyceae (Deflandre 1950). Other newly described algae were tentatively placed in orders of the Chrysophyceae like Rhizochromulina (Hibberd and Chretiennot-Dinet 1979) of the order Rhizochromulinales, or Pedinella (Vysotskii 1887) and Pteridomonas (Penard 1890) of the order Pedinellales. These orders were later combined to form the class Dictyochophyceae (Silva 1980), and a new order was described in this class recently, the Florenciellales (Eikrem et al. 2004). General description The Dictyochophyceae are mostly unicellular algae, with a typical size of less than 10 µm in the largest dimension. The cells can have different shapes, from the amoeboid cells of the order Rhizochromulinales, the radially symmetrical cells of the order Pedinellales, the irregular cell of the order Dictyochophyceae, or the bumpy cells of the order Florenciellales (Fig. 5.26). It is common for cells of the Dictyochophyceae to have tentacles-like cytoplasmic extensions, rhizopodium, or pseudopodium. In the order Pedinellales, a long trailing stalk can extend from the posterior end of the cell and sometimes anchor the cell to a substrate.

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The cells are generally golden-brown when chloroplasts are present. The number of chloroplasts per cell can vary greatly among the lineages and the life cycle stage, but generally a multitude of chloroplasts can be observed in the cells, and they never present an eyespot. The ultrastructural features of the Dictyochophyceae are generally typical of the Heterokontophyta, with a chloroplast enclosed in four membranes, the outermost being continuous with the nuclear envelope, a girdle lamella enclosing the three stacked thylakoids, tubular cristae in the mitochondria. The chloroplasts contain the chlorophyll a, c1, and c2 as primary photosynthetic pigments and fucoxanthins, diatoxanthin/diadinoxanthin as secondary pigments. The Dictyochophyceae of the order Dictyocales are characterized by the formation of an external silicified skeleton (Lipps 1979; Moestrup and Thomsen 1990). The shape of this skeleton is species-specific and varies from a simple ring or triangle but is often complex and composed of multiple different star-like shapes (Fig. 5.27). In the order Pedinellales, cells can be covered by unmineralized scales and spined scales can be present. The number of flagella per cell varies between the different lineages of the Dictyochophyceae. In most species, only a single immature flagellum is present, but the basal body of the mature flagellum is present. The immature flagellum is typical of the Heterokontophyta and bears tripartite hairs. In the order Florenciellales however, a smooth mature flagellum is present along the immature long flagellum (Edvardsen et al. 2007). As most of the single-celled Heterokontophyta, algae of the Dictyochophyceae generally present an alternation of two cell types, vegetative cells and flagellated cells. In the order Dictyotales, a more complex life cycle is observed and was carefully described (Fig. 5.28; Henriksen et al. 1993; Chang et al. 2012). During this cycle, large multinucleated cells are formed by the differentiation of silica skeleton-forming cells. These amoeboid cells will then release uninucleated flagellated cells not forming silica skeleton. However, sexual reproduction has not been described in any Dictyochophyceae. Occurrence The Dictyochophyceae have been primarily described from marine ecosystems, but they are not limited to them and some species are found in freshwater (e.g. Pseudopedinella). They are found in all oceans, and recent metabarcoding analysis revealed that they are cosmopolitan (de Vargas et al. 2015; Carradec et al. 2018). However, some species, notably of the order Dictyochales, appear to be more abundant at high latitudes in colder waters and in some cases

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Fig. 5.26 Diversity of the Dictyochophyceae. a Light microscopy of Pseudopedinella elastica (strain CCMP 716). b Light microscopy of Florenciella parvula (CCMP 2471). c Light microscopy of Rhizochromulina sp. d Light microscopy of the skeleton-bearing life stage of Distephanus speculum (isolated in East Greenland). The cells present the typical six spines of the skeleton and the single emergent flagellum (arrow). a–c from Han et al. (2019), d from McCartney et al. (2014b)

temperatures above 8–10 °C are lethal (e.g. Mesopedinella, Daugbjerg 1996). Upon favorable conditions, they can form blooms in marine to brackish waters from cold (2–5 °C) to warmer temperatures (e.g. Henriksen et al. 1993; Edvardsen et al. 2007). Some of these blooms have been associated with fish death, notably in fish farms, due to the production of an ichthyotoxic substance (Edvardsen et al. 2007; Chang et al. 2014). The Dictyochophyceae can be either exclusively photosynthetic, mixotrophic, or heterotrophic. Evolutionary history The evolution of the traits described above (i.e. ecology and mode of nutrition) has started to be investigated with the development of molecular phylogeny and the discovery of new species (e.g. Edvardsen et al. 2007; Chang et al. 2017). These suggest that mixotrophy evolved in the ancestor of the Rhizochromulinales and Pedinalles. In these orders, some genera (e.g. Ciliophrys; Pteridomonas) have lost the ability to fix inorganic carbon through photosynthesis.

5.1.4.8 Pelagophyceae The pelagophytes are a group of strictly marine algae that are distinct from other Heterokontophyta, notably the chrysophyte (Andersen et al. 1993). The class was named in

reference to pelagos (Greek pelagos - sea) as the earlydescribed taxa (Pelagococcus, Pelagomonas) occurred in the open oceans. Brief history and current taxonomy In the twentieth century, numerous marine taxa of uncertain affiliation were described such as Aureococcus (Sieburth et al. 1988), Pelagococcus (Lewin et al. 1977), and Sarcinochrysis (Geitler 1930). Despite some significant differences, notably of the zoospore morphology, they were included in the class Chrysophyceae. Based on electron microscopic observations of the flagellar apparatus and flagellar hair ultrastructure, a number of these taxa were grouped in the order Sarcinochrysidales (Chrysophyceae) by Gayral and Billard still within the Chrysophyceae (1977a). Investigation of the alga Pelagomonas revealed that its single flagellum covered by bipartite hairs and the structure of its flagellar apparatus were unique and clearly different from that of chrysophyte algae (Andersen et al. 1993). This led to the description of the class Pelagophyceae and the separation of the genera Aureococcus, Pelagomonas, and Pelagococcus from the Chrysophyceae (Andersen et al. 1993; Saunders et al. 1995). With the development of molecular analysis, the unsatisfying position of the Sarcinochrysidales within the Chrysophyceae was resolved with

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Fig. 5.27 Scanning electron microscopy (SEM) photographs of silicified skeletons of Dictyochophyceae. a–d Skeletons of Distephanus speculum, with descriptive morphological terminology. a, b Parts of skeletons. c, d Full skeletons. e Full skeleton of Octactis pulchra from the Seto Inland Sea. f Skeleton of Distephanus crux (Ehrenberg) Haeckel, a fossil Dictyochophyceae from the late Eocene, Oamaru, New Zealand. Modified from McCartney et al. (2014b)

the demonstration of its genetically close relation with pelagophytes, and it was therefore recognized as an order of the class Pelagophyceae (Saunders et al. 1997). From this point on, numerous genera were described and included in this class, like Andersenia (Wetherbee et al. 2015), Aureoumbra (DeYoe et al. 1997), Aureoscheda (Wynne et al. 2014), or Chrysocystis (Lobban et al. 1995). The recent

rediscovery of the alga Sarcinochrysis marina from its type locality in the Canary Islands allowed to firmly anchor the genus Sarcinochrysis in modern taxonomy and also to establish four new genera and a new family (Han et al. 2018). The class Pelagophyceae is deeply divided into two orders: Pelagomonadales and Sarcinochrysidales, and currently contains 18 genera.

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Fig. 5.28 Life cycle of Distephanus speculum (Dityochophyceae). a Skeleton-bearing cells. b Cells connected by cytoplasmic bridges. c Large spherical cells. d Naked cells. e Transition stage. f Multinucleate cell. The illustrations are not to scale. Note that the cycle is “incomplete” as the skeleton-bearing cells are not re-established from other cell types (from Henriksen et al. 1993)

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General description Generally, Pelagophyceae are golden-brown in color (Fig. 5.29). This is due to the photosynthetic pigments contained in their chloroplast (generally 1 or 2 per cell): the chlorophylla, chlorophyllc1, and Chlorophyllc2, fucoxanthin, diatoxanthin-diadinoxanthin, and other accessory pigments. The chloroplasts are typical of the Heterokontophyta with a girdle lamella and a chloroplastic endoplasmic reticulum but without an eyespot. Beyond these ultrastructural characters, it is hard to determine characters to unify the orders Pelagomonadales and Sarcinochrysidales of the Pelagophyceae. The two orders present many differences, notably concerning their respective flagella, and, for clarity, they will be described separately. However, recently the presence of a perforated theca was proposed as a characteristic feature of the Pelagophyceae (Wetherbee et al. 2021). The Pelagomonadales (Fig. 5.30) are single-cell organisms of very small size (e.g. Pelagomonas 1.5
3 µm; Aureococcus 2 µm) that are either flagellate (Pelagomonas, Ankylochrysis) or coccoid (Aureococcus, Pelagococcus). The typical flagella of the Heterokontophyta are present in Ankylochrysis but in Pelagomonas only a single immature flagellum is present. Furthermore, the Pelagomonas flagellar apparatus is distinct from other uniflagellated Heterokontophyta (e.g. Chrysophyceae; Pinguiophyceae) as the basal body of the mature flagellum is totally absent (Andersen

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et al. 1993; Heimann et al. 1995). The presence of a paraxonemal rod between the axoneme and the flagellar apparatus is also particular and is absent in Ankylochrysis (Honda and Inouye 1995). The Sarcinochrysidales (Fig. 5.31) present various morphologies: flagellates, nonmotile single cells, colonies, or filaments. Large colonies can be flowing and reach 3–5 cm (Chrysocystis) or form well-organized sheets of several cm2 (Aureoscheda). The smaller benthic species are mostly coccoid cells (Arachnochrysis, Chrysoreinhardia, Gazia, Glomerochrysis, Pelagospilus, Sarcinochrysis, Sargassococcus, and Sungminbooa), but there is one unbranched filamentous species (Andersenia). A gel usually surrounds the cells or colony (e.g. Han et al. 2018; Wetherbee et al. 2021), and a cell wall is generally present. Interestingly, in the genera Gazia and Glomerochrysis a theca encases the cells in every life stage (i.e. coccoid and zoospore) only letting the flagella out through a pore. Despite no clear description of a theca in other Pelagomonadales genera, a similar structure could have been overlooked in Aureoumbra, Aureococcus, Chrysonephos, Pelagococcus, Pelagomonas, and Sarcinochrysis (see Wetherbee et al. 2021). Zoospore production has been observed in most of the Sarcinochrysidales genera (except Aureoscheda, Aureoumbra, Sargassococcus, Sungminbooa), and zoospores serve to disperse the algae to a favorable microenvironment. The motile cells have two flagella typical of the Heterokontophyta: a long anterior

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b Fig. 5.29 Diversity of the Pelagophyceae. a Vegetative cells of Sarcinochrysis marina strain BEA 0109B with highly lobed chloroplasts

(arrowheads). b Cell of S. marina strain BEA 0109B showing the central nucleus (arrowhead) between the two chloroplasts. Note the gelatinous sheaths from mother cell walls. c Cluster of Sungminbooa caribensis strain CCMP 292. The cells are maintained in pairs with a gelatinous material. d Vegetative cells of Sargassococcus simulans strain CCMP 1996 showing large range in cell size. e Linear colony of S. simulans strain CCMP 1996 cells. Note the hematochrome droplet (arrow). f Filaments of Andersenia nodulosa showing differentiation and knot formation giving the filaments a gnarled appearance. Most cells in a filament divide in several planes to produce multicellular regions, the cells enlarging and eventually forming motile zoospores. Gnarled filaments are also active in producing extracellular mucilage, while some undifferentiated cells are present (arrowheads). g Perforate cluster of Sargassococcus epiphyticus strain CCMP 1995 cells showing the colonial habit found in culture tubes. h Early stage of Sargassococcus epiphyticus strain CCMP 1995 colony development showing cells in packets of 2–8 cells; the entire packet has 32 cells. i Solid, clump-like cell mass of Chrysoreinhardia giraudii strain BEA 0313B formed by adhering packets of cells (arrow). j Cluster of Arachnochrysis demoulinii strain CCMP 2350 showing the highly lobbed chloroplasts. k Two cells of A. demoulinii strain CCMP 2350. Top cell viewed from the top showing the plastid lobes meeting; bottom cell viewed from the side showing the two chloroplasts separated by the nucleus (N). Pyrenoid (arrowhead) is underneath upper two plastids. l Surface view of a mature zoospore showing the insertion of the anterior flagellum (arrow) and posterior flagellum (arrowhead). Note the sinusoidal wave of the anterior flagellum and the stiff sculling posterior flagellum. m A sheet-like growth of Aureoumbra geitleri strain BEA 0312B cells found floating on the water surface. n Cells of A. geitleri strain BEA 0312B showing the stalked pyrenoids (arrows). a–e and g–n from Han et al. (2018); f from Wetherbee et al. (2015)

Fig. 5.31 Drawings of Aureoumbra lagunensis. This alga is representative of the order Sarcinochrysidales, with stalked pyrenoid and two basal bodies near the nucleus (Adapted from DeYoe 1997 / Drawings from Lee 2018)

Fig. 5.30 Drawings of Pelagomonas calceolata. This alga is representative of the order Pelagomonadales with a single immature flagellum (from Andersen 1993 / Drawings from Lee 2018)

immature flagellum bearing tripartite hairs, a short posterior mature flagellum with no hairs. Occurrence Ecologically, pelagophytes are only reported from marine environments. However, despite what the name of the class suggests, most pelagophytes are not found in the pelagic zone, i.e. they are benthic and attached to various substrates including sand (Wetherbee et al. 2021). The pelagic species are important primary producers (Andersen et al. 1996). The coastal planktonic Aureococcus and Aureoumbra are famous for producing marine algal blooms or brown tides, reaching densities of 3.109 cells per liter. In addition, during the

blooms, light cannot penetrate the water column, which causes the destruction of submerged plant beds. Although the brown tide organisms do not produce toxins and are not directly harmful to humans, the blooms do have a negative impact on filter-feeding organisms. They have been linked to high mortality rates in scallops, oysters, or blue mussels that sometimes result in major economical losses (Nicholls 1995). The factors leading to the development of these blooms are still poorly understood, but it appears that increased concentration of phosphorus or nitrogen has no effect whereas input of iron might (Bricelj and Lonsdale 1997). Recently, sequencing of the genome of Aureococcus anophagefferens brought new insights into the genetics of this blooming species (Gobler et al. 2011). Compared to other non-blooming algae, gene families involved in light harvesting, carbon and nitrogen metabolisms, or anti-microbial activities were expended (i.e. contained more genes). Filamentous benthic pelagophytes such as Andersenia and Chrysoreinhardia have also been reported to form large-scale sheaths recovering the seafloor. Occurring from late spring and throughout the summer, these layers although

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only several mm thick can outcompete and replace macroalgae (Hoffman et al. 2000). Evolutionary history The Pelagophyceae have no known fossils, and their evolutionary history has been studied using phylogenies only. Pelagophyceae are members of the SIII clade (Yang et al. 2012; Derelle et al. 2016) and therefore related to other lineages with reduced or absent flagella. However, the presence of two flagella in the order Sarcinochrysidales and Ankylochrysis in the order Pelagomonadales questions a reduction in the common ancestor of the SIII clade. The re-emergence of a mature flagellum after its loss in the Pelagophyceae is unlikely but this was not tested. Furthermore, flagellated stages have not been observed in most genera of the Pelagomonadales, and therefore the unusual flagella of Pelagomonas might represent an exception and a derived character rather than an ancestral one. The recent discovery of the theca of Gazia and Glomerochrysis and its possible widespread presence in the Pelagophyceae could redefine the class. This perforated theca has been proposed as a unifying and distinct character (i.e. synapomorphy) of the Pelagophyceae. However, exploration of more genera using electron microscopy to validate the presence of a theca is needed.

5.1.4.9 Phaeothamniophyceae The phaeothamniophytes are another group of algae that was shown to be distinct from the chrysophytes (Craig Bailey et al. 1998). The class was named after the shrubby (Greek phaeo—gray, dusky and thamnion—small shrub) aspect of the alga Phaeothamnion. Brief history and current taxonomy For decades, the classification of filamentous algae was relatively straightforward for Phaeophyceae, Xanthophyceae, and diatoms. Other filamentous heterokonts were classified in the Chrysophyceae. Evidence that this classification was problematic was discovered in the late 70 s during ultrastructural studies of the flagella and flagellar hairs (Gayral and Billard 1977b). With the emergence of molecular data, these “chrysophyte” filaments were transferred to other classes: Pelagophyceae (see Sect. 5.1.4.8, Andersen et al. 1993; Saunders et al. 1997), Phaeothamniophyceae (here), and the Phaeosacciophyceae (see Sect. 5.1.4.16). Similarly, the freshwater filamentous alga Phaeothamnion was described in Lagerheim (1884) and subsequently was classified in the Chrysophyceae. However, when photosynthetic pigments, molecular and electron microscopy analysis revealed a unique combination of carotenoids, gene sequences, and ultrastructural features for Phaeothamnion, a new class was required. The class

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Phaeothamniophyceae was erected from the chrysophyte order Phaeothamniales (Bourrelly 1954) to acknowledge these differences. At this time, the class Phaeothamniophyceae contained a wide range of primarily freshwater species forming more or less loose colonies (e.g. Phaeoschizochlamys or Stichogloea) and filaments (e.g. Phaeothamnion). Further molecular analysis including larger number of taxa demonstrated the polyphyly of the class Phaeothamniophyceae (Kai et al. 2008; Yang et al. 2012). Consequently, the genera Phaeobotrys, Pleurochloridella, and Tetrasporopsis were removed from the class. However, until recently, the class Phaeothamniophyceae had remained relatively understudied with molecular techniques, and links between historical (pre-molecular) descriptions and modern taxa were missing. In a large study, the genera Chrysoclonium, Phaeoschizochlamys, Phaeothamnion, Stichogloea, Tetrachrysis, and Tetrapion were re-investigated (Graf et al. 2020a). This study allowed to clarify the taxonomy of the class Phaeothamniophyceae and a better understanding of the evolution within this class. However, re-collecting species from their type localities is still necessary to definitively anchor names with biological specimens and validate the phylogeny of the class. General description Morphologically, algae of the class Phaeothamniophyceae form filamentous or loose colonies of various shapes and sizes, usually attached to a substrate (Fig. 5.32). Filaments of Phaeothamnion elongate from a single basal cell and are formed of 2 to 5 initial branches from the basal cell depending on the species. The branches are constituted of elongated vegetative cells usually 6 µm wide and 15 µm long, and the total size of the ranges from 1 to 5 mm. The vegetative cells are usually spherical to slightly ovoid in colonies of Phaeoschizochlamys, Stichogloea, or Tetrachrysis, and their size usually ranges from 4 to 12 µm. Cells have a cell wall and during division, the daughter cell wall is formed within the mother cell wall. The mother cell wall is then either cast off (e.g. in Phaeoschizochlamys), retained around the daughter cell (e.g. in Stichogloea), or surrounds all the cells in Phaeothamnion (i.e. intercalary growth). Furthermore, in all Phaeothamniophyceae, a gelatinous envelope surrounds the filaments or colonies and probably plays a role in the attachment to the substrate. Sexual reproduction is unknown in the Phaeothamniophyceae, and dispersion happens with zoospores or colony and filament fragmentation. Zoospores are released through lateral pores in the cell wall of the zoosporangium. They have two flagella inserted laterally, generally with one chloroplast but with two chloroplasts in some species, and with or without an eyespot. In vegetative cells, too, the number of chloroplasts varies from species to species, but

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Fig. 5.32 Diversity of the Phaeothamniophyceae. a Highly branched filament derived from a single basal cell of Phaeothamnion confervicola strain A15, 092. b Basal cell of P. confervicola strain A15,092 with two branches. Not the ear-like view of the gelatinous pad (arrowhead). c A zoospore of P. wetherbeei strain CCMP 637 showing the immature flagellum (long arrow), the mature flagellum (arrowhead), and the eyespot (short arrow). A second eyespot is also visible (double arrowheads). d A zoospore of P. wetherbeei strain CCMP 637 showing the longer immature flagellum (arrow) and the shorter mature flagellum (arrowhead). e Vegetative cells of Phaeoschizochlamys siveri strain CCMP 635 showing “grandmother” (large arrowhead), “mother” (small arrowheads), and “daughter” cell walls (small arrow). f Vegetative cells of P. santosii strain ACOI-284 with cup-like half-wall silhouettes (arrows). g Dividing cells in a colony of Stichogloea fawleyi strain CCMP 2289 that is lightly stained with brilliant cresyl blue to show the colonial gel (arrows). h Cells of S. dopii strain ACOI-338 showing the 4-cell groups; the typical linear arrangement is disrupted by the cover-slip. i Cells of S. dopii strain ACOI-338 showing dumbbell-like bilobed chloroplasts (arrows). Modified from Graf et al. (2020)

generally a single highly lobbed chloroplast is present. Photosynthesis pigments of the class Phaeothamniophyceae are chlorophyll a and c and the diadinoxanthin-diatoxanthin carotenoid cycle, as well as fucoxanthin and heteroxanthin (Table 5.2). Occurrence Phaeothamniophyceae appear to be widespread and have been reported from freshwater bodies around the world (e.g.

Starmach 1985; Kristiansen and Preisig 2001; Nicholls and Wujek 2015). These are generally attached to different substrates and have been reported as epiphytes of Sphagnum but others are planktonic like some species of Stichogloea. However, so far there is no clear understanding of the ecological niche occupied by the Phaeothamniophyceae and their global repartition. The toughness of their cell wall prevents easy access to their DNA maybe explaining why there are no environmental DNA sequences reported for this class.

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5.1.4.10 Bolidophyceae The class Bolidophyceae was described following the isolation of two small algae from the Pacific Ocean and the Mediterranean Sea (Guillou et al. 1999). These algae were rapid swimmers, and the class was named to acknowledge this behavior (French bolide - racing car). Brief history and current taxonomy The first algae described in this class (i.e. genus Bolidomonas) were picoplanktonic unicellular naked cells (Fig. 5.33d). Molecular and phylogenetic analysis placed them as a sister group to the diatoms (Guillou et al. 1999; Daugbjerg and Guillou 2001). Initially, the absence of any form of silicified structure was puzzling and brought insights into the early evolution of the diatoms. However, this discussion changed radically with the first molecular analysis of algae of the order Parmales. These algae (Fig. 5.33a–c) were historically classified in the

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Chrysophyceae but were demonstrated to be closely related to Bolidomonas and transferred to the Bolidophyceae (Ichinomiya et al. 2011). Even more surprising, further genetic analysis showed that members of the Parmales and the genus Bolidomonas actually formed a monophyletic group and that Bolidomonas and Triparma were synonymous, i.e. alternating life forms of the same organisms (Ichinomiya et al. 2016). Despite this close genetic relationship, morphologically algae of the genus Triparma are extremely heterogeneous. General description As described above, some algae (i.e. “Bolidomonas”) areunicellular biflagellates. Others (Parmales sensu Booth and Marchant) are small (2 to 5 µm) non-flagellate unicells covered with silicified plates of various shapes (Fig. 5.33; Silver et al. 1980; Booth and Marchant 1987). The motile algae possess two flagella of unequal length inserted

Fig. 5.33 Transmission electron micrographs and drawings of cells of the Bolidophyceae. a Pentalamina corona. b Tetraparma pelagica. c Triparma laevis. In a–c) D = dorsal plate, G = girdle plate, S = shield plate, and V = ventral plate. d T. eleuthera. The drawing is not to scale and represents the main feature of the cell. d = Circular DNA molecule, g = Golgi body, h = Tubular hairs of the immature flagellum, l = Girdle lamella, m = Mitochondrion, n = Nucleus, and p = Chloroplast. Cell orientation arbitrary, A = Anterior, D = Dorsal, P = Posterior, and V = Ventral. Scale bars = 1 µm. Modified from Kuwata et al. (2018) (redrawn from Booth and Marchant 1987) and from Guillou et al. (1999)

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ventrally (Fig. 5.33). The length of both flagella (long 4-7 µm, short flagella 0.9–2.2 µm) generally exceeds the size of the cell (1–1.7 µm). The longest flagellum extends forward in a wavelike motion, and its length might help explain the speed of the swimming (1–1.5 mm.s-1). The short flagellum is smooth and hairless, whereas the long flagellum bears tubular bipartite hairs. These bipartite hairs were also observed on the flagellum of Pelagomonas (Pelagophyceae; Fig. 5.29; Andersen et al. 1993) and Oikomonas (Chrysophyceae; Cavalier-Smith et al. 1995). The Bolidophyceae possess a single chloroplast with girdle lamella, no pyrenoid, and no eyespot. The photosynthetic pigments are chlorophylla, chlorophyllc1, chlorophyllc2 and chlorophyllc3, fucoxanthin, the diadinoxanthindiatoxanthin cycle, and ß-carotene (Table 5.2). The nonmotile algae are entirely covered by silicified plates fused together. Four different types of plates can be recognized based on their shape, and they are arranged around the cell in various numbers and combinations depending on the genus (Fig. 5.33a–c). The shield plates are circular as are the ventral plate, but this one is larger than the shield plates. The dorsal plates have three arms fitting around the shield plates. The girdle plates form a ring around the ventral plate and between the shield and dorsal plates. Occurrence Flagellated algae of the genus Triparma were originally isolated from the equatorial region of the Pacific Ocean and the Mediterranean Sea. In contrast, non-flagellated algae of the genus Triparma were principally isolated in polar and subpolar oceans. In recent years, environmental metabarcoding revealed a more widespread distribution of the Bolidophyceae around the world and a more complex diversity than previously thought (reviewed in Kuwata et al. 2018). Prolonged monitoring also revealed seasonal variation in the abundance and distribution in the water column of the Bolidophyceae (reviewed in Kuwata et al. 2018). Evolutionary history Molecular phylogenies have placed the Bolidophyceae as a sister group to the diatoms (Ichinomiya et al. 2011). This suggests that the characteristic silica frustule of the diatom may be derived from an ancestor covered with silicified plates similar to the actual Bolidophyceae; as was proposed before, the Parmales were known (Round and Crawford 1981, 1984). However, the fossil record of the Bolidophyceae is much younger than that of the diatoms. However, the recent re-examination of fossils from the Late Cretaceous (100–66 Mya) revealed that they belonged to the Bolidophyceae (Abe and Jordan 2021), extending the stratigraphic range of the Bolidophyceae by millions of years (Stradner

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and Allram 1982; Konno et al. 2007). Importantly, this discovery seems to reconcile the conflicting fossil records of the closely related Bolidophyceae and diatoms, supporting the hypothesis of a common ancestor to these lineages resembling the actual Bolidophyceae (Abe and Jordan 2021).

5.1.4.11 Pinguiophyceae In a quick succession of papers, the class Pinguiophyceae (Kawachi et al. 2002a) and four new to science algae were described (Kawachi et al. 2002b; O’Kelly 2002; Honda and Inouye 2002; Andersen et al. 2002). Remarkable for their large amounts of omega-3 fatty acids, they were named to represent this trait (Latin pingue – fat, grease). Algae in the Pinguiophyceae are marine, collected from the Atlantic and Pacific Oceans and the coast of Australia; their morphologies differ significantly (Fig. 5.34). Their photosynthetic pigments are typical of the Heterokontophyta with chlorophyll a and c, fucoxanthin, violaxanthin, zeaxanthin, ß-carotene, and traces of antheraxanthin. However, apart from the photosynthetic pigments, the genera of the class Pinguiophyceae are extremely diverse. These algae were either nonmotile unicells (Pinguiochrysis, Fig. 5.34a; Pinguiococcus, Fig. 5.34d), alternating mobile/nonmotile unicells (Phaeomonas, Fig. 5.34b–c), solitary or colonial cells (Glossomastix, Fig. 5.34e–g), or loricate unicells (Polypodochrysis, Fig. 5.34h–i). This diversity is also observed at the ultrastructural level. For example, Phaeomonas has two typical heterokont flagella, whereas Glossomastix and Polypodochrysis lack tripartite hairs. Pinguiochrysis and Pinguiococcus have no known flagellate stage. This high variety is found at almost every level and no specific unifying characters (i.e. synapomorphy) could circumscribe the Pinguiophyceae (Kawachi et al. 2002a). Despite the lack of clear unifying morphological and ultrastructural characters, the molecular and phylogenetic analyses support the establishment of this class (Kawachi et al. 2002a). The monophyly of the class Pinguiophyceae was consistently recovered in subsequent phylogenetic analyses (e.g., Yang et al. 2012). Furthermore, maybe the most peculiar and shared character of the Pinguiophyceae was their exceptionally high concentration of polyunsaturated fatty acid (PUFA) and especially of 20:5 (n-3) eicosapentaenoic acid (EPA). Fatty acid composition of the Pinguiophyceae was primarily of PUFA of which a large proportion is EPA (Phaeomonas, 69.9% PUFA and 56.0% EPA; Pinguiochrysis, 64.5% PUFA and 54.5% EPA; Glossomastix 58.0% PUFA and 39.2% EPA; Pinguiococcus 53.9% PUFA and 34.0% EPA; Polypodochrysis 47.4% PUFA and 23.5% EPA). These levels largely exceeded those of other Heterokontophyta, such as the diatoms (6–28.4% EPA; Brown et al. 1989; Zhukova and Aizdaicher 1995), Raphidophyceae (11.7–

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Fig. 5.34 Diversity of the Pinguiophyceae. a Pinguiochrysis pyriformis. b Phaeomonas parva swimming cell c Phaeomonas parva nonmotile cell. d Pinguiococcus pyrenoidosus. e Glossomastix chrysoplasta vegetative cell. f G. chrysoplasta colony. g G. chrysoplasta zoospore. h Polypodochrysis teissieri loricate vegetative cell. i Polypodochrysis teissieri zoospore. Scale bar = 10 µm (except f) Scale bar = 1 µm). Modified from Kawachi et al. (2002a, b)

16.3%; Nichols et al. 1987), Pelagophyceae (3.5–8%; Bricelj et al. 1989), or Phaeophyceae (8%; Wood 1988). Only Eustigmatophyceae (17.8–39.9%; Wood 1988; Renaud et al. 1991; Zhukova and Aizdaicher 1995). Only the Dictyochophyceae showed levels comparable to those of Pinguiophyceae (26%; Wood 1988) showed levels comparable to those of Pinguiophyceae. The absence of a cell wall and the relatively easy maintenance of the Pinguiophyceae species could have made them a valuable target for commercial extraction of PUFA and animal feeds. However, so far, no optimized growth for increased PUFA amount studies or large-scale extraction experimentations have been conducted on these algae.

5.1.4.12 Schizocladiophyceae The class Schizocladiophyceae was established with the discovery of a filamentous alga in the Mediterranean Sea (Kawai et al. 2003). Isolated from a collection of brown algae from the Island of Ischia, Italy, the morphology of these filaments resembles the filamentous gametophytes of the Phaeophyceae. Its pear-shaped zoospores possess two flagella of unequal length inserted laterally also resembling the motile cells of Phaeophyceae. They contain a single

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chloroplast with girdle lamella, a red eyespot placed at the base of the posterior flagellum, and no pyrenoid. The chloroplasts contain chlorophyll a and c, fucoxanthin, ßcarotene pigments, and potentially other xanthophylls not detected yet. After settlement, the zoospores form a cell wall and germinate into uniseriated filaments. Upon maturation, fertile cells differentiate into zoosporangia that become swollen, containing several chloroplasts and forming several zoospores in individual compartments. However, critical differences exist between Schizocladia and the Phaeophyceae that justify the distinction of the class Schizocladiophyceae: namely (1) the absence of cellulose in the cell wall of Schizocladia, whereas it is probably ubiquitous in Phaeophyceae (McCandless 1981; Painter 1983; Kawai et al. 2003); (2) the absence of plasmodesmata (or any intercellular cytoplasmic connections), whereas they are a distinguishing feature of the Phaeophyceae (Bisalputra 1966); (3) presence of a transitional helix in the flagella, whereas it is absent in Phaeophyceae (Fig. 5.35). The phylogenetic position of Schizocladia within the Heterokontophyta is consistent with the above description as they were always found in a sister position to the Phaeophyceae (Kawai et al. 2003; Kai et al. 2008; Yang et al. 2012; Graf et al. 2020a, b). This position and the differences between the Phaeophyceae and Schizocladia open an interesting perspective to understand the evolution of the multicellularity in the Heterokontophyta. The second-ever record of Schizocladia was recently reported from the Eastern basin of the Mediterranean Sea (Rizouli et al. 2020) indicating that this species and potentially other related taxa might be found in infralittoral and circalittoral marine communities and could have been overlooked due to their small size and maybe their resemblance to brown algae gametophytes.

5.1.4.13 Synchromophyceae The class Synchromophyceae was established from a genus of amoeboid marine algae: Synchroma (Horn et al. 2007). The class was named after the characteristic chloroplast complexes (Greek syn – with, together; Greek chroma – color) of these algae. First discovered on the Canary Islands, these algae were reported from the Atlantic Ocean and the Caribbean and Mediterranean Seas (Schmidt et al. 2012). They are amoeboid cells organized with a central spherical or fusiform core around which a network of simple and branched pseudopodia expends (Fig. 5.36a). These algae were characterized by a strikingly original organization of their chloroplasts. The chloroplasts aggregate in complexes enclosed in a common periplastidial membrane and epiplastid rough endoplasmic reticulum (Fig. 5.36b–c). The pyrenoids are positioned in the center of these complexes. This unique organization was never observed in any other organisms. However, other morphological characters (e.g.

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Fig. 5.35 Light microscopy and TEM of Schizocladia ischiensis. a Multiple branched filaments growing and showing swollen reproductive cells (arrows). b Single filament with swollen reproductive cells (arrows). c Cells of a filament. Note the absence of plasmodesmata in the cell wall. C = chloroplast; N = nucleus; CW = cell wall; arrow = original outer wall; arrowhead = new cell wall. From Kawai et al. (2003)

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chloroplast with girdle lamella) and pigment composition (i. e. chlorophyll a and c2, fucoxanthin, violaxanthin, antheraxanthin, zeaxanthin, and ß-carotene) linked them to the Heterokontophyta. Furthermore, molecular analysis clearly placed the Synchromophyceae within the photosynthetic Heterokontophyta (Horn et al. 2007; Schmidt et al. 2012). Recent phylogenetic studies including a broader range of plasmodium forming Heterokontophyta indicated their close relationship with the Synchromophyceae (Schmidt et al. 2015). However, these taxa (i.e. Chlamydomyxa, Chrysopodocystis, and Guanchochroma) do not form chloroplast complexes, and their affiliation to the Synchromophyceae would necessitate a revision of the class definition.

5.1.4.14 Aurearenophyceae The class Aurearenophyceae was established after the discovery of a unicellular alga, Aurearena, on the sandy beaches of Japan (Kai et al. 2008). The name of the class came from the ecological niche and color (Latin Aurearena golden sand) of this alga. The genus Aurearena is characterized by a cell cycle alternating between a naked, free-living, swimming stage (Fig. 5.37b) and a nonmotile stage where the cell is covered by a cell wall (Fig. 5.37a). Interestingly, during its entire cycle, the cells possess two flagella of unequal length, an

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immature one bearing tripartite tubular hairs, and a smooth mature flagellum. The cells possess a single chloroplast divided into two compartments each surrounded by three membranes and joined together by a fourth membrane. A yellowish eyespot is present in the chloroplast, and it is associated with the short flagellum. The pyrenoid of the chloroplast is placed in the center of this arrangement. The two compartments of the chloroplast each present a deep invagination, and this original architecture marks a distinguishable cross pattern in the pyrenoid in the center of the cell (Fig. 5.37b). The photosynthetic pigments of Aurearena are composed of chlorophyll a, fucoxanthin, diadinoxanthindiatoxanthin cycle, violaxanthin, antheraxanthin, zeaxanthin, and ß-carotene, and strikingly the chlorophyll c has not been detected in this alga. Some of these characteristics support the Heterokontophyta affiliation of Aurearena (e.g. two flagella of unequal length and tripartite tubular hairs on the longest one) but at the same time are making it a unique member of these algae (e.g. a single chloroplast separated in two compartments and without a girdle lamella; absence of chlorophyll c), justifying its treatment as a distinct class. Molecular analysis further confirmed both the position of the Aurearenophyceae within the Heterokontophyta and as a distinct lineage probably affiliated with the class Phaeothamniophyceae (Kai et al. 2008). A position that has been consistently recovered in recent phylogenetic analysis

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not been observed. Rather they develop into a single zoospore that escapes the cell wall leaving it empty. The zoospore possesses two typical Heterokontophyta flagella inserted near the center of the cell. The immature flagellum projects forward and the mature flagellum trails behind. The immature flagellum is 2–3 times the length of the cell and covered with tripartite hairs, whereas the mature flagellum is short and smooth. The eyespot is conserved in one of the chloroplasts after the differentiation. Ultrastructural observations of the cell revealed that the chloroplast is surrounded by only two membranes, not four membranes as with other Heterokontophyta (Fig. 5.38). The nature of these membranes was hard to determine but it appears that they are homologous with the outermost and innermost membranes of the typical Heterokontophyta chloroplast (Fig. 5.38b–d) and that the two other membranes were presumably lost (Wetherbee et al. 2019). The chloroplast stroma is typical of the Heterokontophyta with three-stacked thylakoids and a girdle lamella. The photosynthetic pigments are principally the chlorophyll a, c1, c2, and fucoxanthin, and other accessory pigments found in smaller concentrations were violaxanthin, antheraxanthin, zeaxanthin, and ß-carotene. The discovery of this alga and the unusual number of membranes of its chloroplast are puzzling. Its position within the Heterokontophyta was only confirmed by phylogenetic analysis, and it’s exact branching is still unclear. Despite the major event that was the loss of two membranes, proteins targeted to the chloroplast do not appear to have lost their targeting signal and transit peptide (Wetherbee et al. 2019). This suggests a relatively recent loss and a chloroplast targeting and import system typical to the Heterokontophyta. Investigation of more algae related to Chrysoparadoxa will certainly help understand the evolutionary history of this class. The sand-dwelling habitat of Chrysoparadoxa has not been extensively explored and more taxa might be discovered. Interestingly, algae of the genus Nematochrysis were recently shown to be genetically close to Chrysoparadoxa (Graf et al. 2020b). However, there is no information on the number of their chloroplast membranes.

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Fig. 5.36 Characteristics traits of the cells of Synchroma grande. a Light microscopy capture of S. grande. b Schematic drawing of the chloroplasts complex of S. grande. Several chloroplasts permanently share the outer pair of enveloping membranes that closely follow the outline of the chloroplast complex. The entire chloroplast aggregate constitutes a unity which represents a single, multiplastidic secondary endosymbiont. c Pigmented chloroplast lobes ending in electron-lucent regions in which lamellae are missing, and are attached to the pyrenoids by flattened stipes. Every chloroplast is enveloped by a pair of closely attached inner membranes (white arrowheads) and by an additional outer membrane pair (black arrowheads). The outermost membrane, called epiplastid rough endoplasmic reticulum, is studded with cytoplasmic ribosomes (arrows). The subjacent periplastid membrane borders the periplastidial compartment in which membrane-lined periplastid reticula occur (asterisks). In the region of the pyrenoids, the periplastidial compartment is particularly narrow. C = Chloroplast, P = Pyrenoid, R = Region missing lamellae, and Cv = Capping vesicle (modified from Horn et al. 2007)

(Graf et al. 2020a, b) and some authors have suggested the inclusion of the class Aurearenophyceae as an order of the class Phaeothamniophyceae based on this phylogenetic position (Ruggiero et al. 2015).

5.1.4.15 Chrysoparadoxophyceae The class Chrysoparadoxophyceae was established very recently when an enigmatic sand-dwelling alga was discovered in Australia (Wetherbee et al. 2019). Unusual features and a distinct position in the phylogeny of the Heterokontophyta gave this alga its name (Greek chrysós gold; Greek parádoxos - contrary to expectation).. The alga of the monospecific genus Chrysoparadoxa is unicellular and found attached to sand grains. The cells are tubular, rounded at their ends, 3
7 µm in size, and covered by a cell wall. The cells contain two highly lobbed chloroplasts that cover a large proportion of the cell and stain the cells a yellowish slightly green coloration. One chloroplast contains an eyespot. Division of the benthic cells has

5.1.4.16 Phaeosacciophyceae The class Phaeosacciophyceae was described recently to accommodate a number of taxa (i.e. “Giraudyopsis”, Nematochrysis, Phaeosaccion, and Tetrasporopsis) whose classification has been controversial for years (Graf et al. 2020b). For example, Tetrasporopsis was first described as a green algae, Tetraspora fuscescens (Braun in Kützing 1849), then transferred to Phaeocystis (Haptophyta) in the new generic section Tetrasporopsis (DeToni 1895), and subsequently raised to generic level and classified in the Phaeophyceae (Lemmermann 1899). Similarly, Phaeosaccion

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Fig. 5.37 Drawings of the cells of Aurearena cruciata. a A single flagellated cell escaping from the cell wall. b Cell cycle of A. cruciata. 1—A flagellate cell inside a cell wall. 2—Naked swimming cell emerging through a pore in the cell wall. 3—Swimming cell. 4—Cytokinesis proceeding in a swimming cell. 5—Swimming cell absorbing flagella into the cell. 6—Round nonmotile cell after absorption of flagella. c Walled cell viewed from the lateral side. d Flagellated cell viewed from the lateral side. C = chloroplast, E = Eyespot, G = Golgi body, M = Mitochondrion, N = Nucleus, P = Pyrenoid, W = Cell wall, EV = Electron opaque vesicle, BB = Basal body, LF = Immature flagellum, and SF = mature flagellum. From Kai et al. (2008)

collinsii (Farlow 1882) was first described as a simple brown alga, but then was successively classified in the Chrysophyceae (Chen et al. 1974), Phaeothamniophyceae (Cryan et al. 2015), and Chrysomerophyceae (Gabrielson and Lindstrom 2018). The situation was further complicated by the descriptions of the class Chrysomerophyceae to accommodate these diverse taxa (Cavalier-Smith et al. 1995). This class was named after the genus Chrysomeris (the correct Latin spelling should have been Chrysomeridophyceae), established by Carter (1937), who described two species from a brackish pond in the Isle of Wight, UK. These filamentous algae were epiphytes on Spartina plants and were remarkable for their zoosporangium where numerous naked zoospores were formed in a gelatinous sheath, and then released via breaks or dissolution of the sheath. The zoospores had a single flagellum and no eyespot with two or three band-shaped chloroplasts (Carter 1937). Despite notable differences (e.g. zoospore with two flagella and an eyespot;

one zoospore formed in a vegetative cell and escaped from a lateral pore in the cell wall), Gayral and Haas identified another isolated alga as Chrysomeris (Gayral and Hass 1969). This description got large acceptance and somehow replaced the original description (e.g. Kristiansen and Preisig 2001), and the alga identified by Gayral and Hass (1969) was used to anchor the family Chrysomeridaceae within the order Sarcinochrysidales (Gayral and Billard 1977a) and later the class Chrysomerophyceae (Cavalier-Smith et al. 1995). However, all these successive classifications were conducted without analyzing biological material from the type locality and without even referring to the original description of the genus Chrysomeris. Therefore, the class Chrysomerophyceae is no longer valid. The recent re-investigation of the genera previously included in this class (i.e. Antarctosaccion, Chrysomeris, Chrysowaernella, “Giraudyopsis”, Nematochrysopsis, and Phaeosaccion) led to the description of the new class Phaeosacciophyceae (Graf et al. 2020b).

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Fig. 5.38 Light microscopy and TEM of Chrysoparadoxa australica. a Benthic cells and abandoned walls (arrows) following zoospore release. Two U-shaped chloroplasts are placed at the poles of the cells (u) and then split into two lobes (arrowheads). b–d Chloroplast membranes and chloroplast. b The girdle lamella (Gl) encloses three stacked thylakoids, and the chloroplast is surrounded by two membranes: the inner chloroplast membrane (white arrowheads) and outer chloroplast membrane (black arrowheads) that are adjacent to the plasma membrane (arrows). c High magnification of a region as seen in (b). The train track structure of the membranes is observed and clearly shows the singular structure of the chloroplast membrane in C. australica with an outer chloroplast membrane (oc) and an inner chloroplast membrane (ic). d The inner (white arrowheads) and outer (black arrowheads) chloroplast membranes are observed at the union of two lobes of a chloroplast (C); pm = plasma membrane; * = cell wall. From Wetherbee et al. (2019)

This complex history and affiliation to many different classes over time can be explained by their diversity of shape, form, and organization, with algae forming filaments (Antarctosaccion, Phaeosaccion, Fig. 5.39a), colonies (Tetrasporopsis, Fig. 5.39e–f), or strictly unicellular organisms (Psammochrysis, Fig. 5.39j–l). That is, there was a lack of unifying morphological characters. These algae are also found in a wide variety of environments from freshwater streams in the Northern hemisphere (Tetrasporopsis) to Antarctic infralittoral epiphytes on red algae (Antarctosaccion) to sand dwelling unicells in Australia (Psammochrysis). Therefore, their unity was only revealed recently through molecular analysis that showed that despite the lack of structural features shared across the entire class genetically, they are closely related forming a clear monophyletic group in phylogenetic trees (Graf et al. 2020b).

5.1.4.17 Olisthodiscophyceae The class Olisthodiscophyceae was recently established following the re-investigation of the genus Olisthodiscus with molecular methods (Barcytè et al. 2021). The genus Olisthodiscus was described in 1937 by Nellie Carter based on an alga isolated from the brackish

Bembridge lagoon on the Isle of Wight, United Kingdom. The classification of Olisthodiscus has remained controversial, being placed alternatively in the Xanthophyceae (Carter 1937), the Raphidophyceae (Hara et al. 1985; Inouye et al. 1992), or the Chrysophyceae (Vesk and Moestrup 1987). The cells of Olisthodiscus are round to ovate with a clear polarization of the cell shape by a concave flattened side and a convex side (Fig. 5.40). They possess the two unequal flagella typical of the Heterokontophyta (i.e. immature and mature flagellum) inserted in a ventral depression near the nucleus. Numerous plastids are present in the vegetative cells, and the plastids do not have eyespots. In certain cells, an extraplastidial globule can be observed reminiscent of the one observed in Eustigmatophyceae (Fig. 5.24). Interestingly, molecular phylogenies showed that Olisthodiscus was clearly separate from the Xanthophyceae, Chrysophyceae, or Raphidophyceae and represented an independent lineage within the Heterokontophyta, being sister to the Pinguiophyceae. The molecular phylogenetic analysis, as well as the unique morphological and ultrastructural characteristics of the genus Olisthodiscus, supported the establishment of the new class Olisthodiscophyceae (Barcytè et al. 2021).

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Fig. 5.39 Diversity of the Phaeosacciophyceae. a Thallus of Phaeosaccion collinsii showing gross morphology, with putative holdfast in the lower right corner. b Dividing cells of P. collinsii showing the single parietal chloroplast and the highly organized pattern. c Palmelloid stage of Nematochrysis sessilis var. vectensis with irregularly organized cells. Note also the aplanospore sporangium (arrow). d Early stage of aplanospore formation in N. sessilis var. vectensis with two cells per future sporangium (arrows). e Globular colonies of Tetrasporopsis fuscescens in situ (Eel River, Ca, USA). f Spherical cells of T. fuscescens with 2 or 3 parietal chloroplasts and situated in the outer colonial mucilage. Remnant cell walls (arrows) scattered among vegetative cells. g Thallus of Phaeosaccion multiseriatum formed from basal cell mass and consisting of numerous uniseriate and multiseriate filaments. h Thallus of P. multiseriatum multiseriate branched filaments arising from a central area. i Thallus of P. multiseriatum parenchymatous-like cells with no obvious filamentous origin. j Raft structure of Psammochrysis cassiotisii with benthic cells on the rim, remnant cell walls in the interior. k Small raft of P. cassiotisii, benthic cells, and empty cell walls following zoospore escape are in pairs. l Benthic cells and remnant cell walls of P. cassiotisii at the edge of a raft. a–d and g–l from Graf et al. (2020b); e–f from Stancheva et al. (2019)

276 Fig. 5.40 Light microscopy and TEM of Olisthodiscus luteus strain K-0444. a Globose vegetative cells showing the central nucleus and a large orange extraplastidial globule. b Vegetative cells with pyrenoids in the chloroplasts. c Broadly ellipsoidal vegetative cell showing the two unequal flagella emerging sub-anteriorly. d Lateral view of cell showing the convex dorsal and slightly concave ventral side with the funnel-shaped depression where the typical Heterokontophyta flagella emerge. From Barcytè et al. (2021)

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5.1.5 Perspectives 5.1.5.1 Heterokontophyta in the Genomics Era The long and rich history of knowledge on the Heterokontophyta has been largely shaped by the available technology and scientific methods. The first genome of a Heterokontophyte was fully completed in 2004 (Armbrust et al. 2004). This prompted the start of a new era of the scientific history of this group: the genomic era. Since then, 64 genome sequences have been released (NCBI consulted September 2022). These genome sequences have been accompanied by a large collection of transcriptomes and metagenomes, all together allowing pushing back the boundaries of our understanding of the Heterokontophyta. Much has been learned about the metabolism, physiology, ecology, genetics, and evolutionary history of the Heterokontophyta, notably thanks to the Next Generation Sequencing (NGS) revolution. The genome sequences of the diatoms helped to understand their biology, notably with the discovery of genes

involved in silica, iron, nitrogen metabolisms, or their complete urea cycle, all essential components for the survival of Diatomeae (Armbrust et al. 2004; Bowler et al. 2008; Allen et al. 2011). Genome-wide comparison of cold water and temperate water diatoms revealed adaptations to the Arctic Ocean (Mock et al. 2017). Similarly, the completion of the first brown algal genome (i.e. Ectocarpus siliculosus; Cock et al. 2010) brought insights into the independent establishment of multicellularity in this lineage as well as insights into brown algae metabolism (Cock et al. 2012). Furthermore, genome sequences of the kelps Saccharina japonica (Ye et al. 2015) and Undaria pinnatifida (Graf et al. 2021) have started to explore the emergence of the complex morphologies of kelps, the genomic signatures of their characteristic traits, and notably their original iodine metabolism. The genes involved in this particular metabolism form a gene family of vanadium-dependent haloperoxidase that has been considerably expended and specialized in the kelps (Ye et al. 2015). In the Eustigmatophyceae, the

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genomes of the genera Nannochloropsis and Microchloropsis have been largely mined principally with the objectives of biofuel production and synthetic biology (e.g. Radakovits et al. 2012; Gong et al. 2020). The availability of multiple high-quality genomes also allowed for the exploration of the evolution in this group, notably revealing that their oleaginous production was in part inherited through horizontal gene transfer (Wang et al. 2014). Genomic analyses have not been restricted to nuclear genomes because organelle genomes also have been important for understanding the evolution of certain lineages (e.g. diatoms Yu et al. 2018; Eustigmatophyceae Ševčíková et al. 2019; Chrysophyceae Kim et al. 2019; Laminariales Starko et al. 2019; Starko et al. 2021). Their relatively easy access also makes them a resource for other types of analysis, such as phylogeny (e.g. Graf et al. 2017) or biogeography (e.g. Starko et al. 2019).

5.1.5.2 Evolutionary Trends The rapid accumulation of transcriptome and genome data for species of the Heterokontophyta has allowed the exploration of difficult evolutionary questions. Relationships between classes of Heterokontophyta It is now widely accepted that the Heterokontophyta (i.e. photosynthetic lineages of the Stramenopiles) form a monophyletic group, regrouping a wide diversity of organisms classified in the 17 classes discussed thus far (e.g. Moriya et al. 2002; Derelle et al. 2016). However, one of the most preeminent questions has remained the phylogenetic relationships between the classes of the Heterokontophyta. The hope raised with molecular phylogenetic analysis was not totally fulfilled. A consensus started to emerge in the past decade with the separation of three clades named SI, SII, and SIII that have been recovered by multiple independent studies (Fig. 5.2; Yang et al. 2012; Ševcíková et al. 2015; Derelle et al. 2016; Ševcíková et al. 2016). The SI clade (Fig. 5.2) regroups the Aurearenophyceae, the Chrysoparadoxophyceae, the Phaeophyceae, the Phaeothamniophyceae, the Phaeosacciophyceae, the Raphidophyceae, the Schizocladiophyceae, and the Xanthophyceae; the SII clade (Fig. 5.2), the Chrysophyceae, the Eustigmatophyceae, the Olisthodiscophyceae, the Pinguiophyceae, and the Synchromophyceae. And finally, the SIII clade (Fig. 5.2) regroups the Diatomeae, the Bolidophyceae, the Dictyochophyceae, and the Pelagophyceae. However, despite the use of multiple supermatrix datasets (i.e. a large combination of aligned gene sequences obtained from multiple species), the ancient relationships within the Heterokontophyta remain generally unresolved. Furthermore, most of these supermatrix studies were missing lineages and therefore could not strongly confirm or disprove

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the three clades and their branching. Recently, in an effort to clarify this, new mathematical approaches were applied to multiple datasets of genes from all major lineages (Di Franco et al. 2021). Interestingly, these analyses revealed that the different genomes (i.e. nuclear, mitochondrial, and chloroplastic) had conflicting phylogenetic signals (Fig. 5.41), adding to the complexity of this problem. Further analysis will be required before confidently resolving the phylogenetic tree of the Heterokontophyta; modern transcriptomebased trees including the entire diversity of the Heterokontophyta lineages will surely be necessary. Endosymbiosis and the evolution of photosynthesis The large amount of genomic data that have been accumulated in the last decade has also allowed exploring endosymbiosis and its evolutionary consequences. Based on the presence of chlorophyll c in their photosynthetic members, the Alveolates, Cryptophytes, Haptophytes, and Stramenopiles have been placed in a monophyletic group named Chromalveolates (Cavalier-Smith 1999). This group was hypothesized to have gained their chloroplast through a secondary endosymbiosis event with a red alga (Cavalier-Smith 1999). However, the monophyly of this group was rapidly questioned and with the development of more sophisticated molecular analyses, it emerged that these four lineages were not closely related (Sanchez-Puerta and Delwiche 2008). Therefore, deciphering the evolutionary scenario that led to the presence of chlorophyll c containing chloroplasts in those lineages remains an open question. Molecular phylogenetic analysis of chloroplast-encoded genes generally places the Heterokontophyta as a sister to the red algae (Le Corguillé et al. 2009; Dorrell et al. 2017). Therefore, the chloroplast of the Heterokontophyta has been largely viewed to have originated from a single secondary endosymbiosis with a red algal endosymbiont that took place in the common ancestor of the photosynthetic lineages (see Chap. 2, Endosymbiosis). However, using chloroplast genes represents the evolutionary relationship of the chloroplast and not of the eukaryote lineages themselves (Palmer 2003). Therefore, nuclear genomic analyses have brought a renewed view of the evolution of the chloroplast in the Heterokontophyta and other chlorophyll c containing lineages. Notably, genomic comparison and statistical analysis revealed that successive endosymbiosis events occurred (Bodył et al. 2009; Stiller et al. 2009; Strassert et al. 2021). This suggests that the chloroplast of the Heterokontophyta was the result of a tertiary endosymbiosis with a Cryptophyte endosymbiont (who acquired its chloroplast through a secondary endosymbiosis with a red alga). This serial endosymbiosis hypothesis was further complicated when genomics analysis revealed that an important proportion of the genes found in the genomes of the Heterokontophyta was

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Fig. 5.41 Incongruencies in the phylogenetic trees of the Heterokontophyta. a Mitochondrial dataset—outgroup of 15 species not shown. b Plastid dataset—outgroup of 15 species not shown. c Nuclear dataset—outgroup of 27 species not shown. SSC, monophyly of Synchromophyceae, Synurophyceae, and Chrysophyceae; Synurophyceae is not seen as a lineage separated from Chrysophyceae anymore (see Fig. 5.2); Bacillariophyta corresponds to Diatomeae; reproduced from Di Franco et al. (2021)

acquired through horizontal gene transfer not only from red algal origin (i.e. acquired during the endosymbiosis) but also from green algal origin (Moustafa et al. 2009; Dorrell et al. 2017; Morozov and Galachyants 2019; Sibbald and Archibald 2020). These genes were associated with the chloroplast,

and several of them targeted it, suggesting that they could be the remaining trace of a “cryptic” endosymbiosis event that occurred in the early evolutionary history of the Heterokontophyta (Moustafa et al. 2009; Dorrell and Smith 2011; Dorrell et al. 2017, Dorrel and Bowler 2017; Morozov and

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Galachyants 2019; Sibbald and Archibald 2020). The clear chain of events leading to the establishment of the chloroplast and the genomic mosaic of Heterokontophyta remains the subject of current debate and ongoing research. Secondary loss of photosynthesis has occurred in three groups (i.e., Diatomeae; Chrysophyceae; Dictyochophyceae), with the reduction of the chloroplast to a colorless organelle. Recently, the roles of these non-photosynthetic chloroplasts have been explored by genome sequencing (Kamikawa et al. 2017, 2018; Kim et al. 2020; Kayama et al. 2020; Dorrell et al. 2021). It revealed a rapid loss of genes involved in photosynthesis and suggested that these peculiar chloroplasts are only maintained to support other biosynthesis pathways (e.g. amino acid biosynthesis). These analyses also revealed convergence(s) during the evolution of non-photosynthetic chloroplasts toward similar gene retention to support similar metabolic pathways (Dorrell et al. 2021).

5.1.5.3 Functional Genomics of the Heterokontophyta As discussed above, the genomic era brought fascinating new insights into the evolution of the Heterokontophyta, but the functional aspect of their genomes remains largely unexplored. By nature, genome data is finally just a collection of gene sequences, the vast majority having no known function. Furthermore, the revolution of epigenetics brought a new layer to functional genomics that has only begun to be investigated in the Heterokontophyta. Exploration of the functional aspects of the Heterokontophyta genomes notably requires reverse genetics and quantitative genetics approaches. To date, reverse genetics in Heterokontophyta is limited to the handful of species that have been successfully transformed. Furthermore, these species only belong to the Diatomeae (Dunahay et al. 1995; Huang and Daboussi 2017), Eustigmatophyceae (Radakovits et al. 2012), and recently to the Phaeophyceae (Badis et al. 2021). Therefore, functional genomic approaches have remained restricted to those groups. In the Diatomeae, the past 20 years have seen important progress in understanding the biology of diatoms, notably with the advances of functional genomics (reviewed in Falciatore and Mock 2022). In the brown alga Ectocarpus siliculosus, mutant strains were established that helped investigate some developmental pathways (e.g. Billoud et al. 2015; Macaisne et al. 2017). Quantitative genetics approaches, such as Quantitative Traits Locus (QTL) detection, have been limited to species of the Phaeophyceae (Avia et al. 2017; Wang et al. 2018). The epigenome of the Heterokontophyta is extremely poorly characterized. To this day, only the epigenetic landscape of the diatom Phaeodactylum tricornutum has been comprehensively characterized (Veluchamy et al. 2013, 2015). This helped to explore its role in response to the

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environment and in the Diatomeae biology (reviewed in Zhao et al. 2022). In the brown algae, the epigenome is even less understood. It appears that DNA methylation might not be as fundamental as in other lineages as around 1.5% of the genome of the kelp Saccharina japonica is methylated (Fan et al. 2020; Yang et al. 2021) and was not detected in Ectocarpus siliculosus. However, because the methylome of kelps growing in different environments was different, it appears that DNA methylation despite its low frequency could play a role in local adaptation (Scheschonk et al. 2021). In Ectocarpus siliculosus histones modifications and miRNA have been characterized, suggesting that post-transcriptional regulations are at play in the brown algae (Cock et al. 2017; Bourdareau et al. 2021). The characterization of the epigenome of more Heterokontophyta will be an important step in our understanding of these algae.

5.1.5.4 Genomics of Heterokontophyta in the Global Change Era During their millions years long evolutionary history, the Heterokontophyta have experienced, adapted, and survived drastic local and global environmental changes (e.g. the end of the Last Glacial Maximum; see Graham et al. 2003, 2010; Fraser et al. 2009). However, as we are witnessing adaptations in response to contemporary human-induced changes, it is important to understand how the Heterokontophyta will cope with and adapt to the ongoing global changes, and it is unsurprisingly an important driver of contemporary research. Ocean warming, acidification, and deoxygenation are expected to have profound impacts on the communities of marine organisms, including the Heterokontophyta (Sunday et al. 2017; Gao et al. 2019). These impacts will not be homogeneous in all the oceans and ecosystems as will be the response of the diverse organisms of the Heterokontophyta. Because of their important ecological and economic roles, the Phaeophyceae, and kelps in particular, have been particularly investigated. The increase of pCO2 and resulting acidification can apparently be beneficial as it stimulates primary production in some populations of Phaeophyceae (Celis-Plá et al. 2017; Cornwall et al. 2017), and this can increase their stocks (Linares et al. 2015; Sunday et al. 2017). Analysis of the transcriptome of Sargassum vulgare growing near volcanic vents (i.e. increased CO2 concentration and lowered pH conditions) further support that idea, with the differential expression of genes involved in energy metabolism, photosynthesis, and ion homeostasis (Kumar et al. 2017). However, different responses can be expected from different species, as some have shown no response to similar conditions (e.g. Saccharina latissima and Laminaria digitata [Roleda and Hurd 2012]) or more complex response (e.g. Fucus serratus [Johnston and Raven 1990]). Furthermore, as many of the brown alga species are acclimated to cold water, warming and heat waves are a

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more concerning threat to underwater forests (Wernberg et al. 2019). The resilience of the kelp forests to heat waves was investigated in Australia and revealed their replacement by turf-forming species (Filbee-Dexter and Wernberg 2018) and drastic genetic diversity reduction (Coleman et al. 2020). Adaptation, in the form of differential gene expression analysis, of kelps species subjected to increased temperatures revealed not only complex physiological responses but also their plasticity in the face of this stress (Heinrich et al. 2012; Liu et al. 2014; Heinrich et al. 2015; Machado Monteiro et al. 2019; Zhang et al. 2021). Global warming will also alter locally other factors, such as salinity, that have also started to be investigated with transcriptome analysis (Machado Monteiro et al. 2019; Rugiu et al. 2020; Zhang et al. 2021). Finally, inter-population differences in transcription responses to stresses suggest that tolerance to global changes may differ possibly due to local adaptation (Machado Monteiro et al. 2019; Rugiu et al. 2020). This level is also at the core of population genomics, which aims at studying individuals within a population at a genome-wide scale to understand the evolution of populations. Such genomics approaches have started to be applied in the conservation and restoration efforts of kelp forests (e.g. Wood et al. 2020). Along with the increasing availability of kelp genomes, transcriptomes studies and other omics approaches (i.e. miRNA Lui et al. 2015 or proteome Lui et al. 2019) as well as ongoing large-scale population genomics projects (e.g. Graf et al. 2021) will contribute to identify important loci and populations of interest for conservation and management efforts in the future. On the other hand, the impact of warming and acidification on unicellular Heterokontophyta is less well documented. Most approaches focus on their ability to fix carbon (e.g. Singh and Ahluwalia 2013) and less on the impact of these global changes on their evolution and survival. In the diatoms, the effect of increased pCO2 on growth, photosynthesis, silica fixation, and metabolism in general appears to be dependent on the size of the cell (Finkel et al. 2010; Key et al. 2010; Richier et al. 2018; Sarthou et al. 2005; Wu et al. 2014). It also appears that a gradual pCO2 increase has a more negative effect (Li et al. 2019), and analysis from natural populations showed more drastic negative effects than experiments in controlled environments (Petrou et al. 2019). Transcriptomic analysis under increased temperature and pCO2 of a diatom started to uncover the genetic side of the observed adaptations (Thangaraj and Sun 2021). Most, if not all, of the studies on global changes have focused on marine species, and the freshwater ecosystems have yet to be investigated as the Heterokontophyta (e.g. Chrysophyceae) play an important role in them too. Overall, the response and adaptation of the Heterokontophyta will reshape the communities of both macro and microalgae and

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5.1.5.5 Genomics, Taxonomy, and “Tradition” The promises held by new technologies and their power to study the Heterokontophyta should not totally shadow the more traditional approaches. There has been no slowing down in the discovery of unknown diversity in the Heterokontophyta. It could even be argued that new technologies helped to uncover unexpected cryptic diversity (e.g. Moon-van der Staay et al. 2001; Díez et al. 2001; Seeleuthner et al. 2018; Fawley et al. 2021). However, the tedious work of re-investigating the classic literature and visiting type localities is still necessary for our understanding of the evolution of the Heterokontophyta. This has been particularly important in recent years with a number of publications helping to clarify, correct, and update the taxonomical classification of the Heterokontophyta (e.g. Andersen et al. 2017; Han et al. 2019; Pusztai and Škaloud 2019; Graf et al. 2020a, b; Wetherbee et al. 2020; Pusztai and Škaloud 2021). In parallel, the continued exploration of nature is also essential, and the discovery and description of new algae greatly participated in expending our understanding of the evolutionary history of the Heterokontophyta (e.g. Kai et al. 2008), sometimes adding complexity and more questions for the future, as with the enigmatic Chrysoparadoxa (Wetherbee et al. 2019 and discussion in Graf et al. 2020b). Despite recent progress, many algae still await to be discovered in nature and others to be re-investigated with modern methods. Therefore, the scientific study of the Heterokontophyta will continue to be a blend of new ambitious and traditional approaches.

5.2

Dinoflagellates

Thomas Friedl and Mona Hoppenrath Dinoflagellates are a peculiar group of unicellular biflagellated organisms. Only a few dinoflagellates may form colonies or nonmotile stages. Dinoflagellates are probably the most abundant and diverse eukaryotes in marine surface waters. Only about 10% of the known species live in freshwater environments, from tropical to polar waters (Moestrup and Calado 2018). Due to their abundance, the photosynthetic dinoflagellates represent a fundamental trophic basis for most other marine life and are second only to diatoms as primary producers on Earth (Field et al. 1998; Fukuda and Suzaki 2015). Thus, the dinoflagellates are important primary producers, symbionts, but at the same time, also consumers and parasites. They are also common inhabitants of marine benthic sediment habitats (Hoppenrath

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et al. 2014; Saldarriaga and Taylor 2017). Dinoflagellates structure and sustain benthic reef ecosystems (e.g. the symbiotic genus Symbiodinium; Not et al. 2016). Dinoflagellates are involved as photosynthetic partners in a broad variety of symbioses (or mutualistic photosymbiosis) with heterotroph hosts such as radiolarians and metazoa (Not et al. 2016). Dinoflagellates cause marine bioluminescence (e.g. Noctiluca, Pyrocystis), and most toxic red tides (e.g. Karenia brevis, Fig. 5.44e; Lingulodinium polyedra, Fig. 5.45a, b). They indirectly cause some human diseases like paralytic shellfish poisoning and ciguatera (Asakawa et al. 2015; Saldarriaga and Taylor 2017). Roughly, half of the dinoflagellates are non-photosynthetic, and loss of photosynthesis has occurred repeatedly. Exceptional among all algae, dinoflagellates exhibit several evolutionary transitions from the photosynthetic to the heterotrophic lifestyle. Consequently, only about half of the described species are photosynthetic containing plastids originating from a red algal ancestor through secondary endosymbiosis. Many non-photosynthetic species still contain cryptic, unpigmented plastids, or bear genomic evidence for plastid-derived metabolic pathways (Saldarriaga and Taylor 2017). Mixotrophy is widespread among photosynthetic dinoflagellates, that is, they rely on a combination of photosynthesis and heterotrophic nutrition. The relative importance of the uptake of organic nutrients and photosynthesis for the dinoflagellates’ nutrition is still unknown. The heterotroph non-photosynthetic dinoflagellates contribute as consumers to the microbial loop. Parasitic dinoflagellates play a central role in the collapse of harmful algal blooms.

5.2.1 Organization and Structural Features of Dinoflagellates Cells 5.2.1.1 General Morphology Dinoflagellates range in size from less than 5 µm to more than 1 mm; colonies may even be several millimeters long. Dinoflagellates exhibit a characteristic morphology (Hoppenrath 2017): a traverse furrow (cingulum) dividing the cell like a girdle into an upper part (episome), terminating in the apex (nearly always with characteristic apical ultrastructures), and a lower part (hyposome; Fig. 5.42a). Both parts may be unequal in size (Figs. 5.45g, 5.46b). A longitudinal furrow (sulcus) extends from the cingulum to the posterior end of the cell. In species without cell wall, the sulcus often is with a thin extension that may even reach the cell’s apex. Both ends of the transverse furrow do not always meet at an equal level of the sulcus but can be displaced against each other (Figs. 5.43a, 5.45f). The longitudinal furrow determines the cell orientation, i.e. the side with the sulcus is defined

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as the ventral side thus, the opposite side is called dorsal, and left and right lateral sides can be distinguished. In the genus Prorocentrum, furrows are missing and the two flagella are inserted apically (Fig. 5.46c). Cell wall Dinoflagellate cells may be naked or possess a wall (Fig. 5.42b). A single layer of flattened vesicles (amphiesmal vesicles or alveoli) lies beneath the cell membrane (plasmalemma; Fig. 5.42b). Therefore, the outer cell layer of the dinoflagellate cells is structurally complex and quite distinctive; it is called amphiesma. The layer of alveoli beneath the cell membrane represents a structure homologous in its nature that is present across all lineages of the Alveolata, i.e. it is also present in the ciliates (Ciliophora) and Apicomplexa. In cells that are called naked, athecate, or unarmored, the amphiesmal vesicles are “empty”, and the vesicles themselves play a structural role (Fig. 5.42b). In the walled cells, cellulosic material is deposited inside the vesicles forming a plate with various pores. There is only one plate per vesicle (Fig. 5.42b). Because the plates usually fit tightly together with their margins often overlapping, a robust theca is formed which determines the shape of the cell. Cells with theca are called thecate or armored, and the single elements of the theca are called thecal plates. Notably, the wall of dinoflagellates is intracellular, i.e. it is located beneath the cell membrane. Consequently, growth of a walled dinoflagellate cell is somehow limited and only possible by the addition of wall material along the margins of the thecal plates. Arrangement of the thecal plates gives a certain pattern (tabulation) which is of critical importance in traditional dinoflagellate taxonomy for the distinction of species (Fig. 5.45c, e; e.g. Hoppenrath et al. 2009; Hoppenrath 2017). In prorocentroid dinoflagellates without furrows, e.g. Prorocentrum, the theca is composed of two large plates, i.e. right and left lateral plates, and there are additional small platelets in the apical periflagellar area where both flagella are inserted (Fig. 5.46c). Flagella Most species live as single unicellular flagellates where the motile phase is dominant. But some spend most of their life cycle in a nonmotile (coccoid) form without flagella, with biflagellated motile cells (i.e. zoospores, of the morphology typical for flagellated dinoflagellates) formed during reproduction. A motile cell possesses two flagella characteristics for dinoflagellates that are differentiated morphologically and functionally (dinokont flagellation). The unique transverse flagellum is located inside the cingulum (Figs. 5.42a, 5.44f, 5.45c). It is structured like a

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Fig. 5.42 Schematic drawings showing general morphological features and organelles of dinoflagellates; a ventral view of a flagellated cell of the thecate Alexandrium (Dinophyceae, Gonyaulacales); b a generalized motile dinoflagellate cell showing ultrastructural characteristics; c different views of a hypothetical athecate (gymnodinioid) dinoflagellate with traverse and longitudinal flagellum; upper cell in ventral, lower cell in dorsal view, episome in red, hyposome in blue, and furrows in green; a Original of Takeshi Nakayama, published in Phycologia 43 (2004, issue 3, cover), reproduced with permission of Taylor & Francis publisher; b, c modified from Hoppenrath (2017), drawings by Tanja Wilke

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Fig. 5.43 Characteristic morphology of a dinoflagellate cell; different views of Kirithra sigma, a marine athecate dinoflagellate with a peridinin-chloroplast (Ceratoperidiniaceae, culture strain LIMS-PS-2747 from the southern coast of Korea); a surface of ventral view showing sulcus and cingulum. Note, cingulum ends are slightly overlapping and displaced from each other; b surface focus of right ventrolateral view showing longitudinal flagellum (arrow); c surface focus of left lateral view; d surface focus of dorsal view; from Hu et al. (2020), reproduced with permission of Taylor & Francis publisher and International Phycological Society

ribbon anchored on its inner edge to the cell. It winds to the left side of the cell encircling it. The transverse flagellum has simple hairs of varying lengths (Figs. 5.42a, 5.44f, 5.45c). The hairs, together with the undulating movements of the ribbon, impart spinning movements that let the cells rotate in the direction of the wave. This movement gave dinoflagellates their name (dino is Greek for “whirling”). Besides turning force, there is also forward propulsion by the transverse flagellum. The longitudinal flagellum has few or no hairs and is located in the sulcus (Figs. 5.42a, b, 5.43f, 5.44e). It extends lengthwise, usually beyond the sulcus, so it can be visible behind the swimming cell. In some dinoflagellates, it can contract rapidly up to the cell body. Prorocentroid dinoflagellates have the two flagella inserted apically with the longitudinal flagellum moving anteriorly and the transverse encircling the apex (desmokont flagellation) (Fig. 5.46). Other cell structures and complex organelles Complex organelle structures are characteristic of the dinoflagellates. For example, there are peripheral structures that can be discharged from the cell body into the environment as a defensive response when cells are irritated, and these occur in several types. The most common are trichocysts which are very similar to those of ciliates (Fig. 5.42b). Trichocyst release causes a jet-propulsive response that could be useful in escaping from predators. The mucocyst is a simple sac with granular or fibrous contents, associated with the release of mucoid material (Saldarriaga and Taylor 2017). The pusule is an unusual internal vacuole or array of tubular membranes (ultrastructural different types are known) which is open at the flagellar

base (Fig. 5.42b). Its function is still unknown. It may be used for excretion and uptake processes (Saldarriaga and Taylor 2017). In addition, there is a large variety of eyespot types, unprecedented among all algae (Moestrup and Daugbjerg 2007; Hoppenrath 2017; https://www.dinophyta. org/morphology-of-dinoflagellates/). Mitochondria are with tubular cristae. Some of the most complex ultrastructural organelles known in unicellular organisms are found in a small number of athecate dinoflagellates, especially the warnowiids and polykrikoids (Hoppenrath et al. 2009). These include nematocysts (“harpoons”) used to capture prey and ocelloids (“eyes”) consisting of subcellular structures reminiscent of lenses, corneas, and retinas that are built from different endosymbiotically acquired components (Gavelis et al. 2015).

5.2.1.2 Coccoid Life Stages, Trophic Cysts, and Dormant Stages Some photosynthetic dinoflagellates are predominately metabolically and reproductively active as nonmotile cells without flagella (Fig. 5.5h, i). They represent coccoid life stages which usually lack amphiesmal vesicles. Rather they are surrounded by a continuous wall of cellulose fibers, homologous with the theca (pellicle), and are then difficult to identify as dinoflagellates. In a broad sense, those coccoid cells may be regarded as cysts which are metabolically active rather than dormant (trophic cysts; Saldarriaga and Taylor 2017). Symbiotic dinoflagellates also live in nonmotile stages, and their fibrous continuous wall may be greatly reduced within the host tissue. Another type of cysts is formed in dinoflagellates as dormant stages (resting stages, most often hypnozygotes) which are resistant to

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b Fig. 5.44 General morphology and diversity of athecate dinoflagellates (Dinophyceae, Gymnodiniphycidae, and Noctilucales); a Amphi-

dinium operculatum (Amphidiniales), a marine benthic species with dorsoventrally flattened cells in surface focus, ventral view; note intense color of peridinin-chloroplasts, the epicone is crescent-shaped and left-deflected, hypocone rounded; from interstitial water of sandy sublittoral sediment, Elba, Italy, Mediterranean Sea; b, c Nusuttodinium aeruginosum (Gymnodiniales) from a pond in the Spessart midlands, Germany, a freshwater species with a kleptochloroplast originating from the Cryptista; note blue-green color of the phycobilin-type chloroplasts; b surface focus of ventral view; note cingulum and sulcus with longitudinal flagellum emerging (arrow); c surface focus of dorsal view; d Amphidinium stirisquamtum, a marine benthic species (Amphidiniales, culture strain isolated from wet beach sand collected from the East China Sea), scanning electron microscopy image, ventral view of a cell showing the epicone shape, ventral ridge, points of flagellar insertion; note the cell body surface covered with icicle-shaped body scales; e figure composed of phase contrast microscopy images of three different species with cells in ventral view; (1) Karenia brevis (Kareniaceae), a marine phytoplankton species with fucoxanthin-type chloroplasts originating from the Haptista; it can reach higher density populations and then form red tides with cells that release brevetoxins, deadly for marine vertebrates; note traverse flagellum in the cingulum (arrow), long emergent longitudinal flagellum, and prominent nucleus (nu); (2) Akashiwo sanguinea (incertae sedis Dinoflagellata, culture strain CCMP1593) a species of marine phytoplankton that can form blooms which may cause fish kills; ovoid cells with hypocone can be deeply indented, emergent longitudinal flagellum; (3) a chain of four interconnected cells of the marine planktonic species Gymnodinium catenatum (Gymnodiniales, culture strain CCMP1937) which is the causative organism of paralytic shellfish poisoning (PSP), a neurotoxic poisoning; scale bars, 10 µm; f Gymnodinium baicalense, scanning electron microscopy images (Gymnodiniales, culture strain isolated from the phytoplankton of the ice-covered freshwater Lake Baikal, Russia); upper left cell in left lateral view, lower right cell in dorsal view; note emergent longitudinal flagellum (lf) and transversal flagellum (tf); g Polykrikos kofoidii (Gymnodiniales) a heterotroph marine phytoplankton species without chloroplasts, the species forms pseudocolonies consisting of four zooids that have fused sulci, i.e. with four cinguli, flagella pairs, and two nuclei (nu), upper left cell in surface focus, lower right cell at deeper focus of ventral view; h Noctiluca scintillans (Noctilucales), large colorless cells from a bloom at the coast of Fuerteventura Island, Spain, in 2016. The cells are without chloroplasts and with a characteristic long tentacle (arrow) located together with one scarcely visible emergent flagellum at an invagination (“oral pouch”) of the cell. The species feeds on all kinds of particles and, therefore, can have an enormous impact on the populations of phyto- and zooplankton as well as bacterial populations. In some oceans, the species can harbor a photosynthetic flagellated green alga, Pedinomonas noctilucae (Chlorophyta). The cells can accumulate at the sea surface in large numbers, causing spectacular red discoloration of large areas of the sea; such mass occurrence may cause fish kills by oxygen depletion when the cells are degraded or by release of ammonia (Hoppenrath et al. 2009). b, c courtesy of Burkhard Büdel, originals of Dieter Mollenhauer, d reproduced from Luo et al. (2021), with permission of Algae, the Korean Society of Phycology; e courtesy of Robert A. Andersen; f reproduced from Annenkova et al. (2020) with permission of Taylor & Francis, British Phycological Society; h courtesy of Emilio Soler Onís, with permission of the BEA Culture Collection of Microalgae and Cyanobacteria

adverse conditions (Fig. 5.46e–g). In most cases, sexual processes are leading to those cysts. They have a thickened cell wall and polymerized storage products inside (Fig. 5.46e, f). Resting cysts allow the cells to endure a period of unfavorable environmental conditions. They also serve as a genetic reservoir because they may remain viable in the sediments of coastal waters for years until favorable environmental conditions trigger their germination. The resting cysts can even be fossilized, and virtually all fossil records of dinoflagellates appear to be cysts (Saldarriaga and Taylor 2017).

5.2.1.3 Unique Molecular Traits Dinoflagellates possess numerous unique cellular, molecular, and biochemical traits. Uniquely among all eukaryotes, the dinoflagellate nucleus (dinokaryon) contains chromosomes that are permanently condensed in a liquid-crystalline state throughout the cell cycle. The chromosomes lack histones. In place of them, the DNA packaging system involves unique proteins with the closest similarity to viruses and is non-nucleosomal. These nuclear features coincide with a dramatically increased size of genomes. Dinoflagellate genomes are usually 10 to 100 times larger than the human genome as indicated by flow cytometry measurements (LaJeunesse 2005; Saldarriaga and Taylor 2017). In contrast,

the dinoflagellates’ organelle genomes are small, fragmented, and contain fewer genes than those of other eukaryotes. Dinoflagellate mitochondrial genomes are some of the smallest known; they have been found to encode no more than three proteins. There are even cases where essential mitochondrial functions have apparently been fully transferred to the nucleus. This resulted in functional mitochondria but without a mitochondrial genome (John et al. 2019). The chloroplasts that contain the carotenoid peridinin are most common in the dinoflagellates. Their genomes are fragmented into numerous minicircles. There is usually one gene per circle flanked by a variety of non-coding sequences (Howe et al. 2008; Saldarriaga and Taylor 2017; Sibbald and Archibald 2020). The absolute number of genes coded by the peridinin-chloroplasts is reduced, i.e. no more than 12–14 genes have been found (John et al. 2019), which is about one-tenth of the gene content of other algal chloroplasts. Some of the missing genes have been moved to the nucleus in some species or lost altogether (Wang et al. 2008; Yoon et al. 2005). In some species, the minicircles may be even located in the nucleus, not in the plastids (Moestrup and Daugbjerg 2007; Saldarriaga and Taylor 2017). Collectively, the unique cytological, ultrastructural, and genetic attributes of dinoflagellates present still intriguing topics of research.

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b Fig. 5.45 General morphology and diversity of thecate dinoflagellates (Dinophyceae, Peridiniphycidae); a, d, j living cells with

peridinin-chloroplasts; b, c, e–i scanning electron micrographs of thecae (cell walls); a, b Lingulodinium polyedra (Gonyaulacales), a species that can form spectacular red tides (blooms), e.g. at the coast of California, and then produce toxins harmful to both humans and marine life; it is also capable of bioluminescence; a surface focus on ventral view; b details of theca, ventral view from anterior mainly on the epitheca; c Fragilidium mexicanum (Gonyaulacales, culture strain isolated from Jinhae-Masan Bay, South Korea), morphology of thin theca with smooth surface, the pattern formed by amphiesmal vesicles (alveolae) containing cellulosic plates (indicated by numbers and primes, Kofoid system; see Hoppenrath 2017) is used to describe and identify dinoflagellate species; d Peridinium cf. bipes (Peridiniales), a freshwater species (from a garden pond at Spessart midlands, Germany), focus is of various views, e.g. ventral (upper left cell), and dorsal (upper right cell); e Parvodinium elpatiewskyi (Peridiniales), a common freshwater dinoflagellate, cell of a culture strain established from a pond in Berlin, Germany, laterodorsal view; note the hypotheca is with various spines on its sutures, the pattern of cellulosic plates (tabulation, indicated by numbers and primes) is shown; f Gonyaulax cf. polygramma (Gonyaulacales), a marine phytoplankton species, ventral view, cell with wall composed of cellulosic plates; the cell shows an asymmetry due to a descending cingulum; g Corythodinium reticulatum (Oxytoxaceae), a tropical marine phytoplankton species, left lateral view, elongated hypotheca with an antapical spine, a small epitheca, and a broad cingulum; h Podolampas bipes (Podolampadaceae), a marine phytoplankton species with pyriform cells, and pronounced antapical spines but without a depressed sulcus and cingulum; i Tripos furca (Gonyaulacales), a marine phytoplankton species, dorsal view, characterized by its straight body, the epitheca is gradually tapering into an anterior horn; the two antapical horns are of different lengths and closed; j Tripos horridus (Gonyaulacales), a marine phytoplankton species with peridinin-chloroplasts, a slender almost triangular body and three thecal horns, i.e. one apical horn and two open antapical horns that are bent anteriorly; d, j courtesy of Burkhard Büdel, c reproduced from Li and Shin (2019) with permission of Taylor & Francis, International Phycological Society, e reproduced from Kretschmann et al. (2020) with permission of Elsevier publisher

5.2.2 Reproduction Vegetative reproduction of dinoflagellates is by binary fission, which is accomplished in two ways in thecate species. It either divides the protoplast inside the mother cell wall and both offspring leave the theca, or the protoplast first hatches and then divides. Both cases are followed by the formation of a complete new theca (eleutheroschisis; Hoppenrath et al. 2009). Or the theca may be shared by the offspring with the synthesis of the missing components (desmoschisis). For example, in the genus Ceratium/Tripos, each daughter cell receives half of the parental wall components and regenerates the missing portion. Sexual reproduction is rather diverse. Dinoflagellates follow a haplontic life cycle, with post-zygotic meiosis (for a general scheme, see Fig. 2.9). That is, the vegetative dinoflagellate cells are haploid. The gametes resemble regular motile cells and fusion is slow which makes sexual reproduction hard to observe. Syngamy may involve equal (isogamy) or unequal (anisogamy) motile gametes. Fusion of gametes may occur within a clonal strain (homothallism) or only between different clonal strains (heterothallism). The product of fusion is a zygote with three or four flagella, i.e. with two longitudinal flagella in addition to one or two transverse ones. The young zygote remains still motile for hours or days before in most cases a nonmotile resting cyst is formed. Meiosis can be accomplished before the release of daughter cells from the cyst, or a motile zygote is released first from the cyst which then undergoes meiosis.

5.2.3 Chloroplasts The photosynthetic dinoflagellates present an evolutionarily diverse collection of different chloroplast types of various

origins (Fig. 5.47). The diversity in types of photosynthesis that exist within dinoflagellates is unparalleled within any group of eukaryotes (Schnepf and Elbrächter 1999). Using phylogenetic and genomic information, it is interpreted that the dinoflagellates have a remarkable ability to remodel their genomes through endosymbiosis (Bhattacharya et al. 2004; Yoon et al. 2005). Dinoflagellate chloroplasts (or plastids) have repeatedly been reduced, lost, and replaced by new plastids, leading to a spectrum of ages and integration levels (Hehenberger et al. 2019; Sibbald and Archibald 2020; Sarai et al. 2020). Many dinoflagellates have lost the plastid genome, or even lost the plastid itself. There are five main groups of dinoflagellate chloroplasts based on their general pigment types: Most common are the peridinin-type chloroplasts which originated from a red algal cell through secondary endosymbiosis. The peridinin-type probably represents the ancestral state (Janouškovec et al. 2010). There are four types of chloroplasts which arose from processes where the peridininchloroplasts have been replaced for chloroplasts with completely different origins (Fig. 5.47). Many dinoflagellates have replaced or supplemented the peridinin-plastid with new secondary or tertiary plastids, or kleptochloroplasts. Tertiary endosymbiosis is referred to as the acquisition of another plastid by engulfment of a plastid that evolved by a secondary endosymbiosis (Stoebe and Maier 2002; Bhattacharya et al. 2004; Keeling 2010). Dinoflagellates can also have chloroplasts from two successive (serial) secondary endosymbioses during their evolutionary history (Keeling 2010; Sarai et al. 2020). Only the dinoflagellates have undergone tertiary endosymbiosis. These chloroplasts, like those from serial secondary endosymbioses, all have in common that they are stably integrated with their host and are retained over long periods of evolutionary time (Hehenberger et al. 2019). Tertiary endosymbiosis also

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b Fig. 5.46 General morphology and diversity of thecate dinoflagellates (Dinophyceae) continued; Dinophysales (a–b), Prorocentrales

(c), Gonyaulacales (d), Peridiniales (e, f) and nonmotile stages of athecate Dinoflagellates (g–i); a Dinophysis tripos, marine phytoplankton species with reddish cryptophyte kleptochloroplasts; inset: schematic drawing of dinophysoid tabulation with characteristic serrated sagittal suture splitting the theca in two lateral halves in yellow; b Phalacroma porodictyum (Dinophysales), a marine species with rounded cell with cingulum that divides the cell into a small epitheca and a large hypotheca; c Prorocentrum cassubicum (Prorocentrales, culture strain SAG 40.80) in lateral view, with theca that consists of two major plates (right and left) and flagellation not associated with furrows (Hoppenrath et al. 2013); note brownish color of peridinin-chloroplast with a prominent pyrenoid with starch-sheath visible as ring; inset: schematic drawings of a prorocentroid cell showing the desmokont flagellation (both flagella arise from one apical pore, and prorocentroid tabulation with the two main thecal plates in blue, the periflagellar area with platelets in purple and characteristic sagittal suture splitting the theca in two lateral halves in yellow; d Ceratocorys horrida (Gonyaulacales, culture strain CCMP157), a marine photosynthetic species with bioluminescence, round-shaped body with prominent emanating spines; e, f resting cysts of Protoperidinium sp. (Peridininales), a species of marine phytoplankton (from the German Bight, North Sea); e enclosed within theca; f release of cyst from theca; g nonmotile stage of the athecate Kirithra sigma (Ceratoperidiniaceae; see Fig. 5.43) covered by a hyaline membrane, with cingulum (black arrows) and nucleus (N); h, i Pyrocystis lunula (Gonyaulacales, culture strain SAG 2014, isolated from the North Sea), a nonmotile marine species with spindle-shaped cells covered by a thin wall; the species is well known for its bioluminescence capacity; i dark field microscopy; d courtesy of Robert A. Andersen; h courtesy of Tatyana Darienko, with permission of the SAG culture collections; a, c, (inset) www.dinophyta.org, drawings by Tanja Wilke; g reproduced from Hu et al. (2020), with permission of Taylor & Francis, International Phycological Society; i courtesy of Burkhard Büdel

resulted in fully functional chloroplasts, i.e. with gene transfer to the host’s nucleus so that the chloroplast includes proteins encoded in the host’s nucleus. In dinoflagellates, there are also many temporary associations where an alga or a ciliate is engulfed or sucked out, and its plastid(s) are taken up for a period of time but ultimately digested. These are called kleptochloroplasts or “stolen” plastids. Kleptoplasty is a common phenomenon observed besides dinoflagellates in many other eukaryotic lineages, e.g. the foraminiferans, ciliates, and even some animals such as sea slugs and flatworms (Sibbald and Archibald 2020). The uptaken chloroplasts may persist anywhere from days to months until they are replenished by taking up fresh algal prey (Hehenberger et al. 2019). In Dinophysis, transitional stages from kleptochloroplasts to permanent chloroplasts were documented (Rusterholz et al. 2017; Garcia-Cuetos et al. 2010). However, it is not always easy to distinguish between true chloroplasts and kleptochloroplasts. A particular set of accessory pigments in addition to chlorophylla characterizes the peridinin-chloroplasts (plastids), i.e. chlorophyll c2 and ß-carotene, and a special type of carotenoid, peridinin, which can be found only in dinoflagellates. The peridinin-chloroplasts are shaped as individual plate-like bodies with one or more pyrenoids, which often form a network, sometimes a single stellate chloroplast. They have envelopes consisting of three membranes. The thylakoids are packed into three. There is no girdle lamella as in the stramenopile algae (Heterokontophyta; see Sect. 5.1). Storage products in species with peridinin-chloroplast are starch, produced exterior to the plastid, and oils. Peridinin-chloroplasts have another type of rubisco that has a much lower specificity for CO2 over O2 when compared to the more common “eukaryotic” rubisco found in other algae. The dinoflagellates’ rubisco is a “bacterial type II” rubisco that evidently arose from a lateral gene transfer.

The fucoxanthin (19’hexanoyloxyfucoxanthin)-type chloroplasts, with the chlorophylls a and c, originated from the Haptista (Tengs et al. 2000). They can be found, e.g. in Karenia, Karlodinium, or Takayama. They lack peridinin, but instead use 19’hexanoyloxyfucoxanthin which is a carotenoid typically found only in the Haptista (Ishida and Green 2002; Wang et al. 2008). True haptophyte-derived chloroplasts were found to have two independent origins, i.e. the Karenia and Karlodinium tertiary plastids appear to be derived from two different haptophytes (Tengs et al. 2000). The haptophyte tertiary chloroplasts are surrounded by four membranes instead of three as in the peridinin-chloroplasts. They also contain normal plastid-encoded form I rubisco. Gene transfers to the host’s nucleus have occurred in the taxa with haptophyte-derived plastids through tertiary endosymbiosis (Ishida and Green 2002; Patron et al. 2006). In addition to the true chloroplast of haptophyte origin, there are also haptophyte-derived kleptoplastids in dinoflagellates (Gast et al. 2007; Hehenberger et al. 2019; Sibbald and Archibald 2020). The diatom-derived plastids contain the chlorophylls a and c plus the brown carotenoid fucoxanthin (Inagaki et al. 2000). A whole diatom cell is then located within the cytoplasm of the dinoflagellate host, separated from the host by only a single membrane, lacking any trace of the siliceous diatom wall. Usually, not only the diatom chloroplast is present but also the diatoms’ nucleus and mitochondria (Fig. 5.47). The type of dinoflagellates with chloroplast originating from diatoms is called dinotoms (Imanian et al. 2010, 2012; Yamada et al. 2017). Although dinotoms are just a small group of closely related dinoflagellates, their morphologies are quite diverse. Examples can be found in the thecate flagellated genus Durinskia and the athecate predominantly nonmotile or filamentous colony-forming Dinothrix (Hoppenrath et al. 2014; Yamada et al. 2020). In addition to the dinotoms where the diatom is retained permanently in the dinoflagellates’ cytosol, there is also a

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Fig. 5.47 Diversity of the chloroplast types of dinoflagellates. Their plastids may be derived from a secondary or tertiary endosymbiosis, a serial secondary, or tertiary endosymbiosis, or even a “kleptoplasty” origin; modified from Hoppenrath et al. (2018)

case of a diatom-derived kleptoplastid, i.e. in Durinskia capensis (Yamada et al. 2019; Sibbald and Archibald 2020). Another association with stramenopile algae has been described for species of the dinoflagellate genus Podolampas (Fig. 5.45h). There, members of the class Dictyochophyceae gave rise to chloroplasts (Schweikert and Elbrächter 2004). However, whether the Dictyochophyceaederived chloroplasts are from tertiary symbiosis or represent kleptoplastids is still unclear.

Chlorophylla + b-type chloroplasts are currently known from three lineages of dinoflagellates. In the genus Lepidodinium (Watanabe et al. 1987, 1990) and two more independent so far undescribed dinoflagellate species (Sarai et al. 2020), the chloroplasts are derived from green algae (Chlorophyta), i.e. members of the Pedinophyceae. While the three dinoflagellates are not closely related to each other, their chloroplast likely arose from very close relatives of the same green algal genus, i.e. Pedinomonas (Kamikawa et al.

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2015; Sarai et al. 2020). The chlorophyll a + b chloroplasts are from two successive secondary endosymbioses with an endosymbiotic gene transfer (EGT) of the endosymbiont genes to the host nuclear genome (Fig. 5.47). The two so far unidentified dinoflagellates species even contain residual nuclei, which are interpreted as nucleomorphs. The host-endosymbiont is less advanced in the two dinoflagellates. An EGT is still in progress (Sarai et al. 2020), in contrast to the nucleomorphs of Chlorarachniophyta and Cryptista, which are seen in a post-EGT state (Curtis et al. 2012). The phycobilin-type chloroplasts are cryptophyte kleptoplastids (Fig. 5.47); they arose from cells of the Cryptista (Ishida and Green 2002; Patron et al. 2006). The chloroplasts contain phycoerythrin and alloxanthin but lack peridinin. They have paired stacked thylakoids with electron-dense contents, similar to cryptophyte thylakoids. Prominent examples of phycobilin-type dinoflagellates are the marine potentially toxic phytoplankton species of Dinophysis (Fig. 5.46a). An example from freshwaters is Nusuttodinium aeruginosum (Fig. 5.44a, b; Schnepf et al. 1989). In contrast to Cryptista’s chloroplasts with four membranes, the plastids of Dinophysis are surrounded by only two membranes (Schnepf and Elbrächter 1988). A key for understanding the origin of phycobilin-type plastids in Dinophysis species is their ability to uptake food particles by myzocytosis, by which only the prey’s cell content is drawn into a food vacuole (Hansen 1991; Schnepf and Elbrächter 1992). The phycobilin-type plastids are kleptoplastids (Sibbald and Archibald 2020). Dinophysis species are an example that it has been inconclusive for a long time whether the phycobilin-type plastids represent true plastids or kleptoplastids (Rusterholz et al., 2017; Garcia-Cuetos et al., 2010); it can be difficult to distinguish between them (Hackett et al. 2003).

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transcriptionally active nuclei. Plastid metabolism depends on the nuclear genome for more than 90% of the needed proteins (Rumpho et al. 2008). A large number of genes are necessary to stabilize the function and persistence of an uptaken chloroplast. Kleptoplasty is considered a strategy to receive the nutritional benefits of photosynthesis, possibly an early evolutionary stage in the permanent acquisition of chloroplasts. Alternatively, it may be a strategy that allows for nutritional flexibility to better adapt to environmental or seasonal changes (Gast et al. 2007). Kleptoplasty is not restricted to dinoflagellates. It is also known for ciliates (Alveolates), foraminifers, and Metazoa. A unique strategy to keep the uptaken chloroplasts functional has been developed by the marine ciliate Myrionecta rubra. It steals plastids from the cryptomonad Geminigera cryophile. To compensate for the absence of numerous nuclear-encoded genes involved in regulation, division, and other chloroplast functions, it accumulates prey nuclei which, after feeding, are retained for some time while servicing the plastids derived from multiple cryptophyte cells (Johnson et al. 2007). Interestingly, Dinophysis cannot feed directly on cryptophytes but by myzocytosis on Myrionecta (Nagai et al. 2008; Kim et al. 2012) and thus utilizing kind of “second-hand kleptochloroplasts”. The sea slug taxon Sacoglossa comprising several genera (Gastropoda: Opisthobranchia; Händeler et al. 2009) acquire chloroplasts temporarily by engulfing algae and retaining their chloroplasts in a functional state. Chloroplasts preferentially from large-celled and coenocytic algae, i.e. the ulvophyceaen green algal genus Codium and other members of Caulerpales, Bryopsidales, Cladophorales, and Dasycladales as well as the xanthophyte Vaucheria are retained (Rumpho et al. 2008; Pelletreau et al. 2011). In the sea slug Elysia chlorotica, plastids ingested from its algal food source Vaucheria litorea, combined with horizontal gene transfer, enable the animal to live from photosynthesis for months (see Sect. 6.1).

5.2.4 Kleptoplasty Kleptoplasty (or Kleptoplastidy) is a phenomenon related to photosynthesis acquisition and means the retention of a foreign organelle, i.e. functional chloroplasts from algal prey. Non-photosynthetic dinoflagellates may reacquire photosynthesis through the temporary use of plastids from their prey (stolen chloroplasts; Schnepf and Elbrächter 1999). It can also be connected to horizontal gene transfer. The acquired chloroplasts remain functional only for a limited period of time, usually not more than several days, until they are digested as the rest of the prey and then need to be replaced with chloroplasts from newly fed prey. This is because the full chloroplast functions rely on

5.2.5 Non-Photosynthetic Nutrition With respect to their nutrition strategies, the dinoflagellates are exceptional among all algal life forms. In nearly all different dinoflagellate lineages, heterotrophic and phototrophic forms can be found. Mixotrophy appears to be the most common mode of nutrition. Approximately only half of the dinoflagellate species possess the photosynthetic capability, but completely autotrophic species are rare (Schnepf and Elbrächter 1992). The other half of the species are non-photosynthetic and traditionally considered to lack plastids. Despite being non-photosynthetic, it is

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hypothesized that at least all free-living (not parasitic) heterotroph dinoflagellates still rely in their bio-compound syntheses on plastid organelles that are derived from an ancestral plastid (Janouškovec et al. 2017). Nutrition by phagotrophy, i.e. feeding on particles like taking up entire food organisms (phagocytosis), is very common among Dinoflagellates. Prey capture mechanisms in phagotrophic forms vary greatly (Saldarriaga and Taylor 2017). In large species, the prey is taken up by a distinct cell mouth (cytostome), i.e. the cells are able to stretch as they engulf intact prey cells and digest them within a food vacuole inside the cell. Surprisingly, also some armored dinoflagellates are capable of ingesting whole cells. Other dinoflagellates can extend a delicate, pseudopodial “feeding veil” (pallium) with which they surround their large prey. Digestion occurs outside the theca, and only digested material is taken up; the veil is retracted afterward. Another feeding form is myzocytosis which involves piercing the cell membrane of prey cells with a special organelle (peduncle) and somehow “sucking” the prey cell’s content as if through a straw (Schnepf and Elbrächter 1992; Saldarriaga and Taylor 2017). Besides phagotrophy, also dissolved organic matter can be taken up (osmotrophy).

5.2.6 Bioluminescence A number of marine free-living dinoflagellates are capable of bioluminescence and emit light when disturbed (Hardeland and Hoppenrath, 2013). This bioluminescence results in a “sparkling” effect that can often be observed in coastal seas and bioluminescent bays. It is controlled by circadian rhythms and only occurs at night. Prominent examples are Noctiluca scintillans (Fig. 5.44h) in coastal temperate regions and Pyrocystis lunula (Fig. 5.46h, i) in tropical oceans. The luminescence occurs as brief blue flashes that have been seen to emanate from individual cytoplasmic bodies (scintillons), which contain luciferase, the main enzyme involved in dinoflagellate bioluminescence. The lowering of pH initiated by a mechanical disturbance that deforms the cell’s plasma membrane triggers the light-producing reaction. To explain the benefits of bioluminescence to dinoflagellates, it has been suggested that shearing stress caused by the feeding of copepods on dinoflagellates triggers their bioluminescence (burglar-alarm hypothesis). It is then used by visual predators like fish and squid to find their zooplankton (copepod) prey. Alternatively, bioluminescence may startle predators and discourage them from feeding on dinoflagellates.

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5.2.7 Toxins and Harmful Algal Blooms Dinoflagellates often are the source of dense toxic Harmful Algal Blooms (HABs) worldwide (Lassus et al. 2016). Biotoxin-producing dinoflagellates are photosynthetic. Dinoflagellates represent the main part of toxic species that can bloom. Currently, about 95 toxic species are known. HABs have an enormous impact on ecosystem functions, e.g. alterations of marine trophic structure with the death of marine mammals, fish, and seabirds. They can also pose economic problems for fisheries due to shellfish and fish contaminated with biotoxins and impairment of tourism and recreational activities (Anderson et al. 2012). Among closely related species, there may be toxic and non-toxic ones. Even within the same species, there can be toxic and non-toxic strains. Several marine species of dinoflagellates can produce potent toxins which can cause massive kills of fish and other marine life. In contrast to the situation in marine environments, there are only a few reports of harmful dinoflagellates from freshwaters. Some freshwater species can form blooms, e.g. in subtropical waters which often are associated with water column stratification (Moestrup and Calado 2018). Up to now, the functions of most of the dinoflagellate biotoxins are unknown. It looked like they do not prevent predation on the producers as most of their predators such as copepods (small planktonic crustaceans) or bivalve mollusks remain unharmed. But there are first indications for some substances having an impact on competitors or predators. Toxic and bioactive substances produced by dinoflagellates show high structural diversity. For example, in a strain of Protoceratium reticulatum 90 different yessotoxin derivatives were detected. There are two types of dinoflagellate biotoxins. One type comprises heterocyclic guanidines such as the saxitoxin complex. They are water-soluble small molecular weight substances that block the entry of sodium into the nerves. Saxitoxin affects the nervous system and paralyzes muscles, thus the term “paralytic” shellfish poison. When shellfish (mollusks like clams, mussels, and oysters), which are filter feeders, eat saxitoxin-producing cells, the toxin accumulates in their tissue. Eating the contaminated shellfish can then cause severe illness in humans and high levels of the toxin, even death. Saxitoxin is way more potent than cyanide and multiple times more toxic than curare (Sako et al. 2001). The other type of toxins consists of larger water- or lipid-soluble (polyether) compounds, such as ciguatoxin, maitotoxin and brevetoxin. Toxin from Gambierdiscus toxicus is considered one of the most potent

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biogenic toxins. The brevetoxin complex from Karenia brevis kills fish and causes neurotoxic shellfish poisoning impairing human health (with symptoms including vomiting and nausea). When taken orally by fish with its food, both biotoxins have been shown to accumulate in fish tissue (mostly the liver). Among the dinoflagellates responsible for HABs, the genus Alexandrium is one of the most important in terms of the severity, diversity, and distribution of bloom impacts. At least half of the currently known Alexandrium species are known to be toxic or have otherwise harmful effects (Anderson et al. 2012). Three different families of known toxins are produced among Alexandrium species—this toxigenic diversity is not found in any other HAB genus. The nutritional strategies of the toxic Alexandrium species are diverse, including feeding by ingesting other organisms. Many Alexandrium species have complex life histories that include cyst formation, which offers considerable ecological advantages (see below; Anderson et al. 2012). Ciguatera fish poisoning (CFP) is the most common non-bacterial disease in the world, which results in 50,000– 500,000 cases per year worldwide (Hoppenrath et al. 2014). It is caused by the accumulation of toxins produced by species of the genus Gambierdiscus. Distribution and abundance of the toxic Gambierdiscus species are reported to correlate positively with water temperature. Consequently, there is growing concern that increasing temperatures associated with climate change could increase the incidence of CFP (Tester et al. 2010). Biotoxin-producing dinoflagellates are usually present in marine water but in low numbers and then cause no problems. But when the algae “bloom”, the increased algal biomass becomes a greater food source for shellfish, and more biotoxin accumulates. The dinoflagellate biotoxins do not harm shellfish, so the toxin level in their tissue will rise until the bloom subsides. When the blooms end and the dinoflagellate cells die, biotoxins are released into the water. Then fish can take the biotoxins via their gills, a more direct way into their bloodstream, and in these cases, the effects of the toxin are much more severe, e.g. fish kills (Saldarriaga and Taylor 2017). The ability of dinoflagellates to form resistant cysts related to bloom decline is key to the long-distance transport of dinoflagellates through ocean currents. Cyst germination, i.e. reviving the population of swimming cells, may provide inocula for new proliferation events. These natural processes have been augmented by human activities, e.g. ballast water discharge, leading to significant (human-mediated) range extensions of HAB species. The high nutrient content of coastal surface water and particular climatic conditions which lead to low salinity (which often occurs during rainy periods) and calm seas, e.g.

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rainy days followed by sunny weather represent favorable conditions for bloom formation.

5.2.8 Phylogeny: Classification The dinoflagellates are one of three main lineages of Alveolates, together with the apicomplexans and ciliates. In addition, there are a few little-known ancestral lineages, i.e. the Colpodellida and the heterotrophic lineages Perkinsidae and Colponemida sensu Adl et al. (2019) (Fig. 5.48; John et al. 2019). The apicomplexans (Apicomplexa) are non-photosynthetic, obligate intracellular parasites that are causes of serious human diseases such as malaria and toxoplasmosis. Their remnant plastid (apicoplast) contains a reduced (35 kb) genome. Early in the evolution of the ciliates (Ciliophora), the plastid was presumably lost (Bhattacharya et al. 2004; Yoon et al. 2005). Based on evidence from molecular phylogenies and genome data, all dinoflagellates and the apicomplexans arose from a common ancestor which was photosynthetically active (Fig. 5.47; e.g. Janouškovec et al. 2010; John et al. 2019), but only half of the known core dinoflagellates still maintain photosynthesis (Gómez 2012). There are only two photosynthetic genera of Colpodellida known, i.e. Chromera and Vitrella (Adl et al. 2019; John et al. 2019). More than 2,500 extant species have been assigned to about 300 genera (Taylor et al. 2008; Hoppenrath 2017). About the same number of fossil species have been recorded. The earliest fossil taxa assigned unequivocally to the dinoflagellates are about 250 million years old, but they may be even older (Fensome et al. 1993). The major dinoflagellate lineage is distinguished in so-called core dinoflagellates, Dinophyceae, and several basal lineages such as the Syndinians or Syndiniales (parasitic dinoflagellates without plastids) and the genus Oxyrrhis. The latter is an obligate heterotrophic taxon with phagotrophic nutrition (Fig. 5.48; John et al. 2019). Within the core dinoflagellates, the Noctilucales are most basal (e.g. Noctiluca, an obligate heterotrophic species without plastids, capable of luminescence, Fig. 5.44h). The single monophyletic origin of all dinoflagellates is well supported. There were two contrasting classifications for the dinoflagellates, i.e. one that considers phylogenetic relationships (as revealed by molecular phylogenies) rather than taxonomic hierarchies, and the other following the traditional morphology-based circumscription of higher level taxa such as classes and orders of dinoflagellates; see Hoppenrath (2017) for discussion. For the phylogenetic classification, we follow Adl et al. (2019) here; for the morphology-based classification, Fensome et al. (1993).

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Kryptoperidinium foliaceum Scrippsiella trochoidea Heterocapsa spp. 96/94 Symbiodinium spp. Polarella glacialis Alexandrium spp. Lingulodinium polyedrum Togula jolla Gymnodinium catenatum Karlodinium spp. Karenia brevis Amphidinium carterae Noctilucales Noctiluca scintillans Amoebophrya sp. (parasite of Akashiwo) 95/93 Amoebophrya ceratii (parasite of Alexandrium) Amoebophrya sp. (parasite of Karlodinium) Hematodinium sp. Oxyrrhis marina Colponemida Perkinsus marinus Perkinsidae Plasmodium falciparum Toxoplasma gondii Apicomplexa Cryptosporidium spp. Chromera velia Colpodellida Vitrella brassicaformis Tetrahymena thermophila Oxytricha trifallax Ciliophora Protocruzia adherens Thalassiosira pseudonana Ectocarpus siliculosus Stramenopiles Saprolegnia parasitica Schizochytrium aggregatum

Fig. 5.48 Phylogenetic position of dinoflagellates within the alveolates showing the non-photosynthetic lineages, Syndiniales and Noctilucales, being ancestral to the Dinophyceae (“core dinoflagellates”). The Colpodellida is the earliest (most ancestral) lineage of all photosynthetic alveolates. The phylogeny strongly supports the common ancestor of all alveolates, except for the Ciliophora, which was photosynthetic (green arrow); the phylogeny is based on a concatenated set of 100 conserved nuclear proteins using the maximum likelihood phylogeny inference method; gray filled circles denote maximal statistical (bootstrap) support for the shared (monophyletic) origin of a group of taxa (clade); modified from John et al. (2019)

Several orders as in their traditional circumscription appear as para- or polyphyletic assemblages in molecular phylogenies of the dinoflagellates, e.g. the orders Gymnodiniales and Peridiniales (see below). The Syndiniales and Noctilucales take a basal position within the Dinoflagellata, preceding the two main lineages of the core dinoflagellates, i.e. Gymnodiniphycidae and Peridiniphycidae sensu Fensome

et al. (1993) and Adl et al. (2019). Excellent overviews on the classification and systematics of the dinoflagellates have been presented, for example in Fensome et al. (1993) and Moestrup and Calado (2018). An overview of the current core dinoflagellate classification is shown in Table 5.3.

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Table 5.3 Current core dinoflagellate classification, phylum Dinoflagellata (as based on Adl et al. 2019, not complete and no incertae sedis taxa; numbers in brackets point to schematicdrawings of genera in Fig. 4.49) Phylogenetic lineage

Families and Genera

Noctilucales:

Abedinium, Cachonodinium, Craspedotella (8), Cymbodinium (7), Kofoidinium (4), Leptodiscus, Noctiluca (6), Petalodinium, Pomatodinium (4), Scaphodinium, Spatulodinium

Dinophyceae, Gymnodiniphycidae Amphidiniales

Amphidiniaceae: Amphidinium s.s. (10)

Gymnodiniales sensu stricto

Gymnodinium (18), Barrufeta, Dissodinium (17), Erythropsidinium (22), Greuetodinium (23), Gymnodinium, Lepidodinium, Nematodinium (21), Nusuttodinium, Pellucidodinium, Polykrikos (19), Proterythropsis (20), Pseudocochlodinium, Wangodinium, Warnowia (20) Gyrodiniaceaea: Gyrodinium s.s. (13) Kareniaceae/Brachydiniaceae Brachydinium (12), Gertia, Karenia (11), Karlodinium, Shimiella, Takayama Ceratoperidiniaceae: Ceratoperidinium, Kirithra

Torodiniales

Kapelodinium, Torodinium, Levanderina, Margalefidinium, Cochlodinium s.s. (14)

Ptychodiscales

Achradina, Amphitolus, Balechina, Ptychodiscus, Sclerodinium Borghiellaceae: Baldinia, Borghiella Sphaerodiniaceae: Sphaerodinium

Tovelliales

Tovelliaceae: Bernardinium, Esopotrodinium, Jadwigia, Tovellia Suessiaceae: Ansanella, Asulcocephalium, Biecheleria, Biecheleriopsis, Freudenthalidium, Halluxium, Leiocephalium, Pelagodinium, Polarella, Prosoaulax, Protodinium, Symbiodinium (35), Yihiella

Dinophyceae, Peridiniphycidae Peridiniales

Amphidiniopsis, Archaeperidinium, Blastodinium (46), Diplopelta (43), Diplopsalis (43), Diplopsalopsis (43), Herdmania, Niea (43), Oblea (43), Palatinus, Parvodinium, Peridinium (45), Peridiniopsis, Preperidinium (43), Protoperidinium (42), Qia (43), Vulcanodinium Thoracosphaeraceae: Aduncodinium, Amyloodinium, Apocalathium, Blastodinium, Chimonodinium, Cryptoperidiniopsis, Duboscquodinium, Ensiculifera, Fusiperidinium, Laciniporus, Leonella, Luciella, Naiadinium, Pachena, Paulsenella, Pentapharsodinium, Pfiesteria, Scrippsiella (40), Stoeckeria, Theleodinium, Thoracosphaera (39), Tintinnophagus, Tyrannodinium Kryptoperidiniaceae (“dinotoms”): Blixaea, Dinothrix, Durinskia (47), Kryptoperidinium, Unruhdinium Heterocapsaceae: Heterocapsa (38) Amphidomataceae: Amphidoma, Azadinium (37) Cladopyxidaceae: Cladopyxis (58), Fensomea

Gonyaulacales

Alexandrium (52), Amylax, Carinadinium, Ceratium (55), Ceratocorys (54), Coolia (49), Fukuyoa (49), Fragilidium (52), Gambierdiscus (49), Goniodoma, Gonyaulax (56), Lingulodinium, Ostreopsis (49), Pentaplacodinium, Peridiniella, Protoceratium, Psammodinium, Pyrocystis (53), Pyrodinium (51), Pyrophacus (50), Sourniaea, Tripos (55) Podolampadaceae: Blepharocysta, Gaarderiella, Heterobractum, Lissodinium, Mysticella, Podolampas (41) Oxytoxaceae: Corythodinium, Oxytoxum

Dinophysales

Amphisolenia (34), Citharistes, Dinofurcula, Dinophysis (30), Histioneis (32), Latifascia, Metadinophysis, Metaphalacroma, Ornithocercus (31), Oxyphysis, Parahistioneis (32), Phalacroma (30), Pseudophalacroma, Sinophysis (29), Triposolenia (33)

Prorocentrales

Mesoporus, Prorocentrum (28)

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Fig. 5.49 Extant dinoflagellate diversity. Only about a quarter of the living genera are shown in this tree. (1) Perkinsus, (2) Oxyrrhis, (3) Amoebophrya, (4) Kofoidinium, (5) Pomatodinium, (6) Noctiluca, (7) Cymbodinium, (8) Craspedotella, (9) Haplozoon, (10) Amphidinium, (11) Karenia, (12) Brachydinium, (13) Gyrodinium, (14) Cochlodinium, (15) Plectodinium, (16) Actiniscus, (17) Dissodinium, (18) Gymnodinium, (19) Polykrikos, (20) Proterythropsis, Warnowia, (21) Nematodinium, (22) Erythropsidinium, (23) Greuetodinium (=Leucopsis), (24) Akashiwo, (25) Protoodinium, (26) Oodinium, (27) Chytriodinium, (28) Prorocentrum, Mesoporus, (29) Sinophysis, (30) Dinophysis, Phalacroma, (31) Ornithocercus, (32) Histioneis, Parahistioneis, (33) Triposolenia, (34) Amphisolenia, (35) Symbiodinium, Polarella (36) Woloszynskia, (37) Azadinium (Amphidomataceae), (38) Heterocapsa (Heterocapsaceae), (39) Thoracosphaera, (40) Scrippsiella, (41) Podolampas (Podolampadaceae), (42) Protoperidinium, (43) Diplopsalis-group (e.g. Diplopsalopsis, Diplopsalis, Oblea, and Preperidinium), (44) Glenodinium, (45) Peridinium, (46) Blastodinium, (47) Durinskia (“dinotoms”), (48) Triadinium (=Heteraulacus), (49) Gambierdiscus (also Fukuyoa, Coolia, and Ostreopsis), (50) Pyrophacus, (51) Pyrodinium, (52) Alexandrium, Fragilidium, (53) Pyrocystis, (54) Ceratocorys, (55) Ceratium, Tripos, (56) Gonyaulax, (57) Paleophalacroma, (58) Cladopyxis, (59) Hemidinium, (60) Gloeodinium, and (61) Stylodinium; from Taylor et al. (2008), partly updated

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5.3

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A Cercozoan Secondary Endosymbiosis: Chlorarachniophyta

Angelika Preisfeld and Burkhard Büdel

5.3.1 General Characters The chlorarachniophytes are a small group of algae with presently 16 species, most of which one won’t come upon easily, because they are with a length between 8 and 20 µm quite small to be found, and the main distribution core area is in tropical and temperate seas. This is even made more difficult by the fact that they occur in three different life stages: As single-celled, photoautotrophic, and mixotrophic amoeboids mostly with pseudopodia, as flagellates (zoospores) with one flagellum and as cysts (coccoid). While some species like Chlorarachnion reptans pass through all three stages (Hibberd and Norris 1984), Bigelowiella natans only displays the flagellate type (Moestrup and Sengco 2001). The type of stages in the life cycle is used as a very important criterion for species discrimination. Besides being able to perform photosynthesis, chlorarachniophytes can use means of phagotrophy to engulf bacteria and other small algae, which generally takes place in pseudopods (Hibberd and Norris 1984) or occasionally, in the cell body (Ishida et al. 1996). Chlorarachniophyta include the classes Chlorarachniophyceae, descendants of a secondary endosymbiosis with a green alga of the Ulvophyceae (Suzuki et al. 2016) and a complex plastid, as well as the Filosa with the only genus Paulinella, a descendant of a more recent primary endosymbiosis with a Synechococcus-like cyanobacterium (see Sect. 4.4, Figs. 1.1 + 2.7) that has lost the feeding pseudopods (Keeling 2004). The host leading to the Chlorarachniophytes was proven to be a cercozoan protist (Rogers et al. 2007). The amoeboid cells are naked and do not develop a cell wall, as it could hinder the activity of the pseudopodia. Instead, the cells often form net-like pseudopodia (reticulopoda) that can have extrusomes in some cases (Fig. 5.50).

5.3.2 History of Research The first alga of the Chlorarachniophytes, Chlorarachnion reptans, was described by Geitler (1930) as a heterokont amoeba. Even in the year 1971, it was the only species known in the family Chlorarachniaceae of the order Rhizochloridales within the class Xhanthophyceae (e.g. Fott 1971). The rising awareness that the origin of plants and plant-like organisms was achieved by primary and secondary endosymbioses let to a new interest and subsequently a new view on this peculiar group of algae. Promoted by the

c c 5 μm

Fig. 5.50 Reticulopods (arrows) forming a network of several amoeboid cells of Gymnochlora sp. Note the bright green color of the chloroplasts (C) due to chlorophyll a + b-. Courtesy of David J. Patterson

emerging new molecular phylogenetic tools, many studies dealt with not only new species and genera but also ultrastructure and life cycle of the Chlorarachniophytes, unraveling more and more the biology of this small (by species number) group of algae. Nevertheless, there are still a number of questions to answer, for example the details of their life cycles.

5.3.3 Morphology and Developmental Stages 5.3.3.1 Pseudopods The most distinctive character of the amoeboid stage are the pseudopodia radiating from the cell body, allowing movement and interconnection of several cells forming a reticulate network (Figs. 5.54, a, h). Pseudopodia can be as long as or even longer than the cell and allow phagotrophy. Pseudopods are a key structure in amoeba, used for feeding and locomotion. When they build networks of branching filaments with other cells, they are often called reticulopods (Keeling 2017). Pseudopods are mainly built of actin filaments and microtubules. The protrusion of the cell is usually driven by actin polymerization, promoting the cell membrane to the direction of movement, followed by sliding of the cytoplasm into the projection. Thus, the cell is pushed forward and can surround its prey organism with pseudopods for phagocytosis. The process of movement is supported and maybe even exclusively maintained by microtubules (Dietz and Schnetter 1996). Some species, for example from the genus Lotharella, also develop pseudopods in the coccoid stage (Hirakawa et al. 2011).

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5.3.3.2 Chloroplasts and Nucleomorph Chlorarachniophyceae contain chloroplasts that is surrounded by four membranes and still have retained a relict nucleus (nucleomorph) from the green alga endosymbiont (Figs. 2.6 + 2.7; see Sect. 2.3.1). The number of chloroplasts per cell varies from 1 to several, each associated with a nucleomorph (a feature they share with the Cryptista; see Sect. 5.6). Chloroplasts are bright green (Figs. 5.50, 5.51) and contain chlorophyll a + b. The thylakoids form stacks of two to six layers. The chloroplasts are mostly bilobed and have a central pyrenoid that is often capped by a single-membraned vesicle containing the reserve carbohydrate in the form of beta-1,3-glucans (McFadden et al. 1997).. Presumably, chlorarachniophycean cells arose, when an amoeba host cell absorbed a green alga via secondary endosymbiosis (see Sect. 2.3.1), which also explains the existence of four membranes, an inner and an outer pair. The inner two membranes are the chloroplast membrane of the green alga-symbiont, whereas the third membrane stems from the cell membrane of the green algae. The room between the second and third membranes (the inner and outer pairs of chloroplast membranes) corresponds to the former cytoplasma, sometimes called periplastid space that is widened around the base of the pyrenoids and contains the nucleomorph, a vestigial nucleus of the green alga (Fig. 5.51). The outermost membrane originates from the food vacuole of the host cell (Ishida et al. 2007). The five plastid genomes sequenced so far are typically circular and highly conserved genomes that in comparison to chloroplast genomes are relatively small and range from 67.4 to 72.6 kb. Also, gene content and order are more or less identical in all 5 genomes. Of the thirteen plastid genes that are usually present in chlorophyte plastid genomes, but lacking in the chlorarachniophycean plastid genomes, five have been identified in the nuclear genome, proposing a transfer from the chloroplast genome into the host genome during endosymbiosis, while others might have been lost. The high degree of conservation suggests that the change in genes already happened in the common ancestor of chlorarachniophytes (Archibald 2007). The nucleomorph presents a very rare relict of the nucleus of the endosymbiont (see Sect. 2.3.1). As all other descendants of secondary endosymbioses except for the Cryptista, and latterly few dinoflagellates (Sarai et al. 2020), have completely lost the endosymbiotic nucleus, it is thought that chlorarachniophytes’ chloroplasts represent an intermediate phase between an endosymbiotic alga and a semi-autonomic chloroplast (Archibald 2007; Archibald and Lane 2009). The nucleomorph is quite small, contains only three very small lineal chromosomes with about 300 and in one case more than 1,000 nucleotides, and is itself surrounded by two membranes (McFadden et al. 1994a, b; Ishida et al. 2011). Nucleomorph genomes are considered as the smallest nuclear

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Fig. 5.51 a Ultrastructure of Bigelowiella natans. C = pyrenoid cap (ß 1, 3 glucan); FR = flagellar root; MT = mitochondrion; NM = nucleomorph; PL = plastid; PY = pyrenoid. From Hopkins et al. (2012), Genome Biology and Evolution. b Scheme of cell structure and biology of B. natans, according to Hopkins et al. (2012) and Curtis et al. (2012). c B. natans, cell with two chloroplasts and pyrenoid cap (ring-like structure). Courtesy of David J. Patterson

genomes described. The chromosomes each bear an rRNA operon consisting of 18S, 5.8S, and 28S rDNA in subtelomeric regions linked to telomere repeats. The genome is compactly ordered with a high density of genes and only small intergenic spacers. The genes contain many very small spliceosomal introns (Hirakawa 2017). About 200 of the 300 function-predicted genes of the few genomes that are sequenced are highly conserved and encode proteins

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occupied with housekeeping. The loss of genes during endosymbiosis is compensated for by nuclear genes. Looking at plastids with four chloroplast membranes and four genomes, protein transport might get very tricky. In addition to the housekeeping genes, 17 plastid-associated genes that are involved in protein transport are encoded in all nucleomorph genomes sequenced so far (Suzuki et al. 2016; Tanifuji et al. 2016). Other nucleomorph-encoded proteins are nucleomorph-specific and cannot be matched with any known proteins. They are called ORFans (Suzuki et al. 2016).

5.3.3.3 Pyrenoids and Pyrenoid Caps Pyrenoids are microcompartments of the chloroplast with densely arranged molecules of the photosynthetic enzyme RuBisCO (ribulose-1,5-bisphosphate carboxylase/ oxygenase). Its function is to take part in the CO2-concentrating mechanism (CCM) and to provide a CO2-rich environment for RuBisCO (Raven 2010) to enhance the efficacy of photosynthesis. They can be of different structures with deep or shallow slits and are close to the nucleomorph. They are usually capped by an accumulation of the main storage carbohydrate b-1,3-Glucan (Fig. 5.51). 5.3.3.4 Developmental Stages A number of species display any combination of one or more stages in the life cycle (Keeling 2017) and the type of the main vegetative stage is, beside pyrenoid formation and nucleomorph location, used for taxa distinction. Probably, the ancestor of all chlorarachniophytes exhibited all of the three cell types of the life cycle. In the course of species divergence, some taxa lost one or two stages, explaining the diversity of vegetative stages such as amoeboid, flagellate, and coccoid. Interestingly, it has been assumed that habitats play an important role in the evolution of live cycles. Amoeboid forms are usually found, where they can attach to a surface like sand beaches or coral reefs, while no flagellated forms exist. Planktonic species usually do not present amoeboid forms, which makes sense, since attaching and gliding is not an opportunity. Planktonic cells sampled are rather flagellated (Hirakawa et al. 2011; Ota et al. 2005). Amoeboid phase Amoeboid cells have filose pseudopodia (Figs. 5.54a, b, e, h) and are more or less isodiametric. Amoeboid cells found in benthic environments are angular. Cells without filopodia range in size from 8 to 20 lm. In the genera Chlorarachnion and Lotharella, several filipodia radiate from each cell and form a network of reticulopodia (Hibberd and Norris 1984; Dietz et al. 2003; Ota et al. 2005; Ota and Vaulot 2012). In other genera/species, the reticulopodia remain distinct and unconnected (Calderon-Saenz and Schnetter 1989; Ishida et al. 1996, 2000; Ota et al. 2007b). The amoeboid cells are

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phagotrophic. Pseudopodia take up motile and nonmotile eukaryotes and prokaryotes forming sometimes very large ingestion vesicles. Whether or not amoeboid cells may divide or give rise directly to zoospores or coccoid cells is species-specific. Coccoid phase Algae of the coccoid (coccal) organization type (Fig. 5.54c, d, f) can also be found in benthic environments. Their cells are spherical and reach 5–15 lm in diameter. They have a firm cell wall of variable thickness that is composed of multiple layers (Ishida and Hara 1994; Ota et al. 2005). The chloroplasts are more irregularly shaped than in the amoeboid stage. Depending on the species, cells of the coccoid stage can either divide or give rise directly to the amoeboid stage or to zoospores. Zoospore phase Flagellated cells bear a single flagellum (Figs. 5.51; 5.54) wrapping around the cell while swimming. The planktonic zoospores are morphologically variable with pyriform, ellipsoid, or ovoid cells. Cell size ranges from 4 to 24 lm in length and 3–7 lm in width. The flagellum of the swimming zoospore runs like a rotating belt around the cell body, operating in a concavity. Species specifically, the zoospore can become temporarily amoeboid, forming one or more pseudopodia. They either can divide or may directly give rise to the amoeboid or coccoid stages (Keeling 2017).

5.3.4 Reproduction and Life Cycle Reproduction has been observed in a number of species. The cell division of Bigelowiella natans was observed in detail; the order of division of the cell compartments was pyrenoid, nucleomorph, chloroplast, and finally the nucleus (Fig. 5.52). Except for the collapse of the nucleus envelope with remaining fragments, mitosis is normal (Moestrup and Sengco 2001). The nucleomorph, however, divides amitotically and no chromosomal condensation or microtubules have been witnessed (Ludwig and Gibbs 1989). Although gametes and sexual reproduction have been observed in zoospores of Chlorarachnion reptans and amoeba of Cryptochlora perforans (Grell 1990; Calderon-Saenz and Schnetter 1989), sexual reproduction was poorly understood. So far, the ploidy status is only known for the species Bigelowiella natans: while the nucleus is haploid, the nucleomorph status is diploid (e.g. Hirakawa and Ishida 2014). For almost all of the other species and morphological states, the ploidy status is unknown. More detailed information can be found in the reviews of Ishida et al. (2007) and Keeling (2017).

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McFadden 1999; Ishida et al. 1999). Besides the result that the chlorarachniophytes are monophyletic, the following relationships were elucidated (Fig. 5.53):

amoeboid state

• the genera Lotharella and/or Gymnochlora are at the base of the chlorarachniophyte lineage, • the genus Chlorarachnion is monophyletic, and • flagellated strains form a monophyletic clade. R?

R?

ploidy ?

in Bigelowiella natans n!

coccoid state mitosis

Syngamy ? K?

zoospore state (flagellate) mitosis

Fig. 5.52 Life cycle of Chlorarachniophytes. Amoeboid, coccoid, and zoospore states alternate, but not all genera/species express all states. Each state performs vegetative division via mitosis. The ploidy state known for Bigelowiella natans zoospores is known as haploid (Hirakawa and Ishida 2014). Modified from Keeling (2017)

However, there are still a number of ambiguities, for example it remains unclear if the genus Lotharella is monophyletic. The phylogenetic position of Cryptochlora perforans is unclear as well, because neither molecular nor ultrastructural data are available for this genus. The genus Partenskyella was described in 2009 only (Ota et al. 2009) and is characterized by the absence of a pyrenoid in all developmental stages. According to Keeling (2017), the following classification is suggested (Table 5.3; with information from Hirakawa et al. (2011); Hirakawa (2017), species numbers from Guiry and Guiry (2021). Examples of chlorarachniophytes are shown in Fig. 5.54. Table 5.4 Current classification of the Chlorarachniophyta according to Keeling (2017) Class Chlorarachniophyceae, 8 genera and 15 species (species numbers in brackets)

5.3.5 Phylogeny and Systematics Phylogenetic studies, using nuclear- and nucleomorphencoded SSU rRNA genes, revealed at least partly the relationships among chlorarachniophytes (Gilson and

Amorphochlora (1)

Cryptochlora (1)

Norrisiella (1)

Bigelowiella (2)

Gymnochlora (2)

Partenskyella (1)

Chlorarachnion (2)

Lotharella (5)

Haptophyta Plastid ultrastructure Pyrenoid and nucleomorph position Flagellate Coccoid Amoeboid Organizational level (life cycle)

+

+/-

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+

+

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+

+

+

Partenskyella

+

+

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Lotharella

+

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Amphochlora

+

+

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Gymnochlora

-

-

+

pyrenoid nucleomorph

Chlorarachniophyta

Bigelowiella

Fig. 5.53 Phylogeny, organizational level, and ultrastructural characteristics. Phylogenetic tree based on nuclear 18S rDNA sequences. Red lines in the tress indicate loss of the amoeboid state; blue lines indicate loss of the flagellate and coccoid states. *According to Ota et al. (2009). Modified from Hirakawa (2017)

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a

b

5 µm

5 µm

c

d

10 µm

c e

py

f

5 µm g

cw

fp py fp

fp 10 µm

1 0 µm

10 µm

h

Fig. 5.54 Species of Chlorarachniophytes. a Chlorarachnion sp. cell with pseudopodia; Gymnochlora sp. cell with reticulopods; figures a and b courtesy of David J. Patterson. c Norrisiella sphaerica, Baja California, from Ota et al. (2007a) with permission of Springer Nature. d Lotharella vacuolata, 2012 © Shuhei Ota and Masahiro Suzuki 2010. e L. globosa, from an artificial coral reef, strain LEX01; fp = filipodia; py = pyrenoid. f L. globosa, binary and quaternary cell divisions, arrowhead indicates parental cell wall. g L. globosa, an amoeboid cell extruding a filopodium out of a cell wall pore (cw); note the pyrenoid. Figures e, f, g from Hirakawa et al. (2011), PLoS One (open access). h Unknown chlorarachniophyte from Roscoff, France, probably amoeboid stage of Gymnochlora, auto-fluorescence image, red structures are chloroplasts. From Phycokeys (Baker et al. 2012)

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Euglenids—(Excavates, Discoba, Euglenozoa, and Euglenida)

Angelika Preisfeld

5.4.1 Short Introduction—what Are Euglenids? Why Are They Called Augentierchen? Everybody knows Euglena gracilis, the “Augentierchen” (Fig. 5.55). But why is it called “Tierchen” in German, which means small animal? Does it have eyes (“Augen”) and can it eat the same way that animals do? The light micrograph shows that neither of these two assumptions is true. It does not have eyes—the red dot is an assemblage of dense lipid bodies—and it cannot be classified as an animal, because it is unicellular. In contrast to unicellular organisms,

animals are always built of many cells. The image of the light micrograph shows that Euglena gracilis possesses chloroplasts which it uses to perform photosynthesis. So, is it a plant? It certainly seems to show more characteristics which would allow for classifying it as a plant, but as a unicell, it does not fit into the definition of plants, either. Therefore, the taxonomic position of Euglena and other euglenids varies, depending either on the perspective of the researcher or the observed taxa (see Box 5.1). The euglenids are indeed a rather diverse and ancient assemblage of flagellated eukaryotes that occur in marine, freshwater, and soil habitats. Members of euglenids can subsist either heterotrophically by absorbing food or autotrophically by performing photosynthesis. Hence, they show characters that have been considered plant- or animal-like, which resulted in the ambiregnal status of the taxon.

Box 5.1: Overview on biological classification—A problem with two codes The aim of biological classification (taxonomy) is to promote the naming and organizing of the diversity of life into taxa, whose members share important properties and to bestow generally accepted names to identified taxa. It was Aristoteles who began dividing organisms into two kingdoms, plants and animals. Later on, Carl Linné invented binominal nomenclature, which means that each organism owns two names, the genus and the species. There are some exceptions to the strategy to file all eukaryotes either under the International Code of Nomenclature for algae, fungi, and plants (Turland 2018) or under the International Zoological Code of Nomenclature, where all animals are listed (ICZN 2012). And the euglenids are part of those exemptions, as they contain an ambivalent or ambiregnal status under both codes due to plant- and animal-like features (Patterson and Larsen 1992). Thus, phototrophic euglenids are considered as algae and listed under the International Code of Nomenclature for algae, fungi and plants (Turland 2018) as Euglenophyta PASCHER 1932 or Euglenophyceae SCHOENICHEN 1925 emend. Marin and Melkonian 2003 [Euglenea Butschli 1884, emend. Busse and Preisfeld 2002a, b] or, more in common language, euglenophytes (Huber-Pestalozzi 1955; Pringsheim 1936; Leedale 1967; Larsen and Patterson 1991). Algae, which are defined as “oxygenic photosynthesizers other than embryophyte land plants” (Cavalier-Smith 2016) exclude most of the heterotrophic forms. These are classified under the International Zoological Code of Nomenclature, where all animals are listed (ICZN 2012) by the name of Euglenida (BÜTSCHLI 1884) emend. SIMPSON 1997. Some taxa are described under both codes, in some cases even under different names or under names, which exist in both codes, but mean different organisms. Pseudoperanema trichophorum Christen 1962, for example, is the botanical designation for the euglenid zoologically known as Peranema trichophorum (Ehrenberg 1838) Stein 1878. Peranema, from the view of the ICBN again, represents a fern genus.

5.4.1.1 Ambiregnal Status One can easily understand that the status of an ambiregnal taxon can be quite confusing (see Box 5.1). Anyway, there are intelligible explanations for consideration under both codes, most comprehensible the nutritional modes (see Sect. 5.4.5), which diversified during the evolution of euglenids. As single-celled euglenids are classified into a group of protists. Protists comprise a large paraphyletic assemblage of mainly unicellular eukaryotes without a recent common ancestor (see Chap. 1) and are present at the

base of each supergroup of eukaryotes (Caron et al. 2017). A revision to the classification, nomenclature, and diversity of eukaryotic supergroups was proposed by Adl et al. (2005, 2012) and revised in Adl et al. (2019). It includes all eukaryotic diversity subsumed under a single code, without ambivalent rank names, and uses informal names, where positions or support are disputed. The classification system used here is based on Adl et al. (2019) with minor exceptions due to ongoing research.

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* ***

*** ***

***

*** Fig. 5.55 Light micrograph of Euglena gracilis with prominent eyespots (S, stigma), chloroplasts (C), some capped by paramylon (P), flagella (F), reservoir (R), and nucleus (N). Bar 10 µm

5.4.2 Taxonomic Classification

****

Taxonomy and classification for euglenids are at least heterogeneous, working with descriptions under one and/or the other code. Here, a hierarchical system without formal ranks and affiliation to a certain code is applied according to Adl et al. (2019) with only slight modifications to facilitate the understanding of possible relationships between taxa. This classification cannot include all phylogenetic observations, due to the fact that only a biased taxon sampling is available. A schematic phylogenetic tree shows the euglenid lineages discussed in the present research (see Sect. 5.4.7, Fig. 5.73).

****

* **

Euglenozoa Cavalier-Smith (1981), emend. Simpson (1997) Euglenida Butschli (1884), emend. Simpson (1997) (continued)

**** ***** *****

Euglenozoa Cavalier-Smith (1981), emend. Simpson (1997) Symbiontida Yubuki et al. (2009) (Note: Still highly controversial, may be placed outside of euglenids! See Sect. 5.4.7). Cells with epibacteria Petalomonadida Cavalier-Smith (1993) Heteronematina Leedale (1967) (Note: Highly controversial, since this is a paraphyletic assemblage only shown to provide a certain amount of order. Petalomonadida have been moved as a monophyletic taxon to the base of the tree, according to Lax and Simpson (2020 a) and Lax et al. (2021). No valid data could justify the erection of more clades at this time. Anisonema, Atraktomonas, Biundula, Calycimonas, Decastava, Dolium, Dinema, Dylakosoma, Entosiphon, Heteronema, Jenningsia, Keelungia, Lentomonas, Neometanema, Pentamonas, Peranema, Peranemopsis, Ploeotia, Scytomonas, Serpenomonas, Teloprocta, Tropidoscyphus, and Urceolus. Phagotrophic euglenids Aphagea Cavalier-Smith 1993, emend. Busse and Preisfeld 2002a, b, Astasia, Distigma, Gyropaigne, Menoidium, Parmidium, and Rhabdomonas. Primary osmotrophic euglenids Euglenophyceae Schoenichen (1925), emend. Marin and Melkonian (2003) [Euglenea Butschli (1884), emend. Busse and Preisfeld (2002a, b)]. Phototrophic euglenoids including secondary osmotrophs (traditional botanical classification used for phototropic euglenids) Rapaza Yamaguchi et al. (2012), Rapaza (mixotroph) Eutreptiales Leedale (1967), emend. Marin and Melkonian (2003), Eutreptia, Eutreptiella Euglenales Leedale (1967), emend. Marin and Melkonian (2003) (= Euglenea) Phacaceae Kim et al. (2010), Lepocinclis, Phacus, and Discoplastis Euglenaceae Dujardin (1841), emend. Kim et al. (2010). e.g. Euglena, Colacium, and Trachelomonas

5.4.3 Origin and Fossil Record Since most euglenids do not own structures allowing for fossilization, only very little is known about the fossil record. Only cells building a lorica, such as Trachelomonas,

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Monomorphina, and Phacus or such cells equipped with very thick pellicle strips are able to leave fossilized traces. Strother et al. (2020) investigated samples of two Moyeria holotypes (Gray and Boucot 1989) from the Upper Silurian (about 415 mya) of the Hagshaw Hills inlier by light and transmission electron microscopy. They observed a wall ultrastructure that fits well into the range of euglenid pellicle diversity (Leander and Farmer 2000, 2001). And even the light microscopical image (Fig. 5.56) demonstrates that the striate surface structure of arches and U-shaped interconnecting segments is highly reminiscent of euglenid pellicles like in the genus Monomorphina. Consequently, they moved Moyeria from the Incertae sedis group Acritarcha, into the euglenids.

5.4.4 History of Research Antoni van Leeuwenhoek (1632–1723) was a well-regarded microscopist, who was able to manufacture the finest lenses to increase the microscope’s resolution. He was the first to recognize the microbial world and to inform the Royal Society about small animals in a letter to the Philosophical Transactions of the Royal Society (1676). The letter was in parts translated imperfectly by the editor (and non-biologist) Henry Oldenburg (1677), who was by the way one of the first to introduce peer-reviewing. In his first report to him, Antony van Leeuwenhoek described “animalcula”, translated as small animals, which he observed in “rain, well, sea and snow water and also in water wherein pepper had lain infused.” He continued, “… when these animalcula or living

Fig. 5.56 Comparison of fossil Moyeria and Monomorphina. a Transmitted white light micrographs of Moyeria from the Upper Silurian. Visible are the pellicular strips from top to bottom of the cell (arrowheads). Image by Peter Strother, reproduced with permission. b Monomorphina pseudopyrum, courtesy of Richard E. Triemer, Woongghi Shin, Jong Im Kim, reproduced with permission. Scale bar = 10 µm

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atoms did move, they put forth two little horns, continually moving themselves.” Since other contemporary scientists were in lack of such fine lenses van Leeuwenhoek used, they did not believe in the microbial world and the little animals he described and a long time passed, until several researchers joined his view. 300 years later, Dobell identified several organisms by the descriptions Leeuwenhoek bequested, like a phototrophic euglenid or the ciliate Vorticella (Dobell 1932, 1958). In the meantime, a green flagellate was described by Mueller (1786), and named Cercaria viridis. It was the well-known euglenid specialist Ehrenberg (1830) who renamed it as Euglena viridis, the type species of the genus Euglena, to which many diverse taxa have been assigned. To bring a certain order into the genus, Dujardin and Perty erected two new genera and transferred the species based on morphological characters to Phacus (Dujardin 1841) or Lepocinclis (Perty 1849). In the following time, a plethora of phototrophic freshwater euglenids was described, for instance Cryptoglena, Colacium, and Trachelomonas by Ehrenberg (1832, 1834), and the two marine genera Eutreptia by Perty (1852) and Eutreptiella by Da Cunha (1913). It was again Ehrenberg, who introduced the osmotroph Distigma (1838) as a euglenid. Subsequently, Dujardin (1841) described the two phagotrophs Ploeotia and Anisonema. This earlier taxonomical work was followed by many other descriptions undertaken by Stein (1878), Klebs (1883), Bütschli (1884), Senn (1900), Lemmermann (1913), Calkins (1933), Huber-Pestalozzi (1955), and Pringsheim (1956). With the advantage of electron microscopy and biochemistry, new information was gathered and published by Leedale (1967) or Buetow (1968). Spurred by the inception of

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molecular phylogeny and the opportunity to use genomic and transcriptomic data, many changes in tree hypotheses and affiliations have occurred. Up to now, more than 1,500 euglenids are described, with phagotrophs still underrepresented (Leander et al. 2017).

5.4.5 General Information and Diversity of Nutrition Modes Euglenids are a rather large and diverse group of unicellular eukaryotes with many individual features. They differ in cell dimensions from less than 10 µm (Petalomonas) to more than 400 µm (Euglena ehrenbergii). Mostly, they are flagellated and motile with only a few exceptions like the sessile Colacium (Fig. 5.58h) that attaches to other Colacium, building small tree-like colonies and/or to the substrate by mucilaginous stalks, which emerge from the anterior end (Huber-Pestalozzi 1955; Leedale 1967). A typical euglenid cell is surrounded by a pellicle, which differs from cell walls known from plants or green algae (Fig. 5.57). They can have chloroplasts or feeding apparatuses or nothing of both, dependent on their nutrition

Fig. 5.57 Cell morphology of Euglena gracilis. a Light micrograph of E. gracilis; b Schematic drawing with most prominent characters. Original drawing Spindler & Preisfeld

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strategy. One or more flagella emerge from an opening in the anterior part, called the canal. Proximally, the canal leads to the reservoir. A vacuole lying close to the reservoir ensures osmotic balance. The nucleus contains visible condensed chromosomes, even in the interphase of the cell cycle, and holds a prominent nucleolus. The carbohydrate storage product paramylon lies distributed in the cytoplasm. Typically, one large “reticulated” mitochondrion shows discoidal cristae.

5.4.5.1 Nutrition—You Are What You Eat To understand the diversity of euglenids, a glance at the variable modalities of nutrition is necessary. As a special euglenid feature, three different forms of feeding developed (Fig. 5.58) that allowed for diversification by inhabiting different ecological niches in four major steps. (i) Molecular and morphological data suggest phagotrophy as the plesiomorphic condition, even though phagotrophs are not monophyletic, but are scattered across a phylogenetic tree. It indicates that the cells are able to engulf bacteria or small eukaryotes by means of a simple (Petalomonas) or rather

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Fig. 5.58 Light micrograph of diversity in phototrophic euglenids. Bactivorous phagotrophs: a Petalomonas minuta, b Entosiphon sulcatum. Eukaryovorous phagotrophs: c Peranema trichophorum, d Dinema platysomum. Primary osmotrophs: e Distigma sp., f Menoidium sp. Phototrophs: g Lepocinclis fusca, h Colacium vesiculosum. Secondary osmotrophs: i Lepocinclis cyclidiopsis, k Euglena quartana. Bar 10 µm

sophisticated (Entosiphon, Ploeotia) feeding apparatus. So-called bacterivorous phagotrophs feed mainly on bacteria, have a rigid form and are relatively small euglenids (Fig. 5.58a–b). So-called eukaryovorous phagotrophs are larger, with a flexible form and feed mainly on small eukaryotes (Leander et al. 2007) (Fig. 5.58c–d). The distinction between eukaryovory and bacterivory is not perfectly accurate, because phagotrophs probably differentiate between small and large prey and not between eukaryote or prokaryote. (ii) At some point, phagotrophs started to take up nutrients in dissolved form by pinocytosis and the feeding apparatus was reduced. This happened once successfully and gave rise to the primary osmotrophs, the Aphagea (Busse and Preisfeld 2002a, Linton et al. 1999; Preisfeld et al. 2000, 2001) (Fig. 5.58e–f). Osmotrophs like Parmidium, Distigma, Menoidium, Rhabdomonas, and Astasia only occur in freshwater habitats, not in marine waters, and probably arose in an environment, where sufficient nutrients could be taken up by pinocytosis, presumably in the reservoir region. (iii) Around 600 million years ago, another milestone was set by a further single successful event, when larger phagotrophs with a flexible pellicle engulfed single-celled green algae of the order Pyramimonadales in a process that is called secondary endosymbiosis (Gibbs 1978, see Chap. 2). The phagotroph euglenid did not digest Pyramimonas or a close relative to it completely (Jackson et al. 2018).

Instead, the chloroplast remained actively performing photosynthesis in the former phagotroph. The secondary endosymbiosis involved complex processes of gene transfer and rearrangement (Gibbs 1978; Leander et al. 2001, 2007) (Fig. 5.58g–h; see Sect. 2.3.1). Traces of the transfer are still visible in the chloroplast genomes of phototrophic euglenids (Dabbagh and Preisfeld 2017; Gockel and Hachtel 2000; Hallick et al. 1993). Phototrophs like Lepocinclis and Euglena are still capable of pinocytosis; they need to take up certain vitamins and other micronutrients from the environment, hence are truly mixotrophic (Bertaux et al. 1991; Leedale 1967). But they lost the ability to eat other cells. However, in many euglenids vestiges of the cytostome are still observable. (iv) A few phototrophs reduced the chloroplasts independently from each other, and became secondary osmotrophs (Fig. 5.58i–k) like Lepocinclis cyclidiopsis or Euglena quartana (Bennett and Triemer 2014; Müllner et al. 2001; Preisfeld et al. 2000; Triemer and Farmer 2007). In some of these, remnants of the chloroplasts and their genome still exist (Hachtel 1998). The model organism Euglena gracilis can survive well in the dark by subsisting on pinocytosis, when sufficient carbon sources and vitamins are in the media. In fact, pyrenoids are often reduced when cells are kept in the dark, emphasizing that osmotrophic nutrition by pinocytotic uptake is happening (Briand and Calvayrac 1980; Kiss et al. 1986). When

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Fig. 5.59 Light micrograph of Euglena gracilis. a grown photoautotrophically in the light with many chloroplasts capped by small paramylon grains and small grains in the cytoplasm; b grown heterotrophically in the dark with glucose supplementation. No chloroplasts are visible, but abundant large paramylon grains. F: flagellum, S: stigma, C: chloroplast, N: nucleus, and P: paramylon. Bar = 10 µm

substituted with high quantities of glucose, the heterotrophically grown Euglena gracilis is densely packed with large paramylon granules, which can account for up to 95% of the cell mass (Fig. 5.59).

5.4.5.2 Cells Eating Other Cells—A Variety of Feeding Apparatuses Evolved in Euglenids The diversity of phagotrophic euglenids and the amazing cellular developments of unicellular organisms are well reflected in multiple forms of feeding apparatuses. They are generally composed of the cytostome and by bundles of microtubules, which usually are part of at least one rod embedded in a cement-like matrix. The architecture spans from short simple pocket- or tube-like formations fortified by microtubules (microtubule-reinforced pocket complex, MTR/pocket) to highly elaborated forms aided by microtubular rods and vanes which ran deeply into the cell (Leander et al. 2007; Triemer and Farmer 1991a, b; Triemer 1997). The simplest and least complex form of an MTR/pocket can be found in petalomonids. Petalomonas and some relatives entail a simple cytostome in the form of an invagination of the pellicle which is merely reinforced by a row of microtubules. It is located adjacent to the reservoir which opens as a canal subapically at the anterior surface, as is typical for euglenids (Fig. 5.60a–d).

In Ploeotia, the feeding apparatus is large and already visible by a light microscope (Fig. 5.60e). It comprises four distinct components: Two rods, a cytostome, a comb, and vanes, which all work together when prey is captured. Most impressive are two supporting rods with an amorphous mass in the center sheathed by cement-like material (Fig. 5.60f). Both rods are connected by a cement-like crosspiece at the anterior end forming the ventral lip. The chute-formed cytostome lies between a comb and the rods and descends into the cell (Fig. 5.60g). The comb is a multilayered structure with three rows of microtubules, surrounded by a cement matrix (Fig. 5.60g, arrowhead). It is connected via fibers to the pellicle and serves as the dorsal lip when prey is engulfed (Belhadri and Brugerolle 1992; Linton and Triemer 1999). The plicate vanes are located between the rods (Fig. 5.60g, h). They probably pull the prey deeper into the cell for digestion in food vacuoles (Linton and Triemer 1999). The feeding apparatus of peranemids is also complex with two rods as shown in Peranema (Fig. 5.61a–d). The rods consist of interlinked microtubules and are embedded in amorphous material (inlay Fig. 5.61b). They separate the reservoir from the cytostome and run in parallel alongside the longitudinal axis (Fig. 5.61c). They are fused together at the anterior end close to the cytostome by a bridge and form a slightly movable plunger to pull the prey into the cell. It has been reported that Peranema also feeds by piercing the

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Fig. 5.60 Light and transmission electron micrographs of feeding apparatuses of Petalomonas cantuscygni (a–d) and Ploeotia costata (e–h). b Longitudinal section of Petalomonas with cytostome and reservoir. c Transverse section of Petalomonas showing the one flagellum and no other structures supporting the feeding apparatus. d Longitudinal section of the MTR/pocket with longitudinally sectioned microtubules. e Light micrograph of Ploeotia with visible feeding apparatus. f Anterior cell region with two flagella (F1, F2) in the reservoir and two rods (arrowheads) with cement-like material embracing the inner vanes. The rods are connected by a crosspiece. g Almost longitudinal section through P. costata with the feeding apparatus running the length of the cell, closely to the reservoir and the comb (arrowhead). h High magnification of the vanes forming the inner part of the feeding apparatus. Cp: crosspiece connecting the rods; Cy: cytostome; FA: feeding apparatus; F: flagella; FV: food vacuole; R: reservoir; M: mitochondria; Ro: rods; V: vanes (c, d, f, g, h courtesy of A. Vollmer)

prey with the rods. This is called myzocytosis (Nisbet 1974; Triemer 1997). The tube-formed feeding apparatus of the ploeotid Entosiphon sulcatum consists of three rods surrounding a propeller-like structure of vanes that are fused together to form a protrusible siphon (Fig. 5.61f, h). It lies adjacent to canal and reservoir. The microtubules of the rods are highly organized in parallel tiered rows that form triangular bundles (Fig. 5.61g). At the anterior end, the siphon is covered by a

movable cap that is withdrawn, when the siphon is extended toward the cell opening (Fig. 5.61h). This way, prey can be captured and “sucked” in by the help of the vanes that probably act as a diaphragm and support the incorporation into food vacuoles (Triemer and Fritz 1987). Highly reduced remnants of feeding apparatuses also exist in autotrophic and osmotrophic euglenids and support the theory of a plesiomorphic phagotrophic condition (Surek and Melkonian 1986; Willey and Wibel 1985).

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Fig. 5.61 Light and transmission electron micrographs of feeding apparatuses of Peranema trichophorum (a–c) and Entosiphon sulcatum (d–h). b Transverse section through P. trichophorum with two flagella (F1, F2) in the reservoir (R) and two rods (Ro) in close proximity. Inlay: Close-up of a rod with interconnected microtubules embedded in transparent matrix. c Longitudinal section through the anterior region with one rod visible left from the reservoir. The striated fiber on the left supports the cytostome movement. e E. sulcatum: Transverse section through a complete cell at midlevel. CV, contractile vacuole; ER, endoplasmic reticulum; FV, food vacuole; G, Golgi apparatus; R, reservoir; S, siphon. f Longitudinal section through a cell depicting the prominent siphon (S) adjacent to the reservoir. g Close-up of the ingestion apparatus (e) with vanes (V) in the center surrounded by three microtubular bundles (MTB). h Longitudinal section of the ingestion apparatus. The central cap (Cp) covers the siphon with rods (Ro) and 4 vanes (v). Electron micrographs courtesy of A. Vollmer

5.4.6 Characters Uniting Euglenids—An Overview of Morphology and Cell Structure 5.4.6.1 Pellicle and Metaboly Surface architecture and cell shape of euglenids are quite unique. The pellicle is of complex structure with integrated cytoskeleton function and covers the cell in a different way known from cell walls of plants and some other algae. It is considered to be the only true synapomorphy all euglenids share. The pellicle developed in varying manifestations during the diversification of species and was furthered by the

necessity to enable full cell function, locomotion, cell shape maintenance, metaboly, and intracellular transport for nutrition (Fig. 5.62). The pellicle (pellicula [lat.] diminutive of pellis, skin) is composed of four components (Fig. 5.63a). (1) The plasma membrane builds the outer envelope of the cell and coats the complete cell and flagella. (2) Slightly laterally overlapping repeating proteinaceous strips underly the plasma membrane as an epiplasmic layer, which is a special part of the cytoplasm, always close to the membrane (Leander et al. 2001, 2007). (3) The strips are supported by a variable number of subjacent microtubules and (4) in close association with

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Fig. 5.62 Varying cell shapes due to metaboly in a Euglena gracilis and b E. mutabilis. Bar: 10 µm

tubular cisternae of the endoplasmic reticulum, where calcium ions are stored (Sommer 1975) (Fig. 5.63a). In Euglena, at least the major plasma protein IP39 is linked to two major epiplasmic cytoskeletal proteins, called articulin p80 and p86 possibly by hydrophobic bonds (Bouck and Ngô 1996; Marrs and Bouck 1992; Rosiere et al. 1990). Huttenlauch and Stick (2003) detected another epiplasmic protein, called epiplasmin. While articulin and probably epiplasmin exist in some other protists, they are absent in multicellular organisms. Traversing centrin fibers, which connect the strips, are thought to be responsible for the upholding of this pattern and also for movement and metaboly that occur when Ca2+ is released from the pellicle endoplasmic reticulum. Probably, centrin fibers also link the microtubule closely to the heel, where the epiplasmic layers overlap, and to the plasma membrane (Cavalier-Smith 2017; Murray 1981; Wiech et al. 1996). The strips cover the complete length of the cell, end close to the reservoir, and are either arranged longitudinally or helically (Fig. 5.63a–e). They are in some species even visible by light microscopy at high magnification (Fig. 5.63b, c). An isolate of the pellicle purified by ultrasonic waves and sucrose gradient centrifugation shows that the cell membrane, the epiplasmic layer, the microtubules, and probably centrin fibers are so closely attached to each other that the procedure including strong centrifugation does not disconnect them (Fig. 5.63f). Movement happens when strips glide relative to one another along the articulation zone, where two epiplasmic strips overlap and microtubules are positioned. Some pellicles are composed of only 4–5 strips; others show up to 120 strips (Dragoş et al. 1997; Leander 2004; Leander et al. 2007; Leander and Farmer 2000). Flexible and rigid pellicles occur in both, heterotrophic and phototrophic forms. Most bactivorous phagotrophs like Petalomonas, Ploeotia, and Entosiphon and several phototrophs and osmotrophs feature a relatively thick epiplasmic layer with rigid fused strips impeding the flexibility to perform euglenid metaboly.

Considering the diversity of pellicle strips in euglenids, it seems possible that low or high number of strips are the result of doubling or reduction events by half during cytokinesis (Brosnan et al. 2005; Esson and Leander 2006, 2008; Yubuki and Leander 2012). While the pellicle of Euglena consists of many unfused helical strips and thus is rather flexible (Fig. 5.64a), the rigid pellicle of Phacus shows fused strips in fewer numbers (Fig. 5.64b). In the presumably basal phagotroph Petalomonas cantuscygni, the epiplasmic layer below the plasma membrane consists of 8 longitudinally arranged strips which are flat, broad, and fused together, leaving the distinction between arches and heels vague. A row of microtubules in uniform distance subtends the pellicle, and the cisternae of endoplasmic reticulum are in close proximity (Fig. 5.64c). Entosiphon sulcatum consists of only a few strips, a thick epiplasmic layer, and is rigid (Fig. 5.64d), but the pattern is altered by repeating strip units composed of 6 sigmoidal strip doublets with alternating deep and flat heels. Also, Ploeotia costata is rigid and possesses 5 longitudinally organized pellicle strip doublets (Fig. 5.64e). Here, the arches are differing alternately. One arch is flat and wide and the next strip’s arch has a deep groove. A prominent keel divides the heel and arch. Contrastingly, Peranema trichophorum has more than 40 helically arranged pellicular strips, which are not fused together and thus enable extreme euglenoid metaboly (Fig. 5.64f). The structure is highly similar to that introduced for Euglena gracilis (Fig. 5.64a); only a prominent keel is missing. Interestingly, the outer surface is smoothly covered with a glycocalyx. The flexibility of many helically, non-fused strips allow eukaryovorous phagotrophs like Peranema to engulf larger prey (Cavalier-Smith 2017; Leander et al. 2017; Leander and Farmer 2001). Although there exists great variation in strip architecture and articulation, most phototrophic and secondary osmotrophic euglenids show a similar ground pattern to Peranema, strengthening the view that some ancestor or close relative to Peranema (“peranemids”) was host to the

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Fig. 5.63 Pellicle construction. a Schematic drawing of the pellicle. b Euglena fusca. c Pellicle stripes in high magnification of E. velata,. d Scanning electron micrograph of E. gracilis showing the flagellum emerging from the canal (C). e Distigma proteus shows a flexible pellicle with numerous strips, which reach into the canal. In the reservoir, two rows of microtubules lie below the membrane. Both flagella (F1, F2) are visible. f Transmission electron micrograph of isolated pellicles of E. gracilis with plasma membrane (P), microtubules (MT), and vesicles of ER still in place. a Original drawing Spindler & Preisfeld

endosymbiosis with a green alga. The loss of ingestion apparatuses that led to primary osmotrophs (Aphagea) probably happened in another closely related phagotroph with a helical pellicle. Nevertheless, not all phototrophs and

osmotrophs present a flexible pellicle. The phototrophs Lepocinclis and Phacus (Fig. 5.64b) and some osmotrophs such as Parmidium or Rhabdospira show rigid pellicles with fused strips (Leander and Farmer 2001; Leedale 1967).

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Mostly, the number of strips and the extent of metaboly correlate positively (Leander et al. 2007; Leedale 1967; Schwartzbach and Shigeoka 2017).

5.4.6.2 Extrusomes—Trichocysts and Mucocysts Produce Extracellular Matrix Below the pellicle, sometimes small membrane-bound structures are visible that exist in variant forms (Fig. 5.65). These extrusomes can function as thick-walled ejectile organelles mainly in bacteriovorous euglenids and are then called trichocysts (Breglia et al. 2010; Farmer 2011; Lee and Simpson 2014a, Simpson 1997). They contain a highly organized material which can be discharged as long needle-shaped structures lined by a lattice framework (Hilenski and Walne 1983; Leander and Farmer 2000; Mignot 1963; Simpson 1997). Function and stimuli are unknown, although a role in defense or predation strategies is plausible. Another kind of extrusomes are mucocysts in mainly phototrophic forms, which contain a water-soluble mucopolysaccharide as extracellular matrix. They are found beneath fine pores in the articulation zones of the pellicle, through which they release mucus, when, for instance, encystation is going to happen (Esson and Leander 2008). In

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Colacium (Fig. 5.58h), they are responsible for large amounts of extracellular material used to compose the stalks, by which the cells are connected or attached to a surface. In Trachelomonas or Strombomonas mucocysts are supposed to build the material for lorica as extracellular matrix (Leedale 1967; Hilenski and Walne 1983). Some phagotrophs such as Peranema show mucilaginous coating of the cell membrane, presumably stemming from mucocysts, too (Fig. 5.64f).

5.4.6.3 Canal and Reservoir Euglenids display an invagination at the anterior end that lies either apically or (mostly) subapically and is partially lined by pellicular strips and microtubules (Fig. 5.66). The invagination resembles a narrow canal that expands into the reservoir (Leedale 1967). The canal serves as a distal opening of the cell to the extracellular environment and merges proximally with the reservoir. Distally, the pellicle structure follows into the canal, supported by a second row of microtubules, until more proximately the cell membrane becomes smooth, but is still sustained by microtubules (Kivic and Wesk 1972). Close by lies the contractile vacuole that discharges into the reservoir as a means of osmoregulation. The pattern of

Fig. 5.64 Transmission electron micrographs emphasizing the diversity of euglenid pellicles. A more or less regular pattern of strips in an “arches and heels” manner is visible in transverse section. a Euglena gracilis with a flexible, helical pellicle. The plateau-shaped arches (A) with a small overhang (Ov) are separated from the heel (H) by a distinct keel. Rows of microtubules (MT) are running perpendicularly to the strips below the epiplasmic layer (EL). They are followed by cisternae of the endoplasmic reticulum (ER). The plasma membrane covers the strips (P). This pattern is modified in different species in regard to flexibility, thickness of the epiplasmic layer, and number and arrangement of microtubules. b Phacus, rigid, helical. c Petalomonas, rigid, longitudinal. d Entosiphon, rigid, longitudinal. e Ploeotia, rigid, longitudinal. f Peranema, flexible, helical. M: mitochondria; Mu: mucus (c–f courtesy of A. Vollmer)

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Fig. 5.65 Transmission electron micrograph of extrusomes in Peranema. a Transverse section of tubular extrusomes. b Almost longitudinal section. Bar: 1 µm (courtesy of A. Vollmer)

Fig. 5.66 Canal and reservoir region. a Scanning electron micrograph of the canal region with emergent flagellum. b Transmission electron micrograph of the canal and reservoir region in longitudinal section. Ca: canal; R: reservoir; F: anterior flagellum (b, courtesy of A. Vollmer)

microtubules underlining the reservoir is here less structured (Kivic and Wesk 1972; Leedale 1967; Leander et al. 2017; Rosati et al. 1991). At the bottom of the reservoir usually two flagella emerge that are continuous with the membrane of the reservoir.

5.4.6.4 Thick Flagella Mark the Euglenids The flagellar apparatus arises from a basal body complex with a ventral and a dorsal basal body connected by a striated fiber and three asymmetrically arranged roots (Leedale 1967). In most phototrophs, osmotrophs, and a few phagotrophs, only one flagellum (dorsal, F1) emerges and the ventral flagellum (F2; see Fig. 5.67) is reduced and remains inside the reservoir (Dawson and Walne 1994). Petalomonas is an exception with just one flagellum (Cann and Pennick 1986). The flagella arise at the base of the reservoir and protrude (one or all) through the canal opening (Fig. 5.66a). Extremely speaking, a flagellum is a long protrusion of the cell with an axoneme structure and is hence surrounded by the cell membrane, albeit mostly with specific structures. Phototrophs and osmotrophs usually live as plankton and are able to swim with one, two, or four emergent flagella; phagotrophs often are attached to a surface or sediment and glide mostly with a leading or trailing flagellum (Fig. 5.67a–f) (e.g. Dawson and Walne 1994; Saito et al. 2003). Sometimes, as with Peranema trichophorum, the trailing flagellum lies within a ventral groove and is closely appressed to the cell.

The pellicle of the groove shows a different pattern with more microtubules and a widened heel. Attached to the flagella membrane are hairlike, non-tubular protein structures, called mastigonemes (Bouck et al. 1978; Deflandre 1934; Melkonian et al. 1992). They appear in two forms as relatively short hairs all around the flagellum and as single rows of longer hairs (Fig. 5.67g, h). It is thought that hairs, the axoneme, and another special feature inside the flagella, the paraxonemal rod, support the movement by gliding on surfaces or swimming (Saito et al. 2003). The inner architecture of all eukaryotic flagella is a microtubule-based cytoskeleton structure known as axoneme (Fig. 5.67g). Together with many protein complexes and motor proteins, it is responsible for the lengthy structure of flagella as well as for the motility. It is composed of 9 doublet microtubules in concentrical form and two central microtubules, also called 9 + 2-structure (Porter and Sale 2000). In non-emergent flagella, the central microtubules may be absent (Anderson et al. 1991). A synapomorphic feature of the Euglenozoa is a cylindrical, paracrystalline protein structure parallel to the axoneme (Fig. 5.67g, h), called paraxonemal rod (PAR, sometimes also referred to as paraxial rod) (Ngô and Bouck 1995). It is on one side attached to the axoneme doublets and on the other to the membrane and probably to the mastigonemes (Bouck et al. 1978; Saito et al. 2003). It might be

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Fig. 5.67 Flagella diversity in euglenids. a One emergent flagellum in Euglena (a) and Trachelomonas (b). Two flagella in Distigma (c), Ploeotia pseudanisonema (d), Notosolenus (e; only one visible), and Eutreptiella (f). g Transverse section of dorsal (F1) and ventral (F2) flagella of Entosiphon showing unequal width due to differing paraxonemal rod structures (PAR). h Negative staining micrograph of an isolated flagellum of Euglena. The membrane was removed by Sarkosyl treatment. Still, the mastigonemes stay attached to the PAR

missing in non-emergent flagella. The PAR in the dorsal flagellum has a whorled lattice-like structure that seems to be rather amorphous in ultrastructural analyses, whereas the PAR of the ventral flagellum is highly structured and paracrystalline with a three-dimensional lattice (Fig. 5.67g). Although PARs exist in other taxa (i.e. dinoflagellates) as well, their biochemical and ultrastructural composition differ widely, so they are not regarded as homologous to the euglenozoan paraxonemal rod (Anderson et al. 1991). It is assumed that the PAR improves cell motility and helps to stiffen the flagellum while gliding (Saito et al. 2003; Talke and Preisfeld 2002; Walne and Dawson 1993). The major PAR proteins are encoded by the two paralogous genes par1 and par2 as a result of a gene duplication prior to the separation of kinetoplastids and euglenids (Talke and Preisfeld 2002).

5.4.6.5 Reproduction and Nucleus Clonal (asexual) reproduction is almost the only mode of reproduction in Euglenida. Only one description reports the fusion of two cells of Scytomonas pusilla by light microscopy, but was never confirmed (Mignot 1962). Euglenids

reproduce through longitudinal binary fission by mitosis and following cytokinesis. Euglenids like many other protists perform a “closed mitosis” with the nucleus envelope persisting throughout the complete process and a characteristic separation pattern of the chromatids (Barsanti and Gualtieri 2020). The mitotic spindle is intranuclear and aided by several subspindles, which contain chromosomal and non-chromosomal microtubules (Boettcher and Barell 2013; Buetow 2011; De Souza and Osmani 2007; Heath 1980; Raikov 1994; Triemer and Farmer 1991a, b). The normally one nucleolus (also called endosome) stays intact and divides after elongation into two daughter nucleoli (Buetow 2011; Zakryś 1986). However, even if the framework for euglenid mitosis seems to be consistent, there is a great variety in detail in different species (Triemer and Farmer 1991a, b). The assumed 42 chromosomes of Euglena gracilis (Dooijes et al. 2000) remain permanently condensed even in interphase, accompanied by an always prominent nucleolus (Fig. 5.69d). The size of the nuclear genome remains still unknown and estimates vary between 500 Mbp (Ebenezer et al. 2019) and 24 Gbp (Ebenezer et al. 2017) in the genus Euglena. Interestingly, in addition to vertically

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inherited genes, many genes and introns of green algae and land plants could be identified, presumably reflecting the acquisition of endosymbiotic transfer of genes from the chloroplast to the genome (Busse and Preisfeld 2003; Ebenezer et al. 2017; Zakrys et al. 2017). Other genes laterally transferred into the genome stem mainly from haptophytes, ochrophytes, and cryptophytes. This is interpreted as a result of a complex succession of transient endosymbiosis events. Additionally, the genome is expanded by genes from the endosymbiotic event that led to mitochondria (Ebenezer et al. 2019). Hence, the nuclear genome of Euglena shows multiple origins of nuclear genes seemingly contributing to the complex mosaic genome. The capacity of Euglena to capture genes to a lesser degree also from various red and green algae during presumably many transient endosymbiotic events was identified in the plastid genome as well and is called the shopping bag hypothesis (Curtis et al. 2012; Dorrell et al. 2017; Ebenezer et al. 2019; Howe et al. 2008; Maruyama et al. 2011).

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5.4.6.6 Euglenid Chloroplasts Evolved by Secondary Endosymbiosis Euglenid chloroplasts resemble those from green algae and land plants as they both share an almost identical pigment set of chlorophyll a and b, beta-carotene, and several xanthophylls (Fig. 5.68). Because of their biochemical characters, green euglenids have been classified within the Chlorophyta for a long time, although several morphological and biochemical differences had been acknowledged (Round 1971). The most distinct characteristic are the three membranes, which surround the chloroplasts (Fig. 5.68e). Gibbs (1978) proposed early on that the chloroplasts in Euglena stem from a secondary endosymbiosis event, during which a eukaryovorous phagotrophic euglenid ingested a green alga but did not digest it, which led to the evolution of Euglenophyceae (see Sect. 2.3.1). With the advantages of molecular and genomic phylogenetics, analyses of the chloroplast genomes revealed that Pyramimonas parkeae is probably the closest extant

Fig. 5.68 Light and transmission electron micrographs of phototrophic euglenids and chloroplasts. a Euglena mutabilis with flagella (F) emerging from the reservoir (R) and the stigma (S) in close proximity, chloroplasts (C), and paramylon grains (P). b Phacus longicauda. c Euglena velata with chloroplasts and pyrenoids (Py). d TEM micrograph of E. gracilis featuring pyrenoids in the chloroplasts, a big nucleus, condensed chromosomes, and a prominent nucleolus (Nu) in the nucleus (N). e High magnification of a chloroplast surrounded by three membranes (CM)

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relative of euglenids’ chloroplasts (Dabbagh and Preisfeld 2017; Hrdà et al. 2012; Schwartzbach and Shigeoka 2017; Turmel et al. 2009; Vanclova et al. 2017). From all that we know about successful endosymbiosis, the incorporation of a cell should result in four membranes surrounding the plastid: 1) the phagosomal membrane of the host, 2) the cell membrane of the green alga (3 and 4), and the two plastid membranes of the green alga. Interestingly, only three membranes can be found in euglenids (Fig. 5.68e). Mostly, a scenario is accepted, where the cell membrane from the ingested green alga (membrane number 2 from outside) was digested in euglenids and the second and third chloroplast membranes are the plastid membranes from the green alga. The first membrane belongs to the phagosome membrane of the host cell, which was the phagotrophic euglenid (Archibald and Keeling 2002; Keeling 2010). Nonetheless, any additional membrane calls for a highly complex transport system for metabolites, proteins, or ions to make cell functions possible. Number, location, and morphology of chloroplasts between genera and even between species are highly diverse. Thylakoids are usually arranged in stacks of three (Vanclová et al. 2017). Chloroplasts can be of discoidal, lobed, spherical, or stellate shape, and with or without pyrenoid, all of which were much used for taxonomic considerations (Ciugulea and Triemer 2010; Vanclova et al. 2017). The pyrenoid is an area inside the chloroplast densely packed with ribulose -1,5- bisphosphate carboxylase oxygenase (RuBisCO), an enzyme necessary for CO2 fixation. RuBisCo is constituted of a small nuclear-encoded subunit and a large chloroplast-encoded subunit, which hints clearly at a complex activity of endosymbiotic gene transfer during the process of secondary endosymbiosis (Ciugulea and Triemer 2010; Schwartzbach and Shigeoka 2017). Chloroplast genomes in euglenids The acquisition of chloroplasts possibly occurred around 600 million years ago (Jackson et al. 2017). Thus, it was a relatively recent event, compared to the evolution of euglenids in general, which are assumed to be among the first eukaryotes to diverge. Some phototrophic euglenids lost the ability to perform photosynthesis independently. At least, the colorless Euglena longa still has a chloroplast genome, even though considerably reduced. Together with Euglena gracilis, it was the first euglenid chloroplast genome completely sequenced (Gockel and Hachtel 2000; Hallick et al. 1993; Hachtel 1998). Astonishingly, the cpGenome of E. gracilis is about 140,000 bp more than a third larger than that of Pyramimonas parkeae, while the number of genes was reduced from about 110 in P. parkeae to around 90 in Euglenales, and to about 86 or 87 genes in Eutreptiales during endosymbiosis. This was mostly because these genes

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were transferred to the nuclear genome, but some genes were reduced completely (Schwartzbach and Shigeoka 2017). The contradiction between the size and number of genes could be unraveled by intensive studies of so far 27 cpGenomes of all major lineages in Euglenophytes (Bennett et al. 2012; Bennett et al. 2014; Bennett and Triemer 2015; Dabbagh and Preisfeld 2017; Dabbagh et al. 2017; Gockel and Hachtel 2000; Hallick et al. 1993; Hrdá et al. 2012; Karnkowska et al. 2018; Kasiborski et al. 2016; Pombert et al. 2012; Wiegert et al. 2012, 2013). Further focus was placed on whether a small or a large cpGenome with high or low number of introns was present, when phototrophic euglenids emerged. Introns (from intragenic region, Gilbert 1978) are non-coding regions within a gene that are removed by splicing from the RNA transcript before translation. Thus, they exist in the DNA of many genes in almost all organisms, but not in the mature RNA. Two types of introns are common in cpGenomes of euglenids, group II and group III introns. Group II introns of euglenids are smaller than group II introns in the plastids of cryptophytes and green plants and usually occur in tRNA or protein-coding genes. Two extreme and unique derivates of group II introns are group III introns and twintrons in euglenid plastid genomes. Group III introns are introns evolutionary related to group II introns and have only two degenerated domains (DI and DVI). Twintrons are introns nested within another intron (Copertino and Hallick 1993; Lambowitz and Zimmerly 2004; Michel et al. 1989; Sheveleva and Hallick 2004). Initially, only cpGenomes of Euglenales have been examined, until two Eutreptiales cpGenomes were analyzed and offered very small genome sizes with only 7 and 27 introns, respectively (Bennett and Triemer 2012; Hrdá et al. 2012, Wiegert). A hypothesis was formed that the invasion of introns started with very low intron numbers and as a consequence small cpGenomes in Eutreptiales, which both increased during diversification (Bennett and Triemer 2015; Hrdá et al. 2012; Thompson et al. 1995; Wiegert et al. 2012). This was firstly corroborated by only one intron in Pyramimonas parkeae, as the alleged closest living relative of the euglenid plastid (Turmel et al. 2009). Later on, it was refuted by analysis of different lineages, which resulted on the one hand in large cpGenomes in Euglenales and Eutreptiales with partially more than 110 introns (E. gracilis) and on the other hand in small cpGenomes with low intron numbers in Monomorphina aenigmatica (Bennett and Triemer 2015; Hallick et al. 1993; Pombert et al. 2012; Wiegert et al. 2013). During evolution, the genomes underwent an intensive gene rearrangement, sometimes amended by duplication of certain areas or repetitive sequences, but they still resemble chlorophyte cpGenomes. Consistent with the shopping-bag theory, the cpGenomes hold also genes of red and brown algae origin, which

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probably invaded the genome during various transient endosymbioses. This is further corroborated by some euglenid pigments that are missed in green algae and were probably adopted from transient cohabitants (Casper-Lindley and Björkman 1998; Laza-Martinez et al. 2019). Euglenid nuclear, plastid, and mitochondria genomes are chimeric in nature and support the view that lateral gene transfer occurred multiple times and is a driving force in evolution (Curtis et al. 2012; Ebenezer et al. 2019; Maruyama et al. 2011; Ponce-Toledo et al. 2018).

5.4.6.7 A Photosensory System Enables the Cell to Respond to Light Changes Green and (some) secondary osmotrophic euglenids possess a photosensory system that is composed of an eyespot (stigma) and the paraflagellar body (Leedale 1967). The eyespot consists of membrane-bound orange-red droplets of b-carotene derivatives and is located independent from chloroplasts closely attached to the reservoir in the cytoplasm (Fig. 5.69a) (Tamaki et a. 2020). This is in contrast to green algae, where eyespots are located within the plastids (Kivic and Vesk 1972). The photoreceptor itself is the paraflagellar body (PFB), recognizable as a delicately swollen region in the emergent flagellum opposite to the eyespot (Fig. 5.69b). Phototactic response happens when the cell reacts to light by alternately exposing and shading the paraflagellar body by the eyespot during swimming in rotating movements. The photosensory apparatus allows for negative or positive phototaxis in order to move to proper light qualities for photosynthesis in the environment (Zakrys et al. 2017).

Fig. 5.69 Light and transmission electron micrograph of the photosensory system. a Red eyespot (stigma) in Trachelomonas. b Photosensory system in Euglena gracilis. Lipid bodies (LB) located at the anterior end of the reservoir (R) shade or expose the paraflagellar body (PFB) within the dorsal flagellum (F1)

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5.4.6.8 An Unusual Storage Polymer—Paramylon Chlorophytes and land plants produce starch, a polymer consisting of unbranched a-1,4-linked glucose monomers forming amylose and branched a-1,4/-1,6-linked glucose as amylopectin. Starch is stored inside chloroplasts (Ball and Morell 2003; Zobel 1988). Since phototrophic euglenids inherited the plastid from green algae during secondary endosymbiosis (Gibbs 1978; Turmel et al. 2009), it would be obvious to assume that green euglenids took over the mode of synthesizing and storing carbohydrates from green algae. But many biochemical and ultrastructural as well as some molecular studies show that this is not the case. All phototrophic and osmotrophic together with most phagotrophic euglenids store paramylon (para [gr.] similar to; amylon [gr.] starch) as reserve carbohydrate, which consists of a chain of b-1,3-glycosidic linked glucose monomers (Bäumer et al. 2001; Gottlieb 1850; Kiss et al. 1987). It is not located inside the chloroplast, although it is often closely located to the pyrenoids. Phylogenetic analyses support the hypothesis of an acquisition of paramylon independent from green algae acquisition during secondary endosymbiosis and also from the capability to perform photosynthesis (Leander et al. 2017; Paerschke et al. 2017). But it might be that some transient symbionts played a role in the emergence of paramylon. In deviation from starch, paramylon is found outside the chloroplasts surrounded by a membrane in the cytoplasm. The membrane of paramylon grains contains at least a majority of the protein machinery for synthesis that is encoded by the nuclear genome (Bäumer et al. 2001; Inui et al. 1992; Kiss et al. 1987). Paramylon has a primary structure of a simple, unbranched glucan chain, which forms triple helices drilled to coiled macrofibrils (Kiss et al. 1987). One can compare the complex structure to a thread spool showing a concentric pattern and radial segmentation (Fig. 5.70). The glucan nanofibrils (in Fig. 5.70a and b, the concentrically arranged “thread” around the “spool”) show possible health effects as dietary fibers (Kuda et al. 2009) and by interaction with immune systems (Barsanti et al. 2011; O’Neill et al. 2015) and are especially in Japan used as a superfood (Nakashima et al. 2018). Purified paramylon is as white as starch (Fig. 5.70d) (Bäumer et al. 2001). The complex structure of paramylon is unique in nature. Two other b-1,3-glucans are described in phaeophytes as laminarin and in diatoms and chrysophytes as chrysolaminarin (syn. leucosin). Unlike paramylon, these polysaccharides branched with b-1,6-linkages are water-soluble and stored in vacuoles, and sometimes other sugars are present, such as mannose (Stone and Clarke 1992). Some haptophytes likewise store a linear, water-insoluble b-1,3-glucan in two large granules, but these are of simpler structure and accumulate within the chloroplast (Kiss and Triemer 1988).

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Fig. 5.70 Light and transmission electron micrographs of paramylon. a Negatively stained paramylon granule isolated from Euglena gracilis. b and c Paramylon grains lie surrounded by a membrane in the cytoplasm of Peranema and Entosiphon, respectively. d Purified crystalline paramylon (left), paramylon after dissolving in sodium hydroxide and neutralization with hydrochloric acid, resulting in puffy grains (right) (b, c courtesy to A. Vollmer)

The grains can be small, large, or dimorphic (presence of two size classes of grains within a cell) and have been used largely as morphological characters to infer generic relationships (Ciugulea and Triemer 2010; Gojdics 1953; Monfils et al. 2011). But paramylon is a rarely considered character in phylogenetic reconstruction in regard to major clades within euglenids. Pärschke et al. (2017) suggested a phylogeny including paramylon as a complex character that resulted in monophyletic emergence of paramylon, although the taxon sampling did not include later findings. They researched the presence of paramylon granules in key phagotrophs and found immunogold labeled paramylon in Peranema trichophorum (Spirocuta) and Entosiphon sulcatum (ploeotiids) and none in Ploeotia costata (ploeotiids) and Petalomonas cantuscygni (Petalomonadida). To assume that paramylon was invented twice would certainly be a rather unparsimonious approach.

5.4.6.9 Mitochondria Are Different in Euglenids Mitochondria are membrane-bound organelles that carry out oxidative phosphorylation and produce most of the ATP the cell needs. The morphology of euglenid mitochondria is

mostly in agreement with general concepts with three exceptions: In E. gracilis and presumably in all other euglenids, one large and mobile reticulated mitochondrion complex exists (Fig. 5.71), also called chondriome (Pellegrini 1980). It is possible that other forms exist in euglenids, conceivably dependent on cellular conditions (Leedale 1967; Roy et al. 2007). The second incongruity concerns the invaginations of the inner membrane, the cristae, which are in Euglenozoa slightly constricted at the base and called discoidal cristae (Zimorsky et al. 2017). These are homologous to those of kinetoplastids and most probable to those of Heterolobosea, too (Leander et al. 2017). Also matching the conditions in kinetoplastids is the existence of mitochondrial DNA as small mini circles (nucleoids) attached to the inner membranes (Hayashi and Ueda 1989). It is known that during mitosis and cytokinesis, the nucleoids are halved and divided into both daughter mitochondria, but it is still unknown how they are distributed (Spencer and Gray 2011). The mitochondrial genome is a challenge to science, since it does not follow typical mtGenome arrangement (Hammond et al. 2020). Additionally, the proteome of the mitochondrion is

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Fig. 5.71 Transmission electron micrographs of single reticulated mitochondria with discoidal cristae of a Euglena gracilis and b E. quartana

with more than 17,000 proteins, the largest investigated so far, but lacks the protein machinery that allows RNA editing and processing. It only hosts 7 protein-coding genes, making it necessary to import many proteins from the cytosol (Dobáková et al. 2015; Hammond et al. 2020; Mokranjac and Neupert 2009). Under aerobic conditions, Euglena gracilis synthesizes ATP in the inner membrane, which is a typical process in mitochondria. Under anaerobic conditions, the facultatively anaerobic mitochondria use acetyl-CoA to produce unconventional wax esters that are of biotechnological interest for biofuel production (Barsanti and Gualtieri 2020; Inui et al. 1982, 2017; Müller et al. 2012; Ogawa et al. 2015; Zimorsky et al. 2017).

5.4.6.10 Ribosomal Operon Euglenozoan ribosomal DNA Euglenozoans, at least kinetoplastids and euglenids, show some unfamiliar characteristics that differ from other eukaryotes regarding nucleotide sequence length, gene copy number, localization, and organization of ribosomal genes. One difference are large insertions within the gene for the small ribosomal unit (SSU). Distigma sennii’s, for example, has a length of more than 4,500 nucleotides, which is more than double the usually found 1,800 base pairs. Interestingly, these insertions are not introns, but still exist in the mature RNA and are folded laterally in the secondary structure to inhibit interaction with the active center of the ribosome (Busse and Preisfeld 2002b). In variance from typical eukaryotes, the rDNA of euglenids is not completely organized in the chromosomes as multiple copies of tandem repeat units of 18S, 5.8S and 28S rDNA and interrupted by spacer regions (Fig. 5.72a). Instead, in Euglena only about four of these copies exist in the chromosomes. Around 800 to 4,000 copies lie extrachromosomally as small rings in the

cytoplasm with a length of about 11,000 bp and a massively fragmented 28S rRNA gene (Fig. 5.72b) (Greenwood et al. 2001; Ravel-Chapuis 1988; Schnare and Gray 1990). This leaves the ribosomal operon of euglenids rather complex and complicated, and it is still unknown, whether these peculiar features are a relic of very ancient eukaryotes, because some trypanosomatids show similar characters albeit in lesser magnitude (Schnare and Gray 1990, 2011; Spencer et al. 1987; Torres-Machorro et al. 2010), or an evolutionary side step, which we still do not understand completely.

5.4.7 Phylogenetic Position—Euglenida Euglenids are treated as one of the three (or four) major clades of the Euglenozoa CAVALIER-SMITH 1981 emend. SIMPSON 1997. They embrace Euglenida, Kinetoplastida HONIGBERG 1963, Diplonemida (CAVALIERSMITH 1993) SIMPSON 1997, and as a heavily argued candidate Symbiontida within or outside of the euglenids (Cavalier-Smith 2016; Lax et al. 2019; Yubuki et al. 2009, 2013; Yubuki and Leander 2018). Symbiontida do not share a pellicle with euglenids, the possibly sole synapomorphic character existing. They live in anaerobic environments enveloped by symbiotic bacteria (Yamaguchi et al. 2012; Breglia et al. 2013). A position outside the euglenids seems likely. Euglenozoa are united by several distinct characters: A closed mitosis with a special spindle apparatus that stays inside the nucleus, two basal bodies (kinetosomes) that give rise to three asymmetrically organized microtubular roots, a flagellar structure with a heteromorphic paraxonemal rod (PAR) paralleling the axoneme, tubular extrusomes (also called trichocysts or mucocysts), and usually one or two flagella emerging from an apical/subapical flagellar pocket (Adl et al. 2012, 2019; Cavalier-Smith 1981; Farmer 2011; Leander 2004; Simpson 1997; Yubuki and Leander 2012;

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Fig. 5.72 Ribosomal DNA operon: a Tandem repeat unit of typical eukaryotic rDNA. ETS: External transcribed spacer; IGS: Intergenic spacer; ITS: Internal transcribed spacers; NTS: non-transcribed spacer. b Extrachromosomal rDNA operon circle of Euglena gracilis (modified after Greenwood et al. 2001)

Yubuki et al. 2013). Molecularly, euglenozoans are combined by trans-splicing during RNA processing together with a capped spliced leader RNA and polycistronic transcription (Campbell et al. 2000). The paraflagellar rod is of tubular or whorled structure in the anterior flagellum and of lattice-like structure in the posterior flagellum (Adl et al. 2019). They differ widely in locomotion, as some forms swim, others glide attached to a surface, and in feeding modes, which range from phagotrophic and osmotrophic over phototrophic to parasitic life forms (Adl et al. 2012; Leander 2004; Yubuki et al. 2013). For the position of Euglenozoa in the tree of life, see Chap. 1 (Fig. 1.1 Opuntia tree of life). The euglenozoans are closely related to Heterolobosea (Page and Blanton 1985), Jacobida (Cavalier-Smith 1993) Adl et al. 2005, and Tsukubamonadida, all together forming the clade Discoba SIMPSON (Adl et al. 2019; Hampel et al. 2009), with members noted as “outgroup” in the schematic drawing of Euglenozoan evolution (Fig. 5.73). The tree shows combined current results and tendencies of phylogenetic analyses and is not the final understanding of evolutionary trends in Euglenozoa and Euglenida. Some positions of taxa are not clear yet and will be altered by future work. This is indicated by dotted lines (Fig. 5.73). Data for phototrophic and osmotrophic euglenids have been sampled extensively, so that relationships are generally well characterized. The mixotrophic species Rapaza viridis represents the basal lineage, while the remaining phototrophic euglenids are divided into Eutreptiales and Euglenales. The Eutreptiales are marine and include only the two genera Eutreptia and Eutreptiella, which have two or four flagella. Euglenales live mostly in freshwater and are split into the two families Euglenaceae and Phacaceae with only one

emergent flagellum; the other is reduced in length and stays in the reservoir. Phacaceae embrace the three monophyletic genera Phacus, Lepocinclis, and Discoplastis, whereas the Euglenaceae are divided into 8 genera (Colacium, Cryptoglena, Euglena, Euglenaformis, Euglenaria, Monomorphina, Strombomonas, and Trachelomonas). Euglena is a rather large and well-researched group and can quite safely be regarded as polyphyletic (Zakrys et al. 2017; Leander et al. 2017). Phylogenetically, phototrophs including secondary osmotrophs, the osmotrophic Aphagea (Busse et al. 2003), and a (yet unknown) number of phagotrophs with a helical pellicle are enclosed in the clade Spirocuta (Cavalier-Smith 2016; Lax et al. 2021) (Fig. 5.73). Phagotrophs, instead, are poorly sampled and no unambiguous data exist for a comprehensive reconstruction of evolutionary pathways. This counts especially for the base of the euglenid tree (Cavalier-Smith 2016; Lax et al. 2019; Adl et al. 2019, Lax et al. 2021) and also pertains to the inclusion or exclusion of Symbiontida (Adl et al. 2012; Breglia et al. 2010; Lax et al. 2019, 2021; Walker et al. 2011; Yubuki et al. 2009; Yubuki et al. 2013). Furthermore, some genera like Ploeotia or Peranema are definitely paraphyletic, and more research is needed to unravel their evolutionary history (Adl et al. 2019; Lax et al. 2021).

5.4.8 Ecology—Where Do We Find Euglenids? Euglenids show no typical habitat, but are true cosmopolitans with widespread abundance in freshwater, marine and brackish water and sediments (Boenigk and Arndt 2002; Walne and Kivic 1990). Especially, phagotrophs are known as important consuments in particularly marine sediments

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Fig. 5.73 Schematic tree of euglenids mainly based on SSU rDNA phylogenies and some transcriptomic data. Phototrophs and secondary osmotrophs are colored green, primary osmotrophic forms blue and phagotrophs red. Due to many uncertainties and incongruent data, most of the phagotrophs show dotted lines. Monophyletic are the Euglenophyceae, Aphagea, Petalomonadida, and Symbiontida. All other taxa are probably paraphyletic and/or researched only in very limited numbers

(Tian et al. 2020; Zhang et al. 2016a, b). Euglenids even succeed in rather extreme habitats, like in furs of Brazilian sloths of the genus Bradypus (Suutari et al. 2010) or under very salty conditions as in the Great Salt Lake, USA (Jones 1944), and at low temperatures on snow (Hoham and Blinn 1979) or endozoic in the damselfly hindgut (Willey 1972). Despite their ubiquitous distribution, they are predominantly found in water–air and water-mud interfaces. Most phototrophic forms are pelagial (Esson and Leander 2008). In ecological research studies, many undescribed environmental species have been discovered, resulting in a big gap between described species and environmental sequences (Forster et al. 2016; Lukesová et al. 2020; Łukomska-Kowalczyk et al. 2016). The remarkable tolerance against pH, light, salinity, or temperature euglenids show (Pereira and Rino 2001) might be due to a protective mucus

layer secreted from mucocysts. Acidophilic Euglena mutabilis is frequently used as bioindicator in freshwater habitats and is able to bind and store iron and other metals (Casiot et al. 2004). Under suboptimal environmental conditions, some euglenids form cysts or palmellae until living conditions improve. In these stages, the flagella are reduced and the cell is enclosed in thick mucus layers (Buetow 1968; Hindák et al. 2000; Rosowski 1977).

5.4.9 Description of Easily Observed Taxa Euglenids can be found easily in all watery environments including mud and soil. They can be observed in green films from the surface of puddles or from green sediments during low tides.

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Fig. 5.74 Choice of easily identifiable euglenids. a Eutreptiella gymnastica, b E. gracilis, c E. mutabilis, d Phacus monilatus, e Lepocinclis acus, f Trachelomonas hispida, g Colacium vesiculosum, h Euglena sanguinea, i Euglena spirogyra, k Menoidium cultellus, l Peranema trichophorum, m Entosiphon sulcatum, and n Ploeotia corrugata. Bar: 10 µm

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The best way to sample phototrophic and osmotrophic species is by drawing a plankton net several times through the euphotic zone or against the stream of watercourses or to scrape green films off substrates. Phagotrophs can be found best in sediments or can be collected by scraping off greenish layers from rocks or decaying plants, where also a wide range of phototrophs can be discovered. A promising method is to put some surface sediment into plastic trays, let it settle, remove most of the liquid, and cover the sediment with lens tissue. On top of this, coverslips are placed and left for about 24 h. Between the coverslips and the sediment gliding and creeping phagotrophs will accumulate and can be observed using a microscope (Preisfeld 2009). Eutreptiella gymnastica belongs to the Eutreptiales and can be found swimming with two flagella of unequal length in marine or brackish habitats displaying vivid metaboly (Fig. 5.74a). Members of the genus Euglena are discovered in almost every freshwater habitat. Euglena gracilis is easily recognized by its metaboly and rapid swimming with one emergent flagellum (Fig. 5.74b). Contrastingly, E. mutabilis swims quite slowly and even performs intensive metaboly at a very slow pace (Fig. 5.74c). Many species of the genus Phacus can be detected in puddles, ponds, and ditches. Phacus monilatus (Fig. 5.74d) shows a short caudal projection and knobbly ornamentation on the pellicle in longitudinal rows. Lepocinclis acus (Fig. 5.74e) has a rigid form with a distinct hyaline posterior spine. The anterior flagellum spins like a lasso while the cell circles constantly. Trachelomonas species are recognizable by a slightly brown lorica, which serves as a rigid envelope with a defined collar through which the flagellum emerges. Loricas are often ornamented with depositions of ferric hydroxide and blunt spines, which become obvious when Trachelomonas hispida is just leaving the lorica (Fig. 5.74f). Colacium vesiculosum can be found as small colonies attached to a substratum (algae, plants, copepods) by mucilaginous stalks at the anterior end of the cell (Fig. 5.74g). Euglena sanguinea is a relatively large phototroph that is able to accumulate enormous portions of the red pigment astaxanthin under certain (stress) conditions. When the population grows very dense, the resulting bloom turns the water “blood red” (Fig. 5.74h). Ehrenberg (1831), who described the species, assumed that the blooms inspired the biblical legend about the time of Moses that water in Egypt was turned into blood. Lepocinclis spirogyroides is another large member that is easily recognized by the spiral ornamentation of the pellicle with brown warts and the slightly twisted cell body (Fig. 5.74i). The osmotroph Menoidium cultellus is flattened and of bean-like shape. It swims by turning around frequently and has large paramylon grains (Fig. 5.74k). Peranema trichophorum is swiftly moving by swimming or gliding, performing thereby profound metaboly. The anterior flagellum is thick and extended forward, while the posterior

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flagellum is very thin and lies almost hidden in a groove (Fig. 5.74l). Entosiphon sulcatum is a small ovoid and somewhat flattened phagotroph. It can be distinguished by the visible feeding apparatus and the anterior flagellum that is directed forward with a quivering tip; the posterior flagellum is tugged behind (Fig. 5.74m). The small, gliding phagotroph Ploeotia corrugata is of rhomboid cell form and has two flagella of unequal length. One rapidly beating anterior flagellum is directed forward. The very long posterior flagellum trails behind (Fig. 5.74n).

5.5

Haptista

Burkhard Büdel, Angelika Preisfeld

5.5.1 General Description The phylum Haptista is formed by the subphyla Haptophyta and the Centroplasthelida, of which only the Haptophyta perform photosynthesis and belong to the algae (Burki et al. 2021; Cavalier-Smith et al. 2015). As Centroplasthelida do not perform photosynthesis and do not have chloroplasts, they do not fall within the definition of algae and, thus, will not be treated in detail here. Haptophyta are unicellular and biflagellate (heterokont) algae characterized by an additional flagellum-like structure that is specialized for the uptake of small food particles and/or substrate attachment, named the haptonema. Their photosynthetic pigments are chlorophyll a + c, as well as b-carotin and xanthophylls. Characteristically, their cell surface is covered by scales of either cellulose (Pavlovophyceae) or, as is the case in more than 300 species of them, have calcified scales and thus are referred to as coccolithophores (Coccolithophyceae). This group of phototrophic organisms occurs almost exclusively in the oceans of the world where they are predominantly planktonic. Only a few live as freshwater algae.

5.5.2 Fossil Record The coccolithophore algae have formed a significant part of the oceanic phytoplankton since the Jurassic period, 201– 145 Mya (De Vargas et al. 2007). Their diversity, however, has apparently peaked already 225 Mya (Triassic, 252–201 Mya). Due to certain biomolecules that are extraordinarily resistant to decay; this can be concluded from the presence of their calcareous nanofossils (Bown 1998). Along the coasts of both sites of the English Channel, the White Cliffs of Dover, England, and the Normandy, France, a massive quantity of calcified cells has been sedimented throughout

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Fig. 5.75 Coccolith algae formed the impressive chalk cliffs on both sides of the English Channel. Cliffs at Etretat, Normandy, France. Courtesy of Evelin Büdel

geological time (Fig. 5.75). Coccolithophores contribute to sequester atmospheric CO2 as limestone and the species Emiliania huxleyi is thought to be the most important species. It is distributed worldwide and frequently forms blooms that cover > 100,000 km2 of the ocean’s surface (Fig. 5.81; Tyrrell and Merico 2004).

5.5.3 Molecular Clock Record Molecular clock studies suggest that the whole group of the Haptophyta diverged from other eukaryotes already in the Proterozoic, 1,200 Mya and the divergence of the two classes Pavlovophyceae and Coccolithophyceae happened about 800 Mya (Medlin et al. 1997). Several other molecular clock studies date the origin of all coccolithophores back to 195 Mya, while the divergence of Coccolithus happened some 64 and of Cruciplacolithus 50 Mya. According to Cuvelier et al. (2010), the picoplankton group Phaeocystales diverged from all other Coccolithophyceae about 480 Mya and then the Prymnesiales diverged from the Coccolithales plus Isochrysidales at around 280 Mya. Interestingly, the warm water Phaeocystis species diverged from the coldwater species about 30 Mya, when the Drake Passage opened, isolating the Antarctic waters. Dispersal to the Arctic across the equator occurred again during a cooling trend around 15 Mya, but a subsequent warming trend isolated the two polar species again (Medlin and Zingone 2007). A detailed description of the molecular clock reconstruction of the

evolutionary history of the haptophytes can be found in Eikrem et al. (2017) and the interested reader is referred to that fundamental work.

5.5.4 Morphology and Ultrastructure The cell shape of unicellular haptophytes ranges from spherical, saddle-shaped to elongated forms. Their two flagella range from iso- to slightly heterokont and insert ventrally, with the posterior flagellum sometimes being reduced and the anterior flagellum with fine, non-tubular hairs (Fig. 5.76). Although the monadal (unicellular with flagella) lifestyle dominates, temporarily colonial forms occur (Fig. 5.79e). The main characteristic feature of the group is their haptonema-named flagella-like appendage that play a role in the uptake of food particles. Cell movement is achieved by two more or less similar flagella without hairs, but are instead covered by submicroscopic scales or knops. Each cell has one or two golden-brown chloroplasts, and many possess organic body scales also used for species identification (Fig. 5.76). The yellow, yellow–brown, and brown chloroplasts contain chlorophyll a and c, b-carotin, and xanthophylls. Storage products inside the cells are chrysolaminarin, oil, and rarely a beta-1,3-glucan, similar to paramylon from euglenids. The chloroplasts are located in plications of the endoplasmatic reticulum (Fig. 5.76). Apart from their flagella, all cells have a thread-like filament named the haptonema (Fig. 5.76). It can be very long,

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Fig. 5.76 Morphology of a representative cell of a coccolithophore alga. Modified from Tsuji and Yoshida (2017), with permission of Elsevier Ltd

haptonema dveloping organic scale coccolith forming vesicle

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in some cases several times the length of the cell body. So far known, the haptonema may be involved in food uptake and/or can also be involved in the substrate attachment of the cell body (for an overview, see Eikrem et al. 2017). The haptonema construction resembles that of a flagellum, but is quite different in fine structure. It is surrounded by the plasmalemma, followed by a fenestrated cylinder of the endoplasmatic reticulum. In the center is a ring composed of 6 or 7 longitudinally running microtubules, which are probably responsible for the coiling of the haptonema upon mechanical stimulation. Coiling has been shown to be strongly reduced by microtubule stabilizers, like taxol (Nomura et al. 2019). The process of food uptake was described in detail by Inouye and Kawachi (1994) and is shown in Fig. 5.77. In several groups of haptophytes, the outer cell surface is covered by scales made from complex carbohydrates and protein. These scales are formed in Golgi vesicles, subsequently transported to the cell wall where they are exuded to the extracellular cell surface forming the coccolith cover. Within the order Coccolithales, the scales are calcified and

termed coccoliths. The function of coccoliths is insufficiently known. Several authors suggest, for example, food protection, buoyancy regulation, light supply, or calcification to support photosynthesis. Some species even have an eyespot (Kawai and Inouye 1989), which is usually only found in Pavlovophyceae, lying on the inner or outer surface of the plastid (Bendiff et al. 2011; Eikrem et al. 2017). Those species whose cells are covered by calcified scales (= coccoltih[s]) are summarized with the term “Coccolithophore”. Their tiny composite exoskeleton is named the coccosphere and is made of multiple coccoliths. Different types of cell surface covering, scale-like structures are known (from Eikrem et al. 2017): (a) holococcoliths (Coccolithaceae, Helicosphaeraceae), non-interlocking structures composed of rhombohedral crystallites of uniform size (ca. 0.1 lm in diameter). Each holococcolith is made up of numerous identical calcite elements in the form of minute rhombohedral or hexagonal prisms.

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haptonema

food particle a flagellum a

a

b

c

d a

Fig. 5.77 Uptake of food particles in Haptolina hirta. a captured food particles at the distal haptonema are transported toward the basis where they aggregate to larger particles (= a); b–c) transport of the particle aggregate toward the haptonema tip; d haptonema deposits the food aggregate at the posterior cell surface where it is ingested. Modified from Inouye and Kawachi (1994)

(b) heterococcoliths (Papposphaeraceae, Prymnesiophyceae) elaborate interlocking structures composed of multiple strongly modified calcite crystals. (c) nannoliths (Ceratolithaceae) differ from both, holo- and heterococcoliths in structure and architecture. (d) unmineralized scales (Prymnesiales, Isochrysidales), composed of microfibrils usually arranged in two layers.

5.5.5 Life Cycle The heteromorphic life cycle, which is common in the Haptophyta, entails two morphologically differing cells between the diploid and haploid stages. The alternation of a haploid stage with a diploid stage has been documented in all orders and many families within the Coccolithophyceae, but has not been found in the Pavlovophyceae (Eikrem et al. 2017). Haptophytes usually reproduce asexually by binary fission (Fig. 5.78): (a) Phaeocystis species show an alternation between a nonmotile planktonic palmelloid phase and motile swarmers (2n). The palmelloid phases, the microflagellates as well as the slightly larger mesoflagellates— the mating cells—are haploid. The macroflagellates are diploid and occur during vegetative cycles. A nonmotile zygote has been observed in some species. (b) Heteromorph diplo-haplontic, isogamy or anisogamy (Fig. 5.78b). Represented by the coccoliths with haploid and diploid generations. The different generations are characterized by a specific cell covering. While

diploid generations bear heterococcoliths, haploid generations, depending on the family/genera, are covered by holococcoliths (Coccolithaceae, Helicosphaeraceae) or nannoliths (Ceratolithaceae). Others (Pleurochrysidaceae and Hymenomonadaceae) can have a non-calcifying benthic stage or a non-calcifying motile stage (Noëlhaerhabdaceae) (Billard and Inouye 2004). (c) The genus Prymnesium develops two distinct cell types with differing scale morphology and cell size. Nonmotile cells may occur and in P. parvum even a silicified cyst occurs (Fig. 5.78a). (d) The genera Chrysotila and Ochrosphaera show an alternation of a nonmotile stage (“Apistonema stage”) with one or more motile forms. The species Chrysotila pseudoroscoffensis produces motile spores without coccoliths after meiosis that give rise to a benthic filamentous phase. It is suggested that the filamentous phase releases isogametes with flagella and haptonema. From the zygote diploid, coccolith-bearing motile cells are released within 24 h. For a more detailed review of life cycles, see Eikrem et al. (2017) and references therein. A number of species belonging to several genera have been discovered that are composed of a combination of holococcoliths and heterococcoliths. In terms of their coccolith cover, these combined cells are interpreted as a transition from the haploid holococcolith stage to the diploid heterococcolith stage (Geisen et al. 2004). There is good evidence that the combined cells are of zygotic nature. Given this stipulation can be documented, the “species pairs” have to be reduced to one single species using the oldest name for the consortium.

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Fig. 5.78 a Heteromorphic diplo-haplontic life cycle with diploid cyst, Prymnesium parvum; b Heteromorphic diplo-haplontic life cycle of Coccolithus pelagicus. Modified from Eikrem et al. (2017). c Coccolithophorid life cycle with alternating diploid heterococcolithophorids and haploid holococcolithophorids. Modified from (Geisen et al. 2002)

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Phylum Haptophyta (cells with haptonema) Class Pavlovophyceae (cells with knob scales), 13 species in 4 genera (Guiry and Guiry 2021; Bendif et al. 2011). Order Pavlovales (e.g. Pavlova, Diacronema, Exanthemachrysis, and Rebecca) Class Coccolithophyceae (= Prymnesiophyceae); cells with organic scales or without; * 710 species (Guiry and Guiry 2021) or 318 species in about 76 genera (Jordan et al. 2004). Order Phaeocystales (Phaeocystis). Order Isochrysidales (e.g. Isochrysis, Emiliania). Order Coccolithales (e.g. Coccolithus, Syracosphaera) species. Order Prymnesiales (e.g. Prymnesium, Chrysochromulina).

Calcification

Fibrillar organic scales, isokont flagella

Within the Haptista, the Haptophyta are grouped together with the Centroplasthelida (centrohelid Heliozoa). This is motivated by the fact that both feed by microtubule-supported filiform appendages (the single haptonema or multiple axopodia), and both groups often cover their cell body with complex mineralized scales (siliceous in centrohelids, usually calcareous but sometimes siliceous in haptophytes) (Cavalier-Smith et al. 2015, see also Burki et al. 2021). The Haptophyta form a phylogenetically well-defined monophyletic group with two classes that can be distinguished: the Pavlovophyceae and the Coccolithophyceae (formerly Prymnesiophyceae). The Pavlovophyceae are considered to be the most primary (plesiomorphic) group among them and are characterized by two unequal flagella, small knobby organic scales, as well as distinctive sterols and pigments (Tsuji and Yoshida 2017). Species of the Coccolithophyceae have two (more or less) equal flagella, are single-celled or colonial, and are distinguished in orders with calcified scales, silicified scales, or no scales (Figs. 5.79 and 5.80). The Centroplasthelida encompass phagotrophic and non-photosynthetic protists with one large spherical or near-spherical centrosome at the center of the cell and numerous radiating axopodia. They are usually covered with siliceous scales or with organic spicules. By applying environmental DNA sequencing, a high haptista diversity has been demonstrated for the marine picoand nanoplankton, and many novel species and lineages might be involved (Eikrem et al. 2017; Burki et al. 2021).

Following Guiry and Guiry (2021), roughly 1290 species exist. However, different species numbers are found in the literature. According to theses authors, the Haptophyta include some 80 extant genera with roughly 330 species in two classes. The class Coccolithophyceae (former Prymnesiophyceae) comprises about 76 genera with 318 species (Jordan et al. 2004) and the Pavlovophyceae have 4 genera with 13 species (Bendif et al. 2011). It is estimated, however, that since they exist, the coccolithophores, have diversified into more than 4000 morphological species, most of which are now extinct (De Vargas et al. 2007). The recent classification of the Haptophyta is presented below (Tsuji and Yoshida 2017; Jordan et al. 2004; Cavalier-Smith et al. 2015):

Coccolithales

Isochrysidales Braarudosphaera

Coccolith producing groups

5.5.6 Phylogeny and Classification

Chrysochromulina s.s. Silicification Colonial, pentacle chitin fibrils

Prymnesiales s.s.

multiple axopodia

Phaeocystales Pavlovophyceae Centroplasthelida

Fig. 5.79 Phylogenetic tree of the Haptista. Modified from Tsuji and Yashida (2017)

Non-calcifying groups

Haptonema

microtubule-supported filiform appendages

Chrysoculter

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a

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b Fig. 5.80 a Emiliana huyxleyi, numerous characteristically ornamented, calcified scales (coccoliths) cover its surface. Courtesy of Björn Rost.

b Palusphaera probertii, broadly elliptical cell with numerous spines that are characterized by basal plates, NE Atlantic Ocean; from Archontikis and Young (2021), open access, Phycologia, Taylor and Francis. c Fossil of Coccolithus pelagicus from the Atlantic Ocean at 48° N and -20°E, from Wikipedia, picture from Richard Lampitt and Jeremy Young, The Natural History Museum London. d Prymnesium parvum, arrow indicates haptonema, Colorado River, Texas (UTEX Strain 2797); from Manning and Claire (2010), open access, Marine Drugs. e Phaeocystis globosa, colony from Marsdiep tidal inlet, the Netherlands; photo credit: Jolanda van Iperen (with permission of the Royal Netherlands Institute for Sea Research). f Pavlova sp., flagellated cells with haptonemata (arrows) (from Wikipedia, picture by CSIRO, http://www.scienceimage.csiro.au/ image/7604)

5.5.7 Eco-Physiology The Haptista are dominant marine primary producers that are responsible for 30–50% of total chlorophyll a biomass in Oceans (Liu et al. 2009). Some haptista form massive blooms and some of them are even toxic (Fig. 5.81). Haptophytes are thus significant players in the global carbonate cycle via their photosynthesis and calcification. Some of their biomolecules are extraordinarily resistant to decay and are thus used by geologists as sedimentary proxies of past climatic conditions (Eikrem et al. 2017).

5.5.7.1 Photosynthesis, Calcification, and CO2— Concentrating Mechanism In seawater, most dissolved inorganic carbon exists as HCO3− under the current atmospheric CO2 level. However, the concentration of the Rubisco substrate CO2 is very low (y15 µM at 15 °C) (e.g. Badger et al.1998). Combined with the slow diffusion rate of dissolved CO2 in water and the slow equilibrium between HCO3− and CO2, limited CO2 acquisition is common in the aquatic environment. To overcome this, also marine phytoplankton has developed a Fig. 5.81 Algal bloom along the coast of Iceland. The turquoise haze in the sea indicates billions of Emiliana huxleyi cells in the phytoplankton. Photo: NASA/GSFC, MODIS Rapd Response

CO2-concentrating mechanism (CCM; see Chap. 3, Box 3.1) to actively take up dissolved organic carbon from the marine environment, providing it efficiently to Rubisco (for a detailed overview, see Tsuji and Yoshida 2017). We already mentioned the calcified coccoliths covering the cell surface of many haptophyte species. Interestingly, there is an obvious link between the calcification of scales and photosynthesis. For Emiliana huxleyi, an intracellular accumulation of dissolved organic carbon (DIC) has been reported, but the photosynthetic affinity for DIC is generally rather low compared to other algae. The observed mismatch between active dissolved organic carbon uptake and the low affinity for dissolved organic carbon of E. huxleyi can be best explained by the inefficiency of the CCM possibly due to a high loss of unfixed CO2 from cells. The CO2 efflux rate is controlled by light, and E. huxleyi can switch the CCM mode from net CO2 uptake to efflux with increasing light intensity (Tchernov et al. 2003). The ability of the CCM to switch between CO2 uptake and efflux is a potential key for acclimation to a broad range of light intensities. The energy consumption by nutrient assimilation is suppressed and the efficient dissipation of excess light energy enables the alga to survive.

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Fig. 5.82 Carbon allocation of Emiliana huxleyi; red letters and arrows indicate carbon flux from photosynthetically fixed carbon; blue letters and arrows indicate calcium flow. Modified from Tsuji and Yoshida (2017)

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