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Comparative morphology and molecular evolution of marine interstitial cercozoans Chantangsi, Chitchai 2009

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  COMPARATIVE MORPHOLOGY AND MOLECULAR EVOLUTION OF MARINE INTERSTITIAL CERCOZOANS   by   CHITCHAI CHANTANGSI   B.Sc., Chulalongkorn University, 2001 M.Sc., The University of Guelph, 2006    A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF   DOCTOR OF PHILOSOPHY   in   THE FACULTY OF GRADUATE STUDIES   (Zoology)    THE UNIVERSITY OF BRITISH COLUMBIA  (Vancouver)   August 2009    © Chitchai Chantangsi, 2009     ii ABSTRACT The Cercozoa is an extremely diverse and poorly understood group of amoeboflagellated microeukaryotes that are united mainly by molecular phylogenetic data; a concrete synapomorphy at the morphological level has yet to be identified for the group.  In order to better understand the biodiversity and evolutionary history of this lineage, I explored several marine benthic habitats in British Columbia, Canada and characterized novel cercozoans with high-resolution light microscopy and electron microscopy.  Comparative ultrastructural studies using scanning and transmission electron microscopy on some of the newly discovered taxa demonstrated several novel features, including putative primary endosymbionts in one lineage (i.e., Auranticordis) and homologous patterns of muciferous bodies beneath the cell surface in another lineage.  I coupled these morphological data with molecular phylogenetic analyses of small subunit (SSU) and large subunit (LSU) rDNA sequences and comparative analyses of polyubiquitin genes.  This approach provided evidence that a concatenation of SSU and LSU rDNA sequences improves the phylogenetic resolution within the Cercozoa and that an insertion of one or two amino acids at the junctions between monomers in the polyubiquitin gene is a universal molecular signature for cercozoans (and foraminiferans).  This study also enabled me to discover and describe eleven new species and five new genera, which underscores how poorly we currently understand the diversity of these marine microeukaryotic predators.  The acquired SSU rDNA sequences from these novel lineages enabled me to provide the cellular identities of several environmental DNA sequence clades previously containing only uncharacterized taxa; these data also demonstrated the effectiveness of using a 600-bp fragment of the SSU rRNA gene for delimiting cercozoan species with limited morphological variation.   iii TABLE OF CONTENTS ABSTRACT ............................................................................................................................ ii TABLE OF CONTENTS ....................................................................................................... iii LIST OF TABLES................................................................................................................ viii LIST OF FIGURES ................................................................................................................ ix ACKNOWLEDGEMENTS.................................................................................................... xi DEDICATION...................................................................................................................... xiii CO-AUTHORSHIP STATEMENT ..................................................................................... xiv CHAPTER 1: INTRODUCTION.............................................................................................1       1.1 Cercozoa and the six eukaryote supergroups ...............................................................1       1.2 General biology of the Cercozoa ..................................................................................1       1.3 Morphological characteristics of the Cercozoa ............................................................2              1.3.1 Microscopic features of the Cercozoa ................................................................2              1.3.2 Ultrastructural features of the Cercozoa .............................................................3       1.4 Biodiversity of the Cercozoa ........................................................................................5       1.5 Classification of the Cercozoa......................................................................................6       1.6 Identification of the Cercozoa ......................................................................................8       1.7 Phylogeny of the Cercozoa...........................................................................................9              1.7.1 The phylogeny of the Cercozoa as inferred from small subunit ribosomal                       DNA sequences ..................................................................................................9              1.7.2 The phylogeny of the Cercozoa and its novel polyubiquitin insertions .............9       1.8 Thesis goals and scope................................................................................................10              1.8.1 Specific aim 1 ...................................................................................................11              1.8.2 Specific aim 2 ...................................................................................................12              1.8.3 Specific aim 3 ...................................................................................................13              1.8.4 Specific aim 4 ...................................................................................................13              1.8.5 Specific aim 5 ...................................................................................................14       1.9 References ..................................................................................................................17   iv CHAPTER 2: AN SSU rDNA BARCODING APPROACH TO THE DIVERSITY OF MARINE INTERSTITIAL CERCOZOANS, INCLUDING DESCRIPTIONS OF FOUR NOVEL GENERA AND NINE NOVEL SPECIES ..................................................24       2.1 Introduction ................................................................................................................24       2.2 Materials and methods................................................................................................26              2.2.1 Sampling and light microscopy (LM) ..............................................................26              2.2.2 DNA extraction and PCR amplification...........................................................27              2.2.3 Sequence alignment ..........................................................................................28              2.2.4 Phylogenetic analyses.......................................................................................29              2.2.5 Genetic distance analyses .................................................................................30              2.2.6 Sequence availability ........................................................................................30       2.3 Results ........................................................................................................................33              2.3.1 Identification of the examined flagellates.........................................................33              2.3.2 The Cryomonadida clade..................................................................................33              2.3.3 The Botuliformidae lineage ..............................................................................36              2.3.4 The Ventrifissuridae clade................................................................................37              2.3.5 The Verrucomonadidae clade ...........................................................................40              2.3.6 The Discomonadidae clade...............................................................................42              2.3.7 DNA barcoding marine benthic cercozoans .....................................................43              2.3.8 Molecular phylogenetic analyses of marine benthic cercozoans......................44       2.4 Discussion...................................................................................................................44              2.4.1 Hidden diversity of marine benthic cercozoans................................................44              2.4.2 The current composition of Protaspis ..............................................................45              2.4.3 Cryothecomonas longipes is more closely related to Protaspis sensu stricto ..46              2.4.4 The benthic Botuliforma benthica gen. et sp. nov. is closely related to the                       planktonic Ebria tripartita................................................................................48              2.4.5 The cellular identities of previously undescribed cercozoans ..........................49              2.4.6 Barcoding marine benthic cercozoans ..............................................................51       2.5 References ..................................................................................................................66 CHAPTER 3: ULTRASTRUCTURE, LIFE CYCLE AND MOLECULAR PHYLOGENETIC POSITION OF A NOVEL MARINE SAND-DWELLING  v CERCOZOAN: CLAUTRIAVIA BIFLAGELLATA N. SP......................................................70       3.1 Introduction ................................................................................................................70       3.2 Materials and methods................................................................................................71              3.2.1 Sample collection .............................................................................................71              3.2.2 Light microscopy (LM) ....................................................................................71              3.2.3 Culture establishment .......................................................................................72              3.2.4 Scanning electron microscopy (SEM)..............................................................72              3.2.5 Transmission electron microscopy (TEM) .......................................................73              3.2.6 DNA extraction and PCR amplification...........................................................74              3.2.7 Sequence alignment ..........................................................................................75              3.2.8 Phylogenetic analyses.......................................................................................75              3.2.9 Sequence availability ........................................................................................76       3.3 Results ........................................................................................................................77              3.3.1 General morphology and life cycle...................................................................77              3.3.2 Main cytoplasmic components .........................................................................78              3.3.3 Molecular phylogenetic position ......................................................................79              3.3.4 Taxonomic descriptions....................................................................................80       3.4 Discussion...................................................................................................................81              3.4.1 Comparison of Clautriavia and Auranticordis .................................................82              3.4.2 Comparison of Clautriavia and Protaspis........................................................83              3.4.3 Emended diagnosis of Clautriavia ...................................................................85       3.5 References ................................................................................................................100 CHAPTER 4: MORPHOLOGY AND MOLECULAR PHYLOGENY OF A MARINE INTERSTITIAL TETRAFLAGELLATE WITH PUTATIVE ENDOSYMBIONTS: AURANTICORDIS QUADRIVERBERIS N. GEN. ET SP. (CERCOZOA) .........................103       4.1 Introduction ..............................................................................................................103       4.2 Materials and methods..............................................................................................105              4.2.1 Sampling and light microscopy (LM) ............................................................105              4.2.2 Scanning electron microscopy (SEM)............................................................105              4.2.3 Transmission electron microscopy (TEM) .....................................................106              4.2.4 DNA extraction and PCR amplification.........................................................106  vi              4.2.5 Sequence alignment and phylogenetic analyses .............................................107              4.2.6 Sequence availability ......................................................................................109       4.3 Results ......................................................................................................................110              4.3.1 General morphology and behaviour ...............................................................110              4.3.2 Main cytoplasmic components .......................................................................111              4.3.3 Molecular phylogenetic position of Auranticordis.........................................112              4.3.4 Taxonomic descriptions..................................................................................113       4.4 Discussion.................................................................................................................115              4.4.1 Comparative morphology ...............................................................................115              4.4.2 Putative primary endosymbionts ....................................................................118       4.5 Conclusion ................................................................................................................121       4.6 References ................................................................................................................136 CHAPTER 5: EVOLUTIONARY RELATIONSHIPS AMONG MARINE CERCOZOANS AS INFERRED FROM COMBINED SSU AND LSU rDNA SEQUENCES AND POLYUBIQUITIN INSERTIONS .....................................................142       5.1 Introduction ..............................................................................................................142       5.2 Materials and methods..............................................................................................144              5.2.1 Source of samples and light microscopy (LM)...............................................144              5.2.2 DNA extraction and PCR amplification.........................................................145              5.2.3 Sequence alignment ........................................................................................146              5.2.4 Phylogenetic analyses.....................................................................................147              5.2.5 Sequence availability ......................................................................................148       5.3 Results and discussion ..............................................................................................148              5.3.1 Preliminary phylogeny of cercozoans as inferred from SSU rDNA                       sequences ........................................................................................................148              5.3.2 Phylogeny of mostly marine cercozoans as inferred from SSU+LSU                       rDNA sequences .............................................................................................149              5.3.3 Patterns of polyubiquitin insertions within the Cercozoa...............................151              5.3.4 Patterns of polyubiquitin insertions within the Rhizaria ................................153       5.4 References ................................................................................................................165 CHAPTER 6: CONCLUSION .............................................................................................169  vii       6.1 Cryptic biodiversity of marine interstitial benthic cercozoans .................................169       6.2 Application of DNA barcoding to species identification of marine interstitial             cercozoans.................................................................................................................169       6.3 Morphostasis within the Cercozoa: a case study of Clautriavia vs. Protaspis ........170       6.4 Ultrastructural characterization and molecular phylogeny of Auranticordis             quadriverberis n. gen. et sp. .....................................................................................172       6.5 Phylogeny of the Cercozoa as inferred from polyubiquitin insertions and             combined SSU and LSU rDNA data ........................................................................173       6.6 Future directions .......................................................................................................174       6.7 References ................................................................................................................176                       viii LIST OF TABLES Table 2.1: Oligonucleotide primers used for amplification and sequencing of SSU rDNA                   in this study...........................................................................................................53 Table 2.2: Triangular matrices showing the number of nucleotide differences and the                   percentage of pairwise sequence divergences between small subunit rDNA                   sequences of 17 cercozoans ..................................................................................54 Table 5.1: Cercozoan protists whose genes were amplified and sequenced in this study...155 Table 5.2: Oligonucleotide primers used for amplification and sequencing in this study. .156                   ix LIST OF FIGURES Figure 1.1: Light micrographs illustrating morphological diversity of marine benthic                     cercozoans and some unknown protists found in this study...............................15 Figure 2.1: Light micrographs of nine novel cercozoans found in this study .......................56 Figure 2.2: Light micrographs of the marine benthic cercozoans examined in this study. ...58 Figure 2.3: Diagrammatic line drawings of the nine novel cercozoans found in this study .60 Figure 2.4: Comparison between Bayesian phylogenies inferred from the 1,617-bp full                     length and 583-bp 5'-half barcoding region SSU rDNA sequence alignments                     of 35 cercozoan taxa ...........................................................................................62 Figure 2.5: Bayesian phylogeny deduced from 923 bp of SSU rDNA sequences of 67                     cercozoan taxa ....................................................................................................64 Figure 3.1: Light and scanning electron micrographs of Clautriavia biflagellata n. sp. ......86 Figure 3.2: Light micrographs of Clautriavia biflagellata n. sp. showing different stages                     in the life cycle....................................................................................................88 Figure 3.3: Light and transmission electron micrographs showing general ultrastructural                     features of Clautriavia biflagellata n. sp. during interphase and division .........90 Figure 3.4: Transmission electron micrographs of Clautriavia biflagellata n. sp. showing                     general ultrastructural features of the uninucleated gliding cells and the large                     multinucleated plasmodia. ..................................................................................92 Figure 3.5: High magnification transmission electron micrographs of                     Clautriavia biflagellata n. sp..............................................................................94 Figure 3.6: Illustration showing the main life cycle stages of                     Clautriavia biflagellata n. sp..............................................................................96 Figure 3.7: A Bayesian phylogenetic tree topology inferred from 1,625 bp of SSU rDNA                     sequences from 36 cercozoan taxa .....................................................................98 Figure 4.1: Light micrographs of Auranticordis quadriverberis n. gen. et sp. showing  x                     cell color, main cytoplasmic components, and variation in cell shape.............123 Figure 4.2: Scanning electron micrographs of Auranticordis quadriverberis n. gen. et sp.125 Figure 4.3: Transmission electron micrographs of Auranticordis quadriverberis                     n. gen. et sp.. .....................................................................................................127 Figure 4.4: Transmission electron micrographs of Auranticordis quadriverberis n. gen.                     et sp., showing different cytoplasmic components...........................................129 Figure 4.5: Transmission electron micrographs showing the ultrastructure of putative                     primary endosymbionts in Auranticordis quadriverberis n. gen. et sp. ...........131 Figure 4.6: A schematic line drawing of Auranticordis quadriverberis n. gen. et sp. ........133 Figure 4.7: Maximum likelihood tree inferred from 32 SSU rDNA sequences, 1,571                     unambiguously aligned sites and a GTR+I+G+8 model of nucleotide                     substitutions ......................................................................................................134 Figure 5.1: Light micrographs of the cercozoans examined in this study...........................157 Figure 5.2: Phylogenetic tree inferred from Bayesian analysis of 1,443 bp of SSU rDNA                     sequences from 85 cercozoan taxa ...................................................................159 Figure 5.3: Comparison of phylogenetic trees inferred from Bayesian analysis of                     a 25-taxon alignment consisting of 23 cercozoan taxa and 2 radiozoans.........161 Figure 5.4: Illustration showing the junction between two ubiquitin monomers within the                     polyubiquitin tract of rhizarians and an array of other eukaryotes...................163         xi ACKNOWLEDGEMENTS   This dissertation would not have been possible without the kind contribution of many individuals, and I would like to express my sincerest gratitude to them.   First of all, I would like to express my deepest gratitude to the Cooperative Research Network (CRN), the Royal Thai Government Scholarship for funding me throughout all of my studies in Vancouver, Canada.  My sincere thanks are extended to all the staff at the Office of Educational Affairs, Washington D.C., who kindly took care of me and processed all of my documents.   Second, but most important, I would like to express my heartfelt gratefulness to my advisor, Professor Brian S. Leander, who provided me with the great opportunity of doing research in his lab.  Thank you very much for your kind advice and support throughout my Ph.D. study.  I also wish to express my thanks to all my labmates: Dr. Mona Hoppenrath, Dr. Naoji Yubuki, Dr. Sonja Rueckert, Dr. Aika Yamaguchi, Dr. Rebecca Rundell, Dr. Celeste Leander, Susana Breglia, Heather Esson, Thierry Heger, Chandni Kher, and Sarah Sparmann. Thank you very much Mona for pioneering study of marine cercozoans at UBC and for providing me some literatures; thank you very much Susana for showing me how to do laboratory work when I arrived UBC; thank you very much Naoji for teaching me how to do electron microscopy work; thank you very much Aika, Chandni, Heather, Mrs. Leander, Rebecca, Sarah, Sonja, and Thierry for sharing your samples, keeping me company, helping me with many English corrections, and being good friends.  In addition, I wish to express my thanks to the EM, the Fast, and the Keeling lab people, particularly Dr. Juan Saldarriaga, Dr. Rowena Stern, Dr. Noriko Okamoto, Dr. Aleš Horák, Gillian Gile, Renny Lee, Garnet Martens, and Derrick Horne for sharing laboratory experience and several useful techniques.   Third, I would like to express my sincerest gratitude to my advisory committee, Professor Martin Adamson, Professor Jim Berger, and Professor Patrick Keeling; the University Examiners, Professor Naomi Fast and Professor Sarah Otto; the External Examiner, Professor Jan Pawlowski; and the Chair, Professor Lorne Clarke.  Thank you very much for your time, kind advice, and valuable comments on my dissertation. I am particularly indebted to the staff at the Antarctic Protist Culture Collection (APCC), in particular Dr. David Caron, Dr. Rebecca Gast, and Dawn Moran for kindly providing a culture of Cryothecomonas sp. (strain APCC MC5-1Cryo).  In addition, my  xii sincere gratefulness is extended to Gillian Gile for cultures of Gymnochlora stellata (strain CCMP 2057) and Lotharella vacuolata (strain CCMP 240) and to Thierry Heger for specimens of Placocista sp. I wish to express my deepest gratitude to several additional people for their kind help and suggestions.  Thank you very much Professor Thomas Cavalier-Smith for some valuable discussions.  Thank you very much Professor Patrick Keeling and Professor Naomi Fast for your kind permission to use some of your lab equipment.  Thank you very much Professor Alexander Myl'nikov and Professor Wonje Lee for some valuable discussions and several literatures. Last but not least, I would like to express my thanks to the Departments of Zoology and Botany, University of British Columbia, for providing me with an excellent environment to conduct research.  To the staff members in the Departments, especially Allison Barnes, Judy Heyes, Sarah Inkster, Alice Liou, and Veronica Oxtoby, I give thanks for their kind help with my documents.  Furthermore, I also wish to thank the staff of UBC libraries and Inter-Library Loan who helped me track down and acquire some references on my research.  In addition, I would like to express my deepest appreciation to all of my friends here and in Thailand. Thank you very much for being nice friends and helping me have a memorable time in Vancouver.  Also, I would like to express my heartfelt appreciation to Kuntida Tangthongchaiwiriya for beautifully illustrating all line drawings of cercozoans, which appeared in this dissertation and for her admirable patience as she tirelessly worked through many revisions to have them perfect for me. Finally, I would like to express my heartfelt gratitude to my parents (Pinyo Chantangsi and Somsri Pornpratimakorn) and to my siblings (Pornsawan, Chokchai, and Chanchai Chantangsi) who have always provided me with the highest degree of love and support.  My special thanks are extended to cercozoan protists for a memorable dedication of their life to my Ph.D. study at UBC.  This dissertation is dedicated to my beloved grandmother: the late Zheng Yu Xiang (鄭玉香) who passed away while I was living and studying in Canada   xiii DEDICATION                  To my beloved country, Thailand                    xiv CO-AUTHORSHIP STATEMENT  All chapters were written in collaboration with Dr. Brian Leander.  Both of us conceived and designed the experiments.  I was primarily responsible for performing experiments, analyzing data, writing all manuscripts, and creating all figures.  Dr. Leander offered suggestions for improving the clarity and organization of the manuscripts and figures. In Chapter 4, Heather J. Esson contributed some light micrographs of Auranticordis quadriverberis, read, and approved the final manuscript. In Chapter 5, Dr. Mona Hoppenrath provided DNA sample of Ebria tripartita for polyubiquitin amplification, read, and approved the final manuscript.               1 CHAPTER 1: INTRODUCTION  1.1 Cercozoa and the six eukaryote supergroups Our current understanding of eukaryotic diversity recognizes at least six major clusters: the Amoebozoa, Archaeplastida, Chromalveolata, Excavata, Opisthokonta, and Rhizaria (Adl et al. 2005; Keeling et al. 2005).  Among these six main groupings, the Rhizaria contains various protists with pseudopodia with supporting microtubules (axopodia) that range from being simple, branching, anastomosing, or radiating.  This supergroup was established by Cavalier-Smith (2002) and presently comprises members of three main subclades, the Foraminifera, the Radiozoa (i.e., Polycystinea and Acantharea), and the Cercozoa.  1.2 General biology of the Cercozoa The Cercozoa was first established on the basis of molecular phylogenetic studies of ribosomal RNA (rRNA) gene sequences (Cavalier-Smith 1998a, b; Cavalier-Smith and Chao 2003).  This group houses a myriad of protists having amoeboid, flagellated, and amoeboflagellated forms.  Members within the group inhabit soil, freshwater, and marine habitats, and both planktonic and benthic lineages are known (Bass and Cavalier-Smith 2004; Massana and Pedrós-Alió 2008; Myl'nikov and Karpov 2004; Nikolaev et al. 2004; Park et al. 2008).  Some groups of cercozoans, such as Phytomyxea (e.g., Phagomyxa and Plasmodiophora), are obligate parasites (Archibald and Keeling 2004; Cavalier-Smith and Chao 2003).  Most cercozoans are heterotrophic; however photosynthetic lineages are also known — the Chlorarachnea and Paulinella chromatophora (Hibberd 1990; Kies 1974; Kies and Kremer 1979; McFadden et al. 1997).  Chlorarachniophytes acquired their plastids from green algal prey cells through secondary endosymbiosis, while Paulinella obtained its  2 photosynthetic machinery directly from cyanobacterial prey via primary endosymbiosis (Keeling 2004).  Many cercozoans are bacterivorous and eukaryovorous, thus playing important roles in nutrient cycles by grazing on bacteria, microeukaryotes, and other organic particulates (Myl'nikov and Karpov 2004).  In fact, some cercozoan taxa [e.g., Allantion and Heteromita] are so common in soil that they may represent half of the total protozoan biomass in these environments (Arndt et al. 2000; Ekelund and Patterson 1997).  Although the Cercozoa is cohesive at the molecular level, unifying morphological features for the group have not yet been identified at the microscopic level (Bass and Cavalier-Smith 2004).  1.3 Morphological characteristics of the Cercozoa 1.3.1 Microscopic features of the Cercozoa Members of the Cercozoa show a diverse array of shapes (Figure 1.1), including asymmetrical, bilateral, and radial (Adl et al. 2005; Karpov et al. 2006; Myl'nikov and Karpov 2004; Nikolaev et al. 2004; Polet et al. 2004; Takahashi and Anderson 2002). Cercozoans are, in general, biflagellated and/or amoeboid.  Amoeboid cercozoans usually possess filopodia (i.e., long, thin, and threadlike pseudopodia) [e.g., Euglypha and Placocista] and axopodia (i.e., radiating pseudopodia) [e.g., Clathrulina and Hedriocystis] (Adl et al. 2005); a chlorarachniophyte with lobopodia (i.e., broad fingerlike pseudopodia) has also been described [i.e., Partenskyella glossopodia] (Ota et al. 2009).  Although flagellated cercozoans usually possess two flagella, some lineages have either one or four, such as the uniflagellated chlorarachniophytes and the tetraflagellated Auranticordis quadriverberis and Cholamonas cyrtodiopsidis (Cavalier-Smith and Chao 2003; Chantangsi et al. 2008; Flavin et al. 2000; Myl'nikov and Karpov 2004; Patterson et al. 2002).  Two heterodynamic flagella (i.e., beating with different patterns) with the anterior one directed  3 forward and the other one directed posteriorly is common in several genera of cercozoans, such as Cercomonas, Heteromita, Katabia, and Protaspis (Myl'nikov and Karpov 2004). These flagella facilitate gliding locomotion along benthic habitats and, in a few lineages, swimming in the water column (Chantangsi et al. 2008; Myl'nikov and Karpov 2004). Cell walls, tests, scales, and siliceous skeletons are present in some groups (i.e., Ebriidea, Imbricatea, and Thecofilosea), although naked cells are also known (Hoppenrath and Leander 2006a, b).  An aperture or slit, from which pseudopodia emerge, is found in a wide range of cercozoans, such as Cryothecomonas, Euglypha, and Protaspis (Hoppenrath and Leander 2006a; Kühn et al. 2000).  Pseudopodia that do not protrude from any specific region have also been observed in several genera, such as Cercomonas and Heteromita (Myl'nikov and Karpov 2004).  These pseudopodia play a major role in food acquisition via phagocytosis.  A majority of cercozoans are uninucleated; however, multinucleated plasmodial stages are not uncommon (Myl'nikov and Karpov 2004).  Like other protists, the nucleus can be located either anteriorly, centrally, or posteriorly, depending on the species. In some large species [e.g., Protaspis grandis], a granular appearance of the nucleus is an indication of permanently condensed chromosomes (Hoppenrath and Leander 2006a). Cytoplasmic inclusions include green plastid, non-plastid pigmented bodies, carbohydrate storage products and lipid droplets.  Cysts are common in many groups, such as Allantion, Cercomonas, Heteromita, Katabia, and Sainouron (Myl'nikov and Karpov 2004). Contractile vacuoles are also usually present in species that inhabit freshwater and soil.  1.3.2 Ultrastructural features of the Cercozoa Morphological information from cercozoans, especially at the ultrastructural level, is very sparse in the literature.  However, ultrastructural studies in representative genera of  4 cercozoans have shown several morphological features, such as mitochondria with tubular cristae, extrusomes, and a multilayered cell wall (Hoppenrath and Leander 2006a; Schnepf and Kühn 2000).  Only the plasmalemma surrounds “athecate” cercozoans, such as Cercomonas spp., Heteromita, and Katabia (Myl'nikov and Karpov 2004).  A nucleus with conspicuous nucleolus and permanently condensed chromosomes has been observed in several cercozoan species, such as Cryothecomonas aestivalis, Ebria tripartita, Protaspis grandis, and Protaspis (ex. Cryothecomonas) longipes (Hargraves 2002; Hoppenrath and Leander 2006a, b; Thomsen et al. 1991).  A microtubular cone (i.e., an array of microtubules around the nucleus originating from the granular end of a rootlet that is associated with a flagellar basal body) has been found in Cercomonas spp.  In addition, a recent comprehensive investigation by Karpov et al. (2006) on the flagellar apparatus of five strains of cercomonads showed a consistent pattern of microtubular roots and a cone of microtubules passing to the nucleus. Although most cercozoans possess mitochondria with tubular cristae, some lineages have flattened tubular cristae [e.g., some Cercomonas] (Karpov et al. 2006), flat or ribbon- like cristae [e.g., Limnofila borokensis, formerly Gymnophrys cometa] (Mikrjukov and Myl'nikov 1998; Nikolaev et al. 2003), or polydiscoid cristae [e.g., Sainouron acronematica] (Cavalier-Smith et al. 2008).  Microbodies (i.e., small single-membrane bounded organelles containing small vesicles and granular matrix) around the nucleus and in parts of the cytoplasm are commonly found in cercozoans, such as Cercomonas, Cholamonas, Heteromita, Katabia, and Massisteria (Myl'nikov and Karpov 2004).  Ejectile organelles or extrusomes of several different kinds, including kinetocysts, microtoxicysts, osmiophilic bodies, and trichocysts are present, depending on the species (Hoppenrath and Leander 2006a; Mikrjukov and Myl'nikov 1998; Myl'nikov and Karpov 2004).  5 Synapomorphies have been identified in some cercozoan subgroups.  For example, cruciform nuclear division has long been recognized in plasmodiophorids (Braselton et al. 1975).  Cavalier-Smith et al. (2008) recently proposed that a flagellar transitional region containing proximal hub-lattices and distal nonagonal fibres is a synapomorphy for all flagellated cercozoans; this ultrastructural complex was found with slight structural variations in the Cercomonadidae, Heteromitidae, Imbricatea, and Sainouridae (Cavalier- Smith et al. 2008).  Although ultrastructural data for representative taxa of different cercozoan subgroups, such as plasmodiophorids and cercomonads, have been extensively studied due to their importance as parasites of economically important plants and essential components of ecological habitats, knowledge of other cercozoans, especially ones living in marine interstitial habitats, is poorly understood.  Therefore, investigating the ultrastructural characteristics of other groups of cercozoans is important and will enable us to better understand the evolutionary history and overall diversity within this particular group of microeukaryotes.  1.4 Biodiversity of the Cercozoa Thus far, fewer than 500 cercozoans have been described, and most of these are from economically important lineages (Adl et al. 2007).  However, Bass and Cavalier-Smith (2004) showed that the diversiy of cercozoans is far greater than currently appreciated and may reach to several thousands of lineages that is on par with two relatively well-established and speciose assemblages, the Ciliophora and the Foraminifera.  This (cryptic) diversity has also been demonstrated with environmental PCR studies (Bass et al. 2009; Brad et al. 2008; Chen et al. 2008; Massana and Pedrós-Alió 2008; Park et al. 2008; Piquet et al. 2008; Šlapeta  6 et al. 2005; Tian et al. 2009).  The rapid discovery of new cercozoan diversity has resulted in major systematic changes within the group over the past few years.  1.5 Classification of the Cercozoa Prior to rDNA sequence data, many current members of the Cercozoa were assigned to several different taxonomic groups that spanned the tree of eukaryotes, due mainly to the lack of diagnostic morphological features and misleading convergent characters shared among evolutionarily distantly related organisms (Bass and Cavalier-Smith 2004; Hoppenrath and Leander 2006a, b).  For example, phaeodareans were previously recognized as radiolarians; plasmodiophorids as fungi or slime molds; Protaspis as euglenids; Ebria as dinoflagellates or silicoflagellates; Gromia as foraminiferans; and Pseudopirsonia as stramenopiles.  Studies based on SSU rDNA sequences have shown that these lineages and several other species previously recognized as enigmatic protists are actually cercozoans (Bass and Cavalier-Smith 2004; Burki et al. 2002; Hoppenrath and Leander 2006a, b; Kühn et al. 2004; Taylor 1990). A comprehensive taxonomic framework for the Cercozoa was attempted by Cavalier-Smith and Chao (2003).  The group is currently composed of two major subclades: the Endomyxa and the Filosa.  Both groups contain free-living and parasitic members and several uncharacterized lineages. The Endomyxa presently contains four major groups: the Ascetosporea, Phytomyxea, Gromiidea, and Proteomyxidea (Bass et al. 2009).  The first two groups contain cercozoan parasites of plants and animals, such as Haplosporidium, Minchinia, Phagomyxa, Plasmodiophora, and Urosporidium; the latter two groups house free-living amoeboid members with branching filopodia or reticulopodia, such as Gromia, Filoreta, and Platyreta (Bass et al. 2009).  Although taxonomic establishment for the Endomyxa is not well  7 established as the Proteomyxidea is currently paraphyletic and requires additional study and revision, the Ascetosporea, Gromiidea, and Phytomyxea are well supported (Bass et al. 2009). The Filosa currently contains most of the described cercozoans: the Chlorarachnea, Granofilosea, Imbricatea, Metromonadea, Thecofilosea, and other subclades, including Auranticordida, Cercomonadida, Glissomonadida, Metopiida, Pansomonadida, Phaeodarea, and Spongomonadida (Bass and Cavalier-Smith 2004; Bass et al. 2005, 2009; Cavalier-Smith and Chao 2003; Chantangsi et al. 2008; Karpov et al. 2006; Polet et al. 2004; Vickerman et al. 2005; Wylezich et al. 2002, 2007).  The Thecofilosea is predominant in both freshwater and marine habitats, and contains uninucleated lineages surrounded by an organic flexible tectum or rigid test with one or two apertures for the emergence of filopodia (Cavalier-Smith and Chao 2003; Chen et al. 2008; Hoppenrath and Leander 2006a, b; Kühn et al. 2004; Lee and Patterson 2000; Lee et al. 2003, 2005; Park et al. 2008; Piquet et al. 2008; Thomsen et al. 1991).  The Thecofilosea is further divided into two major clades, the Tectofilosida and the Cryomonadida; the former houses testate filose amoebae [e.g., Pseudodifflugia], and the latter mostly contains biflagellated cercozoans [e.g., Cryothecomonas and Protaspis] and a testate amoeba genus — Lecythium (Cavalier-Smith and Chao 2003).  The cellular identities and biodiversity of thecofiloseans are poorly understood in comparison to other groups of cercozoans, such as ascetosporeans, cercomonads, chlorarachneans, and phytomyxeans. Improved knowledge of this particular group of cercozoans is required to more comprehensively understand the biodiversity and evolutionary history of cercozoans in general.   8 1.6 Identification of the Cercozoa The absence of unifying features of the Cercozoa at the microscopic level has led to difficulties in identifying these lineages and assigning them to appropriate taxonomic groups. A good example of this difficulty is the taxonomic history of Pseudopirsonia mucosa, which possesses several convergent morphological features with the stramenopile Pirsonia. Pseudopirsonia mucosa was originally placed within Pirsonia, but is now known to be a cercozoan on the basis of molecular evidence (Kühn et al. 1996, 2004).  Several molecular phylogenetic studies have subsequently demonstrated that many protists that were previously placed into different eukaryotic supergroups or were treated as Eukaryota incertae sedis have turned out to be cercozoans, such as Allantion, Allas, Bodomorpha, and Spongomonas (Cavalier-Smith 2000); Cryothecomonas (Kühn et al. 2000); Ebria (Hoppenrath and Leander 2006b); Gymnophrys and Lecythium (Nikolaev et al. 2003); Massisteria (Atkins et al. 2000); Metopion and Metromonas (Bass and Cavalier-Smith 2004); Proleptomonas (Vickerman et al. 2002); and Protaspis (Hoppenrath and Leander 2006a). These results have demonstrated that molecular phylogenetic data are indispensible for the discovery and identification of cercozoans.  This approach is what we currently refer to as DNA barcoding, which works efficiently in the delimitation of animal species and some groups of protists (Barth et al. 2006; Chantangsi et al. 2007; Godfray 2002; Hebert et al. 2003a, b; Lynn and Strüder-Kypke 2006; Saunders 2005; Scicluna et al. 2006; Tautz et al. 2002, 2003).  In fact, a short 20-35 bp region of SSU rDNA sequences has already been used as a molecular signature for the identification of several genera of cercozoans, such as Cercomonas, Neocercomonas, and Heteromita (Ekelund et al. 2004; Karpov et al. 2006). However, this region is much too short for inferring phylogenetic relationships.  Because the Cercozoa is established on the basis of molecular data and the group lacks shared  9 morphological characteristics at the microscopic level, the application of DNA barcoding should be explored more comprehensively in the group.  1.7 Phylogeny of the Cercozoa 1.7.1 The phylogeny of the Cercozoa as inferred from small subunit ribosomal DNA sequences Bass and Cavalier-Smith (2004) used Cercozoa-specific primers to amplify SSU rDNA from environmental samples and demonstrated a great deal of hidden diversity in natural habitats; in other words, several novel cercozoan subclades comprising many uncultured and undescribed taxa were discovered (Bass et al. 2009).  Although SSU rDNA sequences provided useful information about the overall diversity and phylogenetic relationships of cercozoans, the internal phylogeny inferred from this gene remains largely unresolved, especially at deep branching levels.  This has forced biologists interested in this group to find other suitable molecules to infer phylogenetic relationships within the Cercozoa.  1.7.2 The phylogeny of the Cercozoa and its novel polyubiquitin insertions Ubiquitin is a small protein composed of 76 amino acids (Archibald et al. 2003; Bass et al. 2005).  This protein is a small regulatory protein that marks other proteins for destruction. Ubiquitin is found only in eukaryotic organisms and plays essential roles in many biological processes, such as cell cycle regulation, DNA repair, transcriptional regulation, signal transduction, endocytosis, embryogenesis, and apoptosis (programmed cell death) (Hershko and Ciechanover 1998).  The protein is also highly conserved, as indicated by only 3 residue differences between yeast and human ubiquitin amino acid sequences.  The ubiquitin gene has 3 main forms: (1) an individual gene with a single open reading frame; (2) a fusion  10 protein involving a co-translated ribosomal protein gene; and (3) a tandem linear repeat of ubiquitin monomers, giving rise to translation product made up of a chain of ubiquitin proteins, called a polyubiquitin molecule (Bass et al. 2005). Comparisons of ubiquitin amino acid sequences across various eukaryotes have shown a unique amino acid insertion between the monomer-monomer junctions of the polyubiquitin molecule in two major protist groups, the Cercozoa and the Foraminifera (Archibald and Keeling 2004; Archibald et al. 2003; Bass et al. 2005).  The number of residues inserted varies from 1-2 amino acids (Archibald and Keeling 2004; Archibald et al. 2003; Bass et al. 2005).  This molecular signature provides a diagnostic tool to assign lineages to the Cercozoa (or the Foraminifera) (Bass et al. 2005).  Although the variation of the amino acid insertions between polyubiquitin monomers has indicated some interrelationships within the Cercozoa and the Foraminifera, the nucleotide and amino acid sequences of this gene cannot be used to directly construct phylogenetic trees because of the highly conserved nature of this gene.  1.8 Thesis goals and scope The biodiversity and evolutionary relationships among protists have attracted the attention of protistologists for decades.  As a newly established phylum, knowledge on cercozoan diversity and evolution is poorly understood, and the group is being studied only by a small number of protistologists.  In order to expand our knowledge on the morphology, evolutionary relationships, and biodiversity of the Cercozoa, my research involved a great deal of sampling from natural habitats.  The collection of morphological and molecular data from novel cercozoans enabled me to characterize the cellular identities (i.e., provide organismal anchors or reference taxa) for previously established environmental sequence clades.  11 Because several novel lineages of cercozoans were discovered from British Columbia (Bass and Cavalier-Smith 2004), my major sampling sites were concentrated on a wide range of natural habitats within this area, including Agular Point, Bamfield Marine Sciences Centre, Boudary Bay, Brady’s Beach, English Bay, Jericho Park, Spanish Bank, Swamps in Pacific Spirit Regional Park, White Rock, and Wreck Beach.  Psammophilous cercozoans inhabiting sand collected from benthic marine habitats were separated by extraction through meshes of different sizes — 20, 45, 60, 100, and 160 µm — using the melting seawater-ice method (Uhlig 1964).  Cercozoans of interest were isolated using a sterile glass pipette and prepared for light microscopy (LM), scanning electron microscopy (SEM), transmission electron microscopy (TEM), and DNA extraction.  In addition, attempts to culture cercozoans using different media were performed to establish permanent clonal cultures by isolating a single cell using a sterile finely drawn glass pipette.  This would guarantee an availability of examined organisms for further studies.  My research plan had the following five objectives.  1.8.1 Specific aim 1 To explore the hidden biodiversity of cercozoans, in particular those dwelling in marine interstitial habitats. Benthic environments are major ecosystems housing a diverse array of organisms that span the tree of eukaryotes.  From a taxonomic point of view, the “true” biodiversity within this system is largely unknown because studies on benthic habitats have been lacking, compared to those on planktonic environments (Fenchel 1987; Hondeveld et al. 1992).  Some studies have revealed cryptic diversity within these particular habitats (Al Qassab et al. 2002; Bass and Cavalier-Smith 2004; Larsen and Patterson 1990; Lee and Patterson 2000; Lee et al. 2003, 2005).  Comparative and extensive investigation on uncultivated cercozoans inhabiting  12 marine benthic interstitial habitats would help establish morphological features for cercozoan lineages that are either completely unknown or only known from environmental DNA sequences.  1.8.2 Specific aim 2 To test the potential utility and feasibility of a DNA barcoding approach for identifying and delimiting species of marine cercozoans. Molecular sequences, especially SSU rDNA sequences, have long been used as an effective and pragmatic way to identify and classify organisms of uncertain taxonomic affinities (Chantangsi et al. 2007).  The original idea of using these sequences as molecular signatures or DNA barcodes has been proposed for well over a decade (Dawkins 1998; Hebert et al. 2003a, b).  The potential of this approach has been demonstrated for several different lineages of organisms, especially animals (Barth et al. 2006; Chantangsi et al. 2007; Hebert et al. 2003a, b; Lynn and Strüder-Kypke 2006; Saunders 2005; Scicluna et al. 2006). Because in many cases cercozoans lack distinctive morphological features at the microscopic level, organisms within this group are difficult to identify without a DNA barcoding approach.  My goal was to establish a baseline of DNA barcodes for uncultivated marine benthic cercozoans in order to facilitate the identification of species by scientists working in various research areas, especially those whom taxonomy is not their expertise.       13 1.8.3 Specific aim 3 To characterize the morphological features of novel cercozoans at behavioral and ultrastructural levels. Environmental DNA analyses have demonstrated several novel cercozoan subclades (Bass and Cavalier-Smith 2004), but the majority of these lineages are represented by uncultured lineages whose morphological characteristics remain unknown.  An understanding of the ultrastructure of novel taxa has the potential to demonstrate key structural homologies for major cercozoan subclades (i.e., synapomorphies) and other structural innovations associated with endosymbioses, locomotion, and feeding.  These ultrastructural data will also help establish, which cellular features are shared by all cercozoans.  1.8.4 Specific aim 4 To identify patterns of amino acid insertions at the junctions between the monomers of polyubiquitin genes in cercozoans. The presence of 1-2 intermonomeric amino acid insertions of polyubiquitin is recognized as a conserved molecular innovation that is shared only by members of the Cercozoa and the Foraminifera. Amplification and sequencing of polyubiquitin genes in more cercozoans will help establish the robustness of this feature and whether different insertions demarcate the phyletic origins of different cercozoan subclades.    14 1.8.5 Specific aim 5 To better establish the phylogenetic interrelationships of marine cercozoans by combining SSU rDNA and LSU rDNA sequences. Although the cercozoa was established based on molecular phylogenetic analyses of SSU rDNA sequences, many of the internal relationships of the group remain unresolved (Cavalier-Smith and Chao 2003).  Therefore, my goal was to develop a new dataset of molecular markers, namely LSU rDNA sequences, from (mostly uncultivated) marine cercozoans in order to reconstruct a more confident phylogenetic framework for the group. Phylogenetic analysis of a concatenated SSU + LSU rDNA dataset was expected to be an effective and pragmatic way to acquire data from uncultivated lineages and bolster the statistical support for the branches within this framework (Moreira et al. 2007).  This improved framework would form the basis for inferring the evolutionary history of polyubiquitin insertions and specific morphological characters within the Cercozoa.  15      Figure 1.1.  Light micrographs (LMs) illustrating morphological diversity of marine benthic cercozoans (A-K) and some unknown protists (L-O) found in this study.  A. A tetraflagellate cercozoan Auranticordis quadriverberis.  B. Cryothecomonas sp. (strain APCC MC5-1Cryo) with two anterior-directed flagella.  C. Ebria tripartita showing siliceous internal skeletal.  D. Protaspis grandis.  E. Protaspis maior showing two heterodynamic flagella.  F. Protaspis obliqua showing a posterior notch.  G. Protaspis verrucosa.  H. Thaumatomastix sp. showing siliceous cell covering and spines.  I. An undescribed cercozoan whose morphology is similar to Protaspis grandis.  J. An undescribed cercozoan with dorsoventrally flattened cell body and a smooth cell surface.  K. An undescribed cercozoan with sculptured cell surface and several tiny spines.  L. An unknown benthic protist showing anterior-serrated cell shape and pseudopodia emergent from a ventral aperture.  M. An unknown protist found gliding in a benthic marine habitat.  N. An unknown protist with two thick flagella and numerous rod- shaped structures on cell surface.  O. An unknown protist with two anterior-directed flagella inserted subapically.  (Bars = 10 µm)       16     17 1.9 References Adl SM, Leander BS, Simpson AG, Archibald JM, Anderson OR, Bass D, Bowser SS, Brugerolle G, Farmer MA, Karpov S, Kolisko M, Lane CE, Lodge DJ, Mann DG, Meisterfeld R, Mendoza L, Moestrup Ø, Mozley-Standridge SE, Smirnov AV, Spiegel F (2007) Diversity, nomenclature, and taxonomy of protists. 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J Eukaryot Microbiol 54: 347-357   24 CHAPTER 2: AN SSU rDNA BARCODING APPROACH TO THE DIVERSITY OF MARINE INTERSTITIAL CERCOZOANS, INCLUDING DESCRIPTIONS OF FOUR NOVEL GENERA AND NINE NOVEL SPECIES*  2.1 Introduction Marine benthic environments are complex and poorly understood ecosystems containing a diverse array of microorganisms (Fenchel 1987; Hondeveld et al. 1992). Environmental DNA surveys on these systems have demonstrated a significant amount of hidden diversity, especially of so-called cercozoan amoeboflagellates (Bass and Cavalier- Smith 2004; Berney et al. 2004; Šlapeta et al. 2005).  The Cercozoa was initially recognized as a monophyletic assemblage based on molecular phylogenetic studies inferred from rDNA genes (Cavalier-Smith 1998a, b).  Subsequent studies have shown that several protists incertae sedis are also members of this group (e.g., Ebria) and that there are many novel environmental sequence clades that have yet to be described at the microscopic level (Bass and Cavalier-Smith 2004; Hoppenrath and Leander 2006a, b).  Cryomonadida (e.g., Protaspis) is a group of cercozoan biflagellates that glide along substrates with heterodynamic flagella and are common predators in marine benthic habitats (Hoppenrath and Leander 2006a).  These particular cercozoans possess several morphological characteristics (e.g., flattened cell shape, mode of locomotion, and a conspicuous nucleus with condensed chromatin) that resemble distantly related eukaryotes living in the same environments, namely some phagotrophic euglenids, dinoflagellates, and katablepharids. These convergent morphological and behavioral features have led to difficulties in the  * A version of this chapter has been accepted for publication.  Chantangsi C, Leander BS An SSU rDNA barcoding approach to the diversity of marine interstitial cercozoans, including descriptions of four novel genera and nine novel species. Int J Syst Evol Microbiol  25 classification and identification of benthic biflagellates in general, but especially in cercozoan lineages, which are very poorly understood. DNA barcoding has been proposed as an alternative and more precise approach for the delimitation and identification of species; this strategy is expected to be particularly advantageous for understanding the diversity of uncultured microeukaryotic lineages that lack sufficient morphological details for species delimitation (Godfray 2002; Hebert et al. 2003a, b; Tautz et al. 2002, 2003).  Hebert et al. (2003b) proposed the mitochondrial cytochrome c oxidase subunit 1 (cox1) gene as a pragmatic and widely applicable “DNA barcode” for animal species.  Although this gene has proven useful for the identification of animal species and even some protists (Barth et al. 2006; Chantangsi et al. 2007; Lynn and Strüder-Kypke 2006; Saunders 2005), the gene is not applicable for all groups of eukaryotes due to several limitations (Scicluna et al. 2006).  For example, some groups of eukaryotes are amitochondriate and therefore lack the cox1 gene, and most other groups of eukaryotes have yet to be studied at the level of the mitochondrial genome.  Moreover, a few groups of eukaryotes, such as ciliates, have different sized insertions within the cox1 gene that create difficulties in the PCR amplification of the barcode sequences (Cummings 1992; Norman and Gray 1997).  In addition, the mitochondrial genome of a free-living diplonemid Diplonema papillatum shows fragmented coding regions for the cox1 gene of this organism (Marande and Burger 2007). Nuclear small subunit ribosomal DNA (SSU rDNA) sequences have been the most widely used molecular markers to study the phylogenetic relationships of eukaryotes and prokaryotes.  The SSU rRNA gene is present in all organisms and plays a highly conserved role in protein translation that is critical for the survival of cells; therefore, this sequence can be compared across the entire tree of life.  This gene is also present in numerous copies  26 within the genome and contains highly conserved regions that facilitate the design of universal primers for PCR amplification (Long and David 1980; Minelli 1993; Sogin et al. 1986).  SSU rDNA sequences also contain regions of sequence variation that are sufficient and advantageous for DNA barcoding; for instance, Scicluna et al. (2006) demonstrated the utility of a 600-bp segment of SSU rDNA sequences for identifying subtypes of the Blastocystis hominis species complex. In order to better understand the microeukaryotic components of marine benthic ecosystems, I investigated the phenotypic and genetic diversity of uncultured marine interstitial cercozoans that showed morphological and behavioral similarities with previously described species of Protaspis and Cryothecomonas (Hoppenrath and Leander 2006a).  In particular, I examined the potential for coupling high resolution light microscopy with SSU rDNA barcodes (e.g., a 5'-618-bp fragment) for facilitating the systematics of these uncultured lineages.  I also used the light micrographs and SSU rDNA sequences to help establish the cellular identities of several environmental DNA sequence clades.  Overall, this combined approach enabled us to (1) describe nine novel species within four novel genera and one described genus of uncultured marine interstitial cercozoans and (2) establish an efficient and effective protocol for advancing the systematics of this group in the future.  2.2 Materials and methods 2.2.1 Sampling and light microscopy (LM) Sand samples were collected from several habitats around British Columbia, Canada during 2006-2007.  Organisms were extracted from the sand samples through a 48 µm mesh using a melted seawater-ice method described by Uhlig (1964).  Briefly, 2-3 spoons of sand samples were placed into an extraction column wrapped with the mesh, and two to three  27 seawater ice cubes were then put on top of the sand samples and left to melt over several hours.  The organisms of interest were separated through the mesh and concentrated in a Petri dish that was filled with seawater and placed underneath the extraction column.  The Petri dish containing the organisms was then observed using a Leica DMIL inverted microscope. Cells were individually isolated and placed on a slide for light microscopy using phase contrast and differential interference contrast (DIC) microscopy with a Zeiss Axioplan 2 imaging microscope connected to a Leica DC500 color digital camera.  2.2.2 DNA extraction and PCR amplification Cells were individually isolated and washed three times in autoclaved filtered seawater. The numbers of cells from which different SSU rDNA sequences were obtained are as followed: Protaspis obliqua (isolate 1) 1 cell; P. obliqua (isolate 2) 10 cells; the number of isolated cells was not recorded for Protaspis rotunda sp. nov.; Protaspis obaniformis sp. nov. 4 cells; Protaspis oviformis sp. nov. 25 cells; Botuliforma benthica gen. et sp. nov. 14 cells; Ventrifissura artocarpoidea gen. et sp. nov. 1 cell; Ventrifissura foliiformis gen. et sp. nov. 12 cells; Verrucomonas bifida gen. et sp. nov. 15 cells; Verrucomonas longifila gen. et sp. nov. 1 cell; and Discomonas retusa gen. et sp. nov. 1 cell.  DNA was extracted using the protocol provided in the Total Nucleic Acid Purification kit by EPICENTRE (Madison, WI, USA).  Polymerase chain reaction (PCR) with the final reaction volume of 25 µl was performed two rounds in a thermal cycler using puReTaq Ready-To-Go PCR beads (GE Healthcare Bio-Sciences, Inc., Québec, Canada).  The first round PCR was conducted using forward (i.e., NPF1 or PF1) and reverse primers (i.e., R4 or FAD) as listed in Table 2.1. Then, either direct PCR reaction or a gel-purified band of an 1,850-bp region from the first round PCR was used as a template for the second round PCR with the appropriate primers  28 provided in Table 2.1.  The thermal cycler was programmed as follows: hold at 94 oC for 4 min; 5 cycles of denaturation at 94 oC for 30 sec, annealing at 45 oC for 1 min, and extension at 72 oC for 105 sec; 35 cycles of denaturation at 94 oC for 30 sec, annealing at 55 oC for 1 min, and extension at 72 oC for 105 sec; and hold at 72 oC for 10 min.  PCR products corresponding to the expected size were separated by agarose gel electrophoresis and cleaned using the UltraClean™ 15 DNA Purification Kit (MO BIO Laboratories, Inc., CA, USA). The cleaned DNA was cloned into a pCR2.1 vector using the TOPO TA Cloning® kits (Invitrogen Corporation, CA, USA).  Plasmids with the correct insert size were sequenced using BigDye 3.1 and the vector forward and reverse primers, and internal primers with an Applied Biosystems 3730S 48-capillary sequencer (Table 2.1).  2.2.3 Sequence alignment Sequences were assembled and edited using SequencherTM (version 4.5, Gene Codes Corporation, Ann Arbor, Michigan, USA).  Acquired sequences were initially identified by Basic Local Alignment and Search Tool (BLAST) analysis.  New SSU rDNA sequences derived from the newly found cercozoans were aligned using ClustalW (Thompson et al. 1994) implemented in the MEGA (Molecular Evolutionary Genetics Analysis) program version 4 (Tamura et al. 2007) and further refined by eye.  Four multiple sequence alignments were created for phylogenetic analyses.  (1) A 69-taxon global alignment comprising sequences of representatives from all major eukaryotic groups (1,134 unambiguous sites) was constructed to determine the phylogenetic affinities of the newly isolated organisms to other eukaryotic groups.  (2) A 67-taxon cercozoan alignment consisting of cercozoan representatives and extensive short environmental sequences about 1,069 bp in length (923 unambiguous sites) was constructed to determine phylogenetic  29 affinities of the newly isolated organisms to uncharacterized taxa represented by sequences derived from environmental studies.  (3) A 35-taxon cercozoan alignment covering representatives from different cercozoan subgroups and excluding the shorter and unrelated environmental sequences (1,617 unambiguous sites) was constructed to more robustly determine the phylogenetic relationships among the newly described taxa.  (4) A 35-taxon cercozoan alignment including only the barcoding regions of 618 bp in length (583 unambiguous sites) was constructed to compare topologies of phylogenetic relationships inferred from the 1,617-bp alignment 3.  Alignments 3 and 4 are composed of the same composition of examined taxa.  All ambiguous sites were excluded from the alignments prior to phylogenetic analyses. As for barcoding analyses, two alignment files were created for genetic distance calculation: (1) a 17-taxon cercozoan alignment of the almost complete SSU rDNA sequences including 1,798 bp in length (1,745 unambiguous sites) and (2) a 17-taxon cercozoan alignment including only the 5'-half 618-bp barcoding regions (608 unambiguous sites).  Both datasets are composed of the same composition of examined taxa.  The second barcoding alignment file was constructed based on diagnostic barcoding regions of Blastocystis hominis (Scicluna et al. 2006): its starting position is 22 bp inside the beginning of B. hominis barcoding region and goes 65 bp further after the end of the B. hominis barcoding region.  The alignment files are available upon request.  2.2.4 Phylogenetic analyses MrBayes version 3.1.2 was used to perform Bayesian analyses on all four datasets (Huelsenbeck and Ronquist 2001; Ronquist and Huelsenbeck 2003).  Four Markov Chain Monte Carlo (MCMC) chains — 1 cold chain and 3 heated chains — were run for 2,000,000  30 generations, sampling every 50th generation (tree).  The first 4,000 trees were discarded as burn-in.  The remaining trees were used to compute the 50% majority-rule consensus tree. Branch lengths of the trees were saved. Maximum likelihood analyses were performed on all four datasets using PhyML (Guindon and Gascuel 2003).  Input trees for each dataset were generated by BIONJ with optimization of topology, branch lengths, and rate parameters selected.  The General Time Reversible (GTR) model of nucleotide substitution was chosen.  The proportion of invariable sites and the gamma distribution parameter were estimated from the input dataset.  Eight categories of substitution rates were selected.  PhyML bootstrap trees with 100 bootstrap datasets were constructed using the same parameters as the individual ML trees.  2.2.5 Genetic distance analyses Sequence divergences were calculated for 1,798-bp and 618-bp datasets using the Kimura two-parameter (K2P) distance model with complete deletion for positions with gaps in effect (Kimura 1980).  2.2.6 Sequence availability The SSU rDNA nucleotide sequences included in analyses for this paper are available from the GenBank database under the following accession numbers: Allantion sp. (AF411265), Allas diplophysa (AF411262), Auranticordis quadriverberis (EU484393), Bodomorpha minima (AF411276), Cercomonas plasmodialis (AF411268), cercozoa sp. (AF411273), Cryothecomonas aestivalis (AF290539), C. aestivalis (AF290541), Cryothecomonas longipes (AF290540), Ebria tripartita (DQ303922), E. tripartita (DQ303923), Euglypha rotunda (AJ418784), Heteromita globosa (U42447), Lecythium sp.  31 (AJ514867), Limnofila borokensis (previously misidentified as Gymnophrys cometa) (AF411284), Massisteria marina (AF174372), Mesofila limnetica (previously referred to it as Dimorpha-like sp.) (AF411283), Metopion-like sp. (AF411278), Paulinella chromatophora (X81811), Proleptomonas faecicola (AF411275), Protaspis grandis (DQ303924), Pseudodifflugia cf. gracilis (AJ418794), Pseudopirsonia mucosa (AJ561116), Rigidomastix-like sp. (AF411279), Spongomonas minima (AF411280), Thaumatomastix sp. (AF411261), Thaumatomonas seravini (AF411259), uncultured cercozoan [marine BOLA322] (AF372764), uncultured cercozoan [marine BOLA383] (AF372765), uncultured cercozoan (AY180012), uncultured cercozoan [oxygen-depleted marine CCW29] (AY180018), uncultured cercozoan (AY180035), uncultured cercozoan [freshwater F7] (AY620276), uncultured cercozoan [freshwater PP2-3D] (AY620277), uncultured cercozoan [soil 4-3.5] (AY620279), uncultured cercozoan [soil 4-3.7] (AY620280), uncultured cercozoan [freshwater F9] (AY620281), uncultured cercozoan [soil 9-3.3] (AY620295), uncultured cercozoan [marine D6] (AY620316), uncultured cercozoan [marine A11] (AY620320), uncultured cercozoan [marine A14] (AY620321), uncultured cercozoan [marine D10] (AY620322), uncultured cercozoan [marine D14] (AY620323), uncultured cercozoan [marine 7-6.5] (AY620340), uncultured cercozoan [marine 12-3.6] (AY620349), uncultured cercozoan [marine 12-4.4] (AY620350), uncultured cercozoan [marine C15] (AY620351), uncultured cercozoan [marine C6] (AY620352), uncultured cercozoan [marine C9] (AY620353), uncultured cercozoan [marine 7-6.1] (AY620355), uncultured cercozoan [marine 7-6.4] (AY620357), uncultured cercozoan [freshwater CV1_B1_11] (AY821946), uncultured cercozoan [freshwater PCA4AU2004] (DQ243995), uncultured cercozoan [freshwater PCG5AU2004] (DQ243996), uncultured Cryothecomonas (AY628366), uncultured eukaryote [marine TAGIRI-1] (AB191409), uncultured eukaryote [marine  32 TAGIRI-2] (AB191410), uncultured eukaryote [marine NAMAKO-3] (AB252743), uncultured eukaryote (marine NAMAKO-4) (AB252744), uncultured eukaryote [marine NAMAKO-5] (AB252745), uncultured eukaryote [marine NAMAKO-6] (AB252746), uncultured eukaryote [marine NAMAKO-9] (AB252749), uncultured eukaryote [marine NAMAKO-10] (AB252750), uncultured eukaryote [marine NAMAKO-15] (AB252755), uncultured eukaryote [marine NAMAKO-16] (AB252756), uncultured eukaryote [marine DSGM-58] (AB275058), uncultured eukaryote [Río Tinto river RT3n19] (AY082998), uncultured eukaryote [estuary BB01_58] (AY885050), uncultured eukaryote [estuary BB01_98] (AY885055), uncultured eukaryote [marine NOR26.10] (DQ314809), uncultured eukaryote [marine NW617.37] (DQ314810), uncultured eukaryote [marine NOR46.14] (DQ314811), uncultured eukaryote [marine NOR46.27] (DQ314814), uncultured eukaryote [oxygen-depleted marine D1P01B11] (EF100205), and unidentified eukaryote [Lake Ketelmeer LKM45] (AJ130856).   GenBank accession numbers for the 11 new sequences obtained from this study are: Protaspis obliqua isolate 1 (1,826 bp; FJ824121), P. obliqua isolate 2 (1,826 bp; FJ824122), Protaspis rotunda sp. nov. (1,824 bp; FJ824123), Protaspis obaniformis sp. nov. (1,823 bp; FJ824124), Protaspis oviformis sp. nov. (1,823 bp; FJ824125), Botuliforma benthica gen. et sp. nov. (1,827 bp; FJ824126), Ventrifissura artocarpoidea gen. et sp. nov. (1,824 bp; FJ824127), Ventrifissura foliiformis gen. et sp. nov. (1,822 bp; FJ824128), Verrucomonas bifida gen. et sp. nov. (1,827 bp; FJ824129), Verrucomonas longifila gen. et sp. nov. (1,828 bp; FJ824130), and Discomonas retusa gen. et sp. nov. (1,824 bp; FJ824131).     33 2.3 Results 2.3.1 Identification of the examined flagellates Eleven different isolates of uncultured cercozoan flagellates were isolated and characterized with light microscopy (Figures 2.1-2.3) and SSU rDNA sequences (Table 2.2, Figures 2.4-2.5).  All organisms were found gliding in marine interstitial habitats, except for Botuliforma benthica gen. et sp. nov., which showed a rotational pattern of swimming.  The DNA sequence data demonstrated that the newly discovered cercozoan flagellates clustered with a diverse assortment of environmental DNA sequences, forming one previously established clade and four novel clades, as described below (Figures 2.4-2.5).  These molecular data were served as our primary taxonomic guides and enabled us to establish nine new species that are challenging to distinguish at the morphological level.  2.3.2 The Cryomonadida clade Phylogenies inferred from SSU rDNA sequences demonstrated that one previously described species, Protaspis obliqua, and the following three new species of Protaspis clustered within the Cryomonadida clade, which also contained Protaspis grandis and Cryothecomonas spp. (Figures 2.4-2.5).  The genus Protaspis currently contains 11 described species - P. gemmifera, P. glans, P. grandis, P. maior, P. metarhiza, P. obliqua, P. obovata, P. simplex, P. tanyopsis, P. tegere, and P. verrucosa.  Protaspis rotunda sp. nov. Chantangsi and Leander, 2009 Size. Cells 25-27 µm wide and 30-31 µm long (Numbers of cells observed > 100). Diagnosis. Cells dorsoventrally flattened and slightly oval to round in outline with smooth surface; uninucleate biflagellate; flagella inserted subapically separated by an  34 anterior protrusion; nucleus is 11 µm wide and 15 µm long; nucleus is ovoid and located at anterior to the right of the cell; clear and homogeneous cytoplasm; curved furrow at the sub- anterior ventral side; locomotion by gliding; found living in marine interstitial sand habitat. Small subunit rRNA gene sequence [GenBank accession no. FJ824123].  Protaspis rotunda differs from the 11 previously described species in that this species shows a smooth cell surface.  Although similar to P. oviformis in shape and cell appearance, nuclear position of P. rotunda is at anterior to the right of the cell as opposed to at posterior or sometimes middle right of the cell in P. oviformis.  Cell shapes of P. grandis and P. obaniformis are rather oblong than the oval shape of P. rotunda. Type locality. Tidal sand-flat at Pachena Beach (48°47'N, 125°07'W), Vancouver Island, British Columbia, Canada.  The organisms were collected in September 2006.  This species was also found in June 2007 and June 2008. Iconotype. Figures 2.1a, 2.2c, and 2.3a. Etymology. The etymology for the specific epithet, Latin fem. rotunda, round.  The specific epithet reflects the round shape of this organism.  Protaspis obaniformis sp. nov. Chantangsi and Leander, 2009 Size. Cells 30-50 µm wide and 50-65 µm long (Numbers of cells observed = 19). Diagnosis. Cells are broadly elliptical and dorsoventrally flattened; thick wall with smooth surface; uninucleate biflagellate; flagella inserted subapically; nucleus is 18 µm wide and 17 µm long; circular nucleus with granular (permanently condensed) chromosomes; nucleus is located in the middle of the cell, sometimes towards the anterior of the cell; cytoplasm contains numerous spherical light brownish granules; prominent vertically straight slit at the posterior ventral side; locomotion by gliding; found living in marine interstitial  35 sand habitat.  Small subunit rRNA gene sequence [GenBank accession no. FJ824124]. Protaspis obaniformis differs from the 11 previously described species in that this species possesses an 1/5 cell length posterior slit.  Although Protaspis obliqua also shows a ventral furrow/groove at the posterior half of the cell, the posterior notch of P. obliqua has never been observed in P. obaniformis.  In addition, cell shape and cell size of the former (i.e., 10- 27 µm wide and 8-32 µm long) are different from the latter. Type locality. Tidal sand-flat at Boundary Bay (49°00'N, 123°02'W), Vancouver, British Columbia, Canada.  The organisms were collected in May 2007. Iconotype. Figures 2.1b and 2.3b. Etymology. The etymology for the specific epithet, Japanese oban is a kind of ancient Japanese coin with a broad oval shape; Latin fem. forma, shape.  The specific epithet, New Latin fem. obaniformis, depicts the shape of this organism, which is similar to an ancient Japanese coin.  Protaspis oviformis sp. nov. Chantangsi and Leander, 2009 Size. Cell is 25-40 µm wide and 30-45 µm long (Numbers of cells observed = 36). Diagnosis. Cells are dorsoventrally flattened and slightly oval to roundish in outline with smooth surface; slightly thick wall; uninucleate cell with an anterior protrusion; nucleus is 8 µm wide and 8 µm long; nucleus with chromosome appearance is circular in outline and located at posterior or sometimes middle right of the cell; cytoplasm contains numerous spherical brownish, grayish, and yellowish granules; vertical ventral slit positioned at the mid anterior end and towards the mid posterior end; finger-like pseudopodia present; locomotion by gliding; found living in marine interstitial sand habitat.  Small subunit rRNA gene sequence [GenBank accession no. FJ824125].  Protaspis oviformis differs from the 11  36 previously described species in that this species possesses a very oval shape and smooth cell surface.  Nuclear position of this species is located differently from its close relatives, P. grandis (i.e., posterior at the middle of the cell) and P. rotunda (i.e., anterior to the right of the cell).  Unlike P. obaniformis that possesses a posterior ventral slit, this structure of P. oviformis is located at the mid anterior end and towards the mid posterior end of the cell. Type locality. Tidal sand-flat at Boundary Bay (49°00'N, 123°02'W), Vancouver, British Columbia, Canada.  The organisms were collected in March 2007. Iconotype. Figures 2.1c, 2.2d-e, and 2.3c. Etymology. The etymology for the specific epithet, Latin neut. ovum, egg; Latin fem. forma, shape.  The specific epithet reflects the shape of this organism.  Protaspis longipes (Schnepf and Kühn, 2000) Chantangsi and Leander comb. nov. basionym: Cryothecomonas longipes Schnepf and Kühn, 2000.  2.3.3 The Botuliformidae lineage Phylogenies inferred from SSU rDNA sequences demonstrated that one new genus/species clustered strongly with the “Ebriid” clade comprising an Ebria tripartita sequence and an environmental sequence (Figures 2.4-2.5).  Genus Botuliforma gen. nov. Chantangsi and Leander, 2009 Diagnosis. Cells are oblong; slightly thick wall with rough surface; uninucleate biflagellate; flagella inserted subapically; large nucleus with granular (permanently condensed) chromosomes located at anterior end of the cell; colorless cytoplasm; ventral  37 furrow present; very fine filopodia observed; extrusomes present; locomotion by rotational swimming; found living in marine interstitial sand habitats. Type species. Botuliforma benthica. Etymology. The etymology for the generic name, Latin masc. botulus, sausage; Latin fem. forma, shape.  The genus name reflects the sausage shape of these organisms.  Botuliforma benthica sp. nov. Chantangsi and Leander, 2009 Size. Cell is about 20 µm wide and 35 µm long (Numbers of cells observed = 51). Diagnosis. Structure as described for the genus; nucleus is 12 µm wide and 15 µm long; large nucleus is subspherical in outline and located at the anterior end of the cell; aggregation of cells forming a swimming spherical ball observed.  Small subunit rRNA gene sequence [GenBank accession no. FJ824126]. Type locality. Tidal sand-flat at Pachena Beach (48°47'N, 125°07'W), Vancouver Island, British Columbia, Canada.  The organisms were collected in June 2007 and June 2008. Iconotype. Figures 2.1d, 2.2k, and 2.3d. Etymology. The etymology for the specific epithet, New Latin benthica, of the benthos.  The specific epithet reflects the natural habitat of this organism.  2.3.4 The Ventrifissuridae clade Phylogenies inferred from SSU rDNA sequences demonstrated that one new genus and two new species clustered together with several environmental sequences and formed the Ventrifissuridae clade (Figures 2.4-2.5).   38 Genus Ventrifissura gen. nov. Chantangsi and Leander, 2009 Diagnosis. Cells are broadly obovate and dorsoventrally flattened; cells with either smooth surfaces or with numerous pointed warts; uninucleate biflagellate; flagella inserted subapically with or without an anterior protrusion; circular to oblong shaped nucleus located at the anterior end of the cell; colorless cytoplasm; ventral furrow present; filopodia observed; extrusomes present; locomotion by gliding; found living in marine interstitial sand habitat. Type species. Ventrifissura artocarpoidea. Etymology. The etymology for the generic name, Latin masc. ventris, belly; Latin fem. fissura, crack, cleft, or chink.  The genus name reflects the morphological feature of members within this genus of having a slit on ventral side.  Ventrifissura artocarpoidea sp. nov. Chantangsi and Leander, 2009 Size. Cell is about 35-36 µm wide and 43-45 µm long (Numbers of cells observed = 3). Diagnosis. Cells are broadly obovate and dorsoventrally slightly flattened; numerous pointed warts distributed evenly over the cell surface; uninucleate biflagellate; flagella inserted subapically without a protrusion; circular nucleus located at the anterior end of the cell, sometimes toward the left side; colorless cytoplasm, sometimes with food particles; ventral furrow present; locomotion by gliding; found living in marine interstitial sand habitat. Small subunit rRNA gene sequence [GenBank accession no. FJ824127]. Type locality. Tidal sand-flat at Boundary Bay (49°00'N, 123°02'W), Vancouver, British Columbia, Canada.  The organisms were collected in May 2007. Iconotype. Figures 2.1e, 2.2g, and 2.3e.  39 Etymology. The etymology for the specific epithet, New Latin masc. artocarpus, a genus of breadfruit; Latin fem. –oidea, suffix denoting resembling.  The specific epithet reflects the shape of this organism which is similar to a breadfruit.  Ventrifissura foliiformis sp. nov. Chantangsi and Leander, 2009 Size. Cell is about 30-35 µm wide and 40-47 µm long (Numbers of cells observed > 100). Diagnosis. Cells are broadly obovate and (extremely) dorsoventrally flattened with a smooth cell surface; uninucleate biflagellate; flagella inserted subapically between an anterior protrusion; nucleus is 15 µm wide and 10 µm long; oblong nucleus with granular appearance located at the anterior end of the cell, sometimes toward the right side; colorless cytoplasm, sometimes with food particles; small clear vacuoles present; ventral furrow present; filopodia observed; extrusomes present; locomotion by gliding; found living in marine interstitial sand habitat.  Small subunit rRNA gene sequence [GenBank accession no. FJ824128]. Type locality. Tidal sand-flat at Boundary Bay (49°00'N, 123°02'W), Vancouver, British Columbia, Canada.  The organisms were collected in May 2007. Iconotype. Figures 2.1f, 2.2f, 2.2h, and 2.3f.   Etymology. The etymology for the specific epithet, Latin neut. folium, leaf; Latin fem. forma, shape.  The specific epithet reflects the shape of this organism which is flattened like a leaf.     40 2.3.5 The Verrucomonadidae clade Phylogenies inferred from SSU rDNA sequences demonstrated that one new genus and two new species clustered together and formed the Verrucomonadidae clade (Figures 2.4- 2.5).  Genus Verrucomonas gen. nov. Chantangsi and Leander, 2009 Diagnosis. Cells are dorsoventrally flattened and with a rough surface; colored warts ranging from yellowish, red, brownish, and golden observed on cell surface; uninucleate biflagellate; flagella inserted subapically; nucleus is located at the anterior end of the cell; colorless cytoplasm; ventral furrow present; anterior and posterior notches sometimes present; filopodia observed; extrusomes present; locomotion by gliding; found living in marine interstitial sand habitat. Type species. Verrucomonas bifida. Etymology. The etymology for the generic name, Latin fem. verruca, wart; Latin fem. monas, a unit (which refers to the flagellate).  The genus name reflects the morphological feature of having colored warts on the cell surface.  Verrucomonas bifida sp. nov. Chantangsi and Leander, 2009 Size. Cell is about 17-25 µm wide and 30-34 µm long (Numbers of cells observed > 100). Diagnosis. Cells are dorsoventrally flattened with a rough surface; colored warts ranging from yellowish, red, brownish, and golden observed on cell surface; uninucleate biflagellate; flagella inserted subapically; bilobed nucleus, 20 µm wide and 13 µm long, located at the anterior of the cell; colorless cytoplasm; ventral furrow present; anterior and  41 posterior notches sometimes present; fine filopodia observed; extrusomes present; locomotion by gliding; found living in marine interstitial sand habitat.  Small subunit rRNA gene sequence [GenBank accession no. FJ824129]. Type locality. Tidal sand-flat at Boundary Bay (49°00'N, 123°02'W), Vancouver, British Columbia, Canada.  The organisms were collected in May 2007. Iconotype. Figures 2.1g, 2.2i, and 2.3g. Etymology. The etymology for the specific epithet, Latin fem. bifida, two-cleft.  The specific epithet reflects the shape of organism’s nucleus which seems to be divided into two parts.  Verrucomonas longifila sp. nov. Chantangsi and Leander, 2009 Size. Cell is about 15-20 µm wide and 23-37 µm long (Numbers of cells observed = 11). Diagnosis. Cells are dorsoventrally flattened with a rough surface; yellowish and reddish warts observed on cell surface; uninucleate biflagellate; flagella inserted subapically; elliptical nucleus with granular appearance, 8 µm wide and 5 µm long, located at anterior end of the cell; colorless cytoplasm; ventral furrow present; anterior and posterior notches sometimes present; fine filopodia observed; extrusomes present; locomotion by gliding; found living in marine interstitial sand habitat.  Small subunit rRNA gene sequence [GenBank accession no. FJ824130]. Type locality. Tidal sand-flat at Spanish Banks (49°16'N, 123°14'W), Vancouver, British Columbia, Canada.  The organisms were collected in April 2007. Iconotype. Figures 2.1h, 2.2j, and 2.3h.  42 Etymology. The etymology for the specific epithet, Latin masc. longus, long; Latin fem. fila, thread.  The specific epithet reflects the longer flagella of this organism in reference to its close relative, Verrucomonas bifida.  2.3.6 The Discomonadidae clade Phylogenies inferred from SSU rDNA sequences demonstrated that one new genus/species clustered with an environmental sequence and formed the Discomonadidae clade (Figures 2.4-2.5).  Genus Discomonas gen. nov. Chantangsi and Leander, 2009 Diagnosis. Cells are disc-shaped and dorsoventrally flattened; uninucleate biflagellate; flagella inserted subapically; nucleus is located in the middle-anterior of the cell; anterior notch present; colorless cytoplasm, sometimes with food particles at the posterior end of the cell; locomotion by gliding; found living in marine interstitial sand habitat. Type species. Discomonas retusa. Etymology. The etymology for the generic name, Latin masc. discus, flat, circular plate; Latin fem. monas, a unit (which refers to the flagellate).  The genus name reflects the shape of this organism.  Discomonas retusa sp. nov. Chantangsi and Leander, 2009 Size. Cell is about 25 µm wide and 25 µm long (Numbers of cells observed = 5). Diagnosis. Structure as described for the genus; nucleus located at the middle-anterior of the cell is 9 µm wide and 9 µm long; discoidal nucleus with central nucleolus.  Small subunit rRNA gene sequence [GenBank accession no. FJ824131].  43 Type locality. Tidal sand-flat at Boundary Bay (49°00'N, 123°02'W), Vancouver, British Columbia, Canada.  The organisms were collected in May 2007. Iconotype. Figures 2.1i, 2.2l, and 2.3i. Etymology. The etymology for the specific epithet, Latin fem. retusa, notched at the apex.  The specific epithet reflects the notched appearance at the anterior end of the organism.  2.3.7 DNA barcoding marine benthic cercozoans DNA analyses based on the Kimura 2 parameter (K2P) model of 1798-bp full length and 618-bp barcoding regions of SSU rDNA sequences of 17 benthic cercozoans showed relatively similar values of average percent sequence divergences, 7.57 and 6.98, respectively.  Analyses of the barcoding region of isolates of three species, including Cryothecomonas aestivalis, Ebria tripartita, and Protaspis obliqua, showed low intraspecific sequence variation and only a few nucleotide differences (Table 2.2).  In general, percent intergeneric sequence divergences between examined genera are quite high (Table 2.2).  It is also significant to note that Cryothecomonas longipes showed lower percent sequence divergences and fewer numbers of nucleotide differences to Protaspis grandis (Figure 2.2a) than to C. aestivalis, which is consistent with the molecular phylogenetic data (Figures 2.4- 2.5).  Moreover, two isolates of P. obliqua (Figure 2.2b) showed high sequence divergences and large numbers of nucleotide differences to the other species/morphotypes currently recognized as “Protaspis”.     44 2.3.8 Molecular phylogenetic analyses of marine benthic cercozoans Phylogenies deduced from 1,617-bp full length and 583-bp barcoding region of SSU rDNA sequences of 35 cercozoan taxa, including representatives from several major cercozoan subgroups, showed very similar tree topologies (Figure 2.4).  Protaspis obliqua and three new species of Protaspis, namely P. rotunda, P. obaniformis, and P. oviformis, clustered within the Cryomonadida clade, which currently contains three genera – Cryothecomonas, Lecythium, and Protaspis (Figure 2.4).  However, P. obliqua was positioned distantly from the other Protaspis spp., and C. longipes was placed within the main Protaspis lineage with very strong statistical support (Figures 2.4-2.5).  The nearest sister group to the Cryomonadida was a clade consisting of Botuliforma benthica gen. et sp. nov., Ebria tripartita, and several undescribed cercozoans derived from environmental DNA surveys (Figure 2.5).  The remaining five species described here – namely Ventrifissura artocarpoidea gen. et sp. nov., Ventrifissura foliiformis gen. et sp. nov., Verrucomonas bifida gen. et sp. nov., Verrucomonas longifila gen. et sp. nov., and Discomonas retusa gen. et sp. nov. – branched with different undescribed cercozoans derived from environmental DNA surveys, forming three different clades with morphological features that remained unknown prior to this study.  2.4 Discussion 2.4.1 Hidden diversity of marine benthic cercozoans Marine planktonic and benthic habitats house a large number of microeukaryotic lineages (Massana and Pedrós-Alió 2008; Park et al. 2008).  The actual biodiversity in benthic environments, however, is not well understood because studies on these habitats are relatively infrequent compared to studies on planktonic environments (Bass and Cavalier-  45 Smith 2004; Chantangsi et al. 2008; Fenchel 1987; Hondeveld et al. 1992; Hoppenrath and Leander 2006a; Lee 2008).  Nonetheless, several studies based on environmental PCR analyses of SSU rDNA have shown that cercozoans, such as Auranticordis, Cercomonas, Massisteria, Metopion, Metromonas, Protaspis, and Thaumatomonas, are major components of benthic habitats (Al Qassab et al. 2002; Bass and Cavalier-Smith 2004; Chantangsi et al. 2008; Hoppenrath and Leander 2006a; Myl'nikov and Karpov 2004; Park et al. 2008). Protaspis, in particular, has been reported from several aquatic and terrestrial environments worldwide (Auer and Arndt 2001; Ekelund and Patterson 1997; Hoppenrath and Leander 2006a; Larsen and Patterson 1990; Lee et al. 2003, 2005; Lee and Patterson 2000; Vørs 1993).  2.4.2 The current composition of Protaspis Protaspis was originally described by Skuja (1939) and currently contains 11 valid species: P. gemmifera, P. glans, P. grandis, P. maior, P. metarhiza, P. obliqua, P. obovata, P. simplex, P. tanyopsis, P. tegere, and P. verrucosa.  However, my study shows the actual diversity of the group to be far greater.  I have assigned three new species – P. rotunda, P. obaniformis, and P. oviformis – to this genus.  These novel flagellates share several common generic features with the type species: (1) a rigid cell body with two heterodynamic flagella (at least in the first two species), (2) both flagella insert subapically on the ventral side of the cell (at least in the first two species), and (3) cells are dorsoventrally flattened and possess a ventral slit from which pseudopodia can be protruded (Hoppenrath and Leander 2006a; Myl'nikov and Karpov 2004; Skuja 1939).  However, the three new Protaspis spp. described here did not possess the specific cell shapes and diagnostic features found in the previously described species.  46 My molecular phylogenetic analyses demonstrated that the three novel species of Protaspis grouped strongly with P. grandis.  However, two isolates of Protaspis obliqua, whose SSU rDNA sequences were sequenced in this study, were positioned very distantly from the other Protaspis spp. in my molecular phylogenetic analyses; this result was consistent with the large number of nucleotide differences between P. obliqua and the other Protaspis sequences and led us to doubt the taxonomic status of P. obliqua.  These data also suggest that a great deal of genetic diversity is hidden at the morphological level (Weisse 2008).  However, further comparative investigations of ultrastructural features in Protaspis spp. (especially the type species Protaspis glans) might demonstrate differences that are consistent with the molecular phylogenetic data and lead to the future taxonomic reassignment of P. obliqua.  2.4.3 Cryothecomonas longipes is more closely related to Protaspis sensu stricto Cryothecomonas was shown to be the closest relative of Protaspis based on molecular phylogenetic evidence and some ultrastructural features (Hoppenrath and Leander 2006a; Thomsen et al. 1991).  Six species of Cryothecomonas, namely C. aestivalis, C. armigera, C. inermis, C. longipes, C. scybalophora, and C. vesiculata, have been described thus far and only two of them – C. aestivalis and C. longipes – have had their SSU rDNA sequenced (Drebes et al. 1996; Kühn et al. 2000; Schnepf and Kühn 2000; Thomsen et al. 1991).  My phylogenetic analyses demonstrated a closer relationship between C. longipes and the genus Protaspis (excluding P. obliqua) than to C. aestivalis, which was consistent with previous results (Hoppenrath and Leander 2006a). Thomsen et al. (1991) established Cryothecomonas as a new genus on the basis of differences between these species and the previously established genus Protaspis, such as the  47 configuration of the flagellar apparatus and the location of the cell slit/groove (Thomsen et al. 1991).  Flagella are homodynamic and inserted apically in Cryothecomonas, whereas Protaspis spp. have heterodynamic flagella inserted subapically (Skuja 1939; Thomsen et al. 1991).  The cell slit/groove, where pseudopodia emerge, is located posterior-laterally in Cryothecomonas and ventral-medially in Protaspis (Skuja 1939; Thomsen et al. 1991). Ultrastructural studies on the latest member of Cryothecomonas – namely C. longipes – demonstrated morphological differences between this species and the other four species in the genus Cryothecomonas, which were all described together when this genus was first established (Thomsen et al. 1991).  For instance, C. longipes possesses heterodynamic flagella; the anterior flagellum is inserted apically, and the posterior flagellum is inserted subapically.  Moreover, the ventral slit in both C. longipes and Protaspis is located on the ventral side of the cell.  If flagellar orientation and the location of longitudinal groove were the key features that rendered Thomsen et al. (1991) to separate Cryothecomonas from Protaspis, then C. longipes should not belong to the former genus.  Several ultrastructural features shared by C. longipes and P. grandis have also been demonstrated, such as the presence of a multilayered cell wall, a nucleus with a prominent nucleolus and condensed chromosomes, structurally identical extrusomes, and flagellar pits with distinctive funnels (Hoppenrath and Leander 2006a; Schnepf and Kühn 2000).  All of these data are consistent with my molecular phylogenetic analyses that showed C. longipes within the main Protaspis clade.  For these reasons, I have transferred C. longipes to the genus Protaspis: P. longipes comb. nov. (see Results section).    48 2.4.4 The benthic Botuliforma benthica gen. et sp. nov. is closely related to the planktonic Ebria tripartita I have discovered a benthic swimming flagellate, namely B. benthica gen. et sp. nov., that is the nearest sister lineage to the clade consisting of Ebria tripartita and environmental sequence EF100205 as inferred from SSU rDNA sequences (Figure 2.5).  Interestingly, the environmental sequence was generated from the upper 2 cm of oxygen-depleted intertidal marine sediments, which is nearly identical to the habitat where I collected B. benthica gen. et sp. nov.  The distinctively different DNA sequences, habitats and morphological features of Botuliforma and Ebria led us to establish a new genus for this novel benthic organism. The “Ebriida” clade was previously shown to be a member of the Cercozoa and a close sister lineage to the Cryomonadida in SSU rDNA phylogenies (Hoppenrath and Leander 2006b). Although morphological similarities between representatives of these two clades were not initially obvious, shared features have been demonstrated at the ultrastructural level (Hoppenrath and Leander 2006b).  Ebria, Cryothecomonas, and Protaspis all share two unequal flagella, a nucleus with a prominent nucleolus and permanently condensed chromosomes, tubular mitochondrial cristae (at least in the last two genera), and feeding by means of pseudopodia (Hargraves 2002; Hoppenrath and Leander 2006a, b; Thomsen et al. 1991).  All current members of the Cryomonadida (Cryothecomonas, Lecythium, and Protaspis) possess a test, layered wall or theca around the cells; by contrast, Ebria possesses a naked cell with a fine layer of fibrillar material lying outside of the plasma membrane and an internal siliceous skeleton (Hargraves 2002). The combination of features in B. benthica gen. et sp. nov. appears to be transitional between ebriids and cryomonads.  For example, B. benthica has a thick and rough wall that might be homologous to the multilayered cell walls of Cryothecomonas and Protaspis.  In  49 addition, a prominent nucleus with condensed chromosomes, fine pseudopodia and extrusomes were easily observed in B. benthica under light microscopy (Figures 2.1d, 2.2k), and these features are also found in Cryothecomonas and Protaspis (Drebes et al. 1996; Hoppenrath and Leander 2006a; Schnepf and Kühn 2000; Thomsen et al. 1991).  Although ultrastructural data are currently unavailable for B. benthica gen. et sp. nov., the close relationship between this lineage and E. tripartita is very robust in molecular phylogenies inferred from SSU rDNA.  Therefore, transmission electron microscopy of B. benthica gen. et sp. nov. might demonstrate the existence of inconspicuous siliceous skeletal elements that are homologous with ebriids.  2.4.5 The cellular identities of previously undescribed cercozoans I established three new genera in addition to Botuliforma gen. nov. – namely Discomonas gen. nov., Ventrifissura gen. nov., and Verrucomonas gen. nov. – on the basis of their very distant molecular phylogenetic positions inferred from SSU rDNA sequences (Figures 2.4-2.5).  My phylogenetic analyses placed two new species of Ventrifissura – V. artocarpoidea and V. foliiformis – within a very distinct clade consisting of three environmental sequences derived from marine environments: AY180018, AY180035, and AY620350 (Bass and Cavalier-Smith 2004; Stoeck and Epstein 2003).  The two new species of Verrucomonas were separated from the other cercozoans by very long branch-lengths (Figure 2.5).  This Verrucomonadidae clade clustered weakly with two environmental sequences also obtained from marine environments – AY180012 and AY620349 (Bass and Cavalier-Smith 2004; Stoeck and Epstein 2003) – and this more inclusive clade formed the nearest sister group to the Ventrifissuridae clade (Figure 2.5).  Analyses by Bass and Cavalier-Smith (2004) showed that several of these environmental sequences (e.g.,  50 uncultured cercozoan AY620349) are members of the Tectofilosida, and the morphological features I describe here help establish the cellular identities for some of the environmental sequence clades contained therein. The SSU rDNA sequence of Discomonas gen. nov. formed the nearest sister lineage to uncultured cercozoan AY620316 (Figure 2.5).  Bass and Cavalier-Smith (2004) recognized this environmental sequence as an undescribed member of ‘Basal Group T’, which branched closely with thaumatomonads.  My analyses of two different datasets, however, showed incongruent phylogenetic positions for Discomonas gen. nov.  Analyses of the 923-bp dataset showed Discomonas gen. nov. branching with environmental sequence AY620316, and analyses of the 1,617-bp dataset showed Discomonas gen. nov. branching next to the Cercomonadida lineage [e.g., Cercomonas plasmodialis] (Figure 2.4).  This incongruence was also observed in Bass and Cavalier-Smith (2004); they reported that AY620316 can sometimes be a sister lineage to the Metopiida and a few other sequences recognized by them as ‘Novel Clade 6’.  However, these relationships were recovered with only weak statistical support.  Although these organisms need to be revisited in the future using alternative molecular markers, my study provides morphological information for Discomonas gen. nov. that sheds light on the probable cellular identity of AY620316 and ‘Basal Group T’ . My study has shown morphological evidence for at least two previously uncharacterized clades belonging to the Tectofilosida and helps substantiate the establishment of the Thecofilosea, which currently contains Tectofilosida and Cryomonadida (Bass and Cavalier-Smith 2004).  However, the original diagnosis for Tectofilosida by Cavalier-Smith and Chao (2003) [i.e., uninucleate cell surrounded by an organic flexible tectum or rigid test with one or two apertures for filopodia, sometimes including foreign mineral particles (agglutinated); cilia or silica scales absent; tubular mitochondrial cristae]  51 should be amended by adding “flagella present” as both Ventrifissura and Verrucomonas possess flagella and are now shown to be members of the Tectofilosida.  2.4.6 Barcoding marine benthic cercozoans DNA sequences can be effectively used as a diagnostic tool for the identification of species, an approach known as “DNA barcoding” (Hebert et al. 2003a, b).  This general approach is starting to be used extensively for assessing the diversity of microeukaryotes having limited morphological details for species discrimination (Barth et al. 2006; Chantangsi et al. 2007; Lynn and Strüder-Kypke 2006; Saunders 2005; Scicluna et al. 2006). Accordingly, one primary aim of my study was to help demonstrate the potential of the 618- bp SSU rDNA sequence as DNA barcode for identifying marine benthic cercozoans and possibly other cercozoans.  Bass and Cavalier-Smith (2004) previously designed phylum- specific primers covering the 618-bp barcoding region for SSU rDNA, which facilitated PCR amplification of the gene sequences in the group of cercozoans I investigated here. In my analyses, different isolates of E. tripartita and P. obliqua showed 0 and 1 nucleotide differences in their 618-bp barcoding regions, respectively.  This result helped demonstrate the reliability of the DNA barcoding region for species identification.  By contrast, two different isolates of C. aestivalis showed 0.5% sequence divergence (3 nucleotide differences), which was equal to the divergence value between P. grandis and P. longipes (ex. Cryothecomonas longipes) (Table 2.2).  Therefore, if P. grandis and P. longipes constitute separate species as clearly shown by morphological and molecular evidence, then the two isolates of C. aestivalis could also be justifiably established as two (cryptic) species on the basis of this molecular marker.  It turns out that the possibility of cryptic species within C. aestivalis was already mentioned by Kühn et al. (2000) when they found a  52 relatively high number of nucleotide differences between two morphologically indistinguishable strains of this “species”.  These results, therefore, also demonstrate the utility of the DNA barcoding region for discovering and delimiting cryptic species. Moreover, because the 618-bp barcoding region gene produced very similar phylogenetic tree topologies as those derived from the full-length of the gene (Figure 2.4), this short barcoding region can also help systematists infer the broader genealogical relationships of cercozoans. The high copy number of the SSU rRNA gene in eukaryotic nuclear genomes facilitates the barcoding of isolated microeukaryotes even when very limited numbers of organisms can be obtained.  My study has shown that this approach is very useful for exploring the biodiversity of uncultured microeukaryotes; most of the SSU rDNA sequences reported here were derived from a few number of uncultured cells per DNA extraction, and in several cases, I acquired SSU rDNA sequences from only 1 uncultured cell that was manually isolated from the ocean.  My study not only highlighted the cryptic diversity of marine benthic cercozoans, which is extremely problematic for the non-specialist, but also demonstrated an alternative approach for species identification using the DNA barcoding principle.  Moreover, I provided morphological data for several cercozoan subgroups that were previously known only from environmental DNA surveys.  Overall, these data provide insights into the cellular identities of uncultured and undescribed cercozoans that help advance our understanding of cercozoan biodiversity in marine benthic ecosystems.  53 Table 2.1.  Oligonucleotide primers used for amplification and sequencing of SSU rDNA in this study.  Primers Direction Sequence 5'-3' Annealing region1 NPF12 Forward 5'-TGCGCTACCTGGTTGATCC-3' 1-19 PF1 Forward 5'-GCGCTACCTGGTTGATCCTGCC-3' 2-23 525F Forward 5'-AAGTCTGGTGCCAGCAGCC-3' 567-585 917FD2 Forward 5'-GCCAGAGGTGAAATTCTNGG-3' 917-936 1050F Forward  5'-GGGGGAGTATGGTCGCAAG-3' 1130-1148 1050FD2 Forward  5'-GGGGGAGTATGGTCGCRAG-3' 1130-1148 1050MRD2 Reverse 5'-GCCTYGCGACCATACTCC-3' 1150-1133 1134FD2 Forward 5'-CGCAAGGCTGAAACHTRAAGG-3' 1143-1163 nomet1134R Reverse 5'-TTTAAGTTTCAGCCTTGCG-3' 1161-1143 1242RD2 Reverse 5'-GTCYGGACCTGGTAAGTTTTC-3' 1242-1222 1250R  Reverse 5'-TAACGGAATTAACCAGACA-3' 1342-1324 1367RD2 Reverse 5'-TTTAGYAGGBCGAGGTCTCG-3' 1367-1348 R4 Reverse 5'-GATCCTTCTGCAGGTTCACCTAC-3' 1823-1801 FAD Reverse 5'-TGATCCTTCTGCAGGTTCACCTAC-3'  1824-1801  1 Annealing region was provided with reference to SSU rDNA sequence of Protaspis rotunda sp. nov. [GenBank accession no. FJ824123]. 2 Primers newly designed in this study.  54 Table 2.2.  Upper triangular matrix showing the number of nucleotide differences; lower triangular matrix showing the percentage of pairwise sequence divergences between small subunit rDNA sequences based on Kimura’s 2 parameter model.  Data are from 17 cercozoans, 11 of which were generated in this study.  The sequences were 618 bp in length.  [Cae1 = Cryothecomonas aestivalis (GenBank accession number AF290541); Cae2 = Cryothecomonas aestivalis (AF290539); Plon = Protaspis (ex. Cryothecomonas) longipes (AF290540); Pgra = Protaspis grandis (DQ303924); Pob1 = Protaspis obliqua isolate 1 (FJ824121); Pob2 = Protaspis obliqua isolate 2 (FJ824122); Prot = Protaspis rotunda (FJ824123); Poba = Protaspis obaniformis (FJ824124); Povi = Protaspis oviformis (FJ824125); Ebr1 = Ebria tripartita (DQ303922); Ebr2 = Ebria tripartita (DQ303923); Bben = Botuliforma benthica (FJ824126); Vart = Ventrifissura artocarpoidea (FJ824127); Vfol = Ventrifissura foliiformis (FJ824128); Vbif = Verrucomonas bifida (FJ824129); Vlon = Verrucomonas longifila (FJ824130); and Dret = Discomonas retusa (FJ824131)].     55  Taxa Cae1 Cae2 Plon Pgra Pob1 Pob2 Prot Poba Povi Ebr1 Ebr2 Bben Vart Vfol Vbif Vlon Dret Cae1 * 3 21 21 30 29 25 27 19 38 38 33 43 44 74 65 55 Cae2 0.5 * 18 18 27 26 22 24 16 35 35 30 42 41 71 64 52 Plon 3.55 3.03 * 3 21 20 4 7 2 32 32 28 41 38 70 65 50 Pgra 3.55 3.03 0.5 * 21 20 7 7 4 32 32 28 42 39 70 65 50 Pob1 5.13 4.61 3.54 3.54 * 1 25 25 19 37 37 34 46 45 69 66 48 Pob2 4.95 4.43 3.37 3.37 0.16 * 24 24 18 36 36 33 45 44 70 67 49 Prot 4.25 3.73 0.66 1.16 4.24 4.06 * 11 6 34 34 30 45 42 72 69 54 Poba 4.61 4.08 1.16 1.16 4.24 4.06 1.83 * 8 38 38 32 46 43 75 70 53 Povi 3.2 2.69 0.33 0.66 3.2 3.02 0.99 1.33 * 32 32 28 39 36 69 64 48 Ebr1 6.55 6.01 5.47 5.47 6.38 6.2 5.82 6.55 5.47 * 0 16 49 47 67 64 42 Ebr2 6.55 6.01 5.47 5.47 6.38 6.2 5.82 6.55 5.47 0 * 16 49 47 67 64 42 Bben 5.65 5.12 4.76 4.76 5.83 5.65 5.11 5.46 4.76 2.68 2.68 * 40 41 70 69 48 Vart 7.46 7.28 7.09 7.27 8.02 7.83 7.82 8.02 6.72 8.6 8.6 6.92 * 27 78 73 61 Vfol 7.62 7.08 6.53 6.71 7.81 7.63 7.25 7.44 6.17 8.18 8.18 7.08 4.59 * 76 72 54 Vbif 13.36 12.77 12.55 12.55 12.38 12.58 12.94 13.57 12.36 11.98 11.98 12.56 14.2 13.74 * 18 80 Vlon 11.59 11.41 11.59 11.59 11.81 12.01 12.38 12.6 11.4 11.42 11.42 12.38 13.19 12.95 3.03 * 77 Dret 9.7 9.14 8.75 8.75 8.38 8.57 9.5 9.33 8.38 7.28 7.28 8.38 10.85 9.48 14.56 13.98 * 55   56     Figure 2.1.  Light micrographs (LMs) of nine novel cercozoans found in this study.  (a) Protaspis rotunda sp. nov. showing a ventral view with an ovoid nucleus at the anterior right side of the cell and a curved slit in the middle of the cell.  (b) Protaspis obaniformis sp. nov. showing granulated cytoplasm, spherical nucleus in the middle of the cell, and a posterior ventral slit.  (c) Protaspis oviformis sp. nov. showing food particles with varying colors within its cytoplasm.  (d) Botuliforma benthica gen. et sp. nov. showing two anterior flagella, a large anterior nucleus, and a thick and rough cell wall.  (e) Ventrifissura artocarpoidea gen. et sp. nov. showing a cell with numerous pointed warts and an anterior spherical nucleus.  (f) Ventrifissura foliiformis gen. et sp. nov. showing a cell with a smooth cell surface, large food particle, and a small anterior nucleus.  (g) Verrucomonas bifida gen. et sp. nov. showing numerous warts of varying colors on the cell surface, a ventral furrow, a bilobed nucleus, and a notch at its posterior end.  (h) Verrucomonas longifila gen. et sp. nov. showing two flagella, an ovoid nucleus at the anterior of the cell, and several yellowish warts on cell surface.  (i) Discomonas retusa gen. et sp. nov. showing a prominent long anterior flagellum, a discoidal nucleus at the anterior of the cell, and numerous spherical granules distributed around cell periphery of the cell.  (Bars = 10 µm)       57      58 Figure 2.2.  Light micrographs (LMs) of the marine benthic cercozoans examined in this study.  (a) Protaspis grandis showing an ovoid nucleus (N) at the posterior of the cell, a ventral slit (arrowhead), and a thick cell wall (double arrowhead).  (b) Protaspis obliqua showing an anterior nucleus (N), a posterior notch (arrowhead), and a thick cell wall (double arrowhead).  (c) Protaspis rotunda sp. nov. showing two flagella (arrows) and a curved slit in the middle of the cell (arrowhead).  (d) Protaspis oviformis sp. nov. showing a posterior nucleus (N), a ventral slit (arrowhead), and an anterior protrusion (triple arrowhead).  (e) Protaspis oviformis sp. nov. showing a nucleus (N) in the middle of the cell, a thick cell wall (double arrowhead), and numerous ingested green food bodies within the cytoplasm.  (f) Ventrifissura foliiformis gen. et sp. nov. showing two flagella (arrows) and branched pseudopodia (arrowheads).  (g) Ventrifissura artocarpoidea gen. et sp. nov. showing a cell with numerous pointed warts and a ventral slit (arrowhead).  (h) Ventrifissura foliiformis gen. et sp. nov. showing a cell with a smooth cell surface, a large food particle, an anterior nucleus (N), and an anterior protrusion (triple arrowhead).  This image was taken through the dorsal side and focused at the ventral side of the cell.  (i) Verrucomonas bifida gen. et sp. nov. showing an anterior flagellum (arrow), numerous orange warts on the cell surface, and ejected extrusomes (arrowheads).  (j) Verrucomonas longifila gen. et sp. nov. showing the posterior flagellum (arrow), fine pseudopodia (double arrowheads), and ejected extrusomes (arrowheads).  (k) Botuliforma benthica gen. et sp. nov. showing one of two anterior flagella (arrow), ejected extrusomes (arrowheads), very fine and branched pseudopodia (double arrowheads), and a ventral groove (triple arrowhead).  (l) Discomonas retusa gen. et sp. nov. showing a prominent long anterior flagellum and a shorter posterior flagellum (arrows). (Bars = 10 µm)    59      60          Figure 2.3.  Diagrammatic line drawings of the nine novel cercozoans found in this study. (a) Protaspis rotunda sp. nov.  (b) Protaspis obaniformis sp. nov.  (c) Protaspis oviformis sp. nov.  (d) Botuliforma benthica gen. et sp. nov.  (e) Ventrifissura artocarpoidea gen. et sp. nov.  (f) Ventrifissura foliiformis gen. et sp. nov.  (g) Verrucomonas bifida gen. et sp. nov. (h) Verrucomonas longifila gen. et sp. nov.  (i) Discomonas retusa gen. et sp. nov.  (Bar = 10 µm)            61      62       Figure 2.4.  Comparison between Bayesian phylogenies inferred from the 1,617-bp full length (left; mean ln L = -11672.98) and 583-bp 5'-half barcoding region (right; mean ln L = - 4072.00) SSU rDNA sequence alignments of 35 cercozoan taxa; the phylogenetic positions of marine benthic cercozoan studied in this study are highlighted in black boxes.  Each tree is a consensus of 36,002 trees with the GTR+I+G using 4 rate categories implemented.  Two isolates of Protaspis obliqua were labeled as isolate 1 and 2.  Numbers of 0.50 or higher at the nodes indicate Bayesian posterior probabilities and PhyML bootstrap percentages higher than 50%.  Diamonds represent Bayesian posterior probability of 1.00 and PhyML bootstrap value of 100%.  The scale bar corresponds to 0.02 substitutions per site.  * Limnofila borokensis was previously misidentified as Gymnophrys cometa (AF411284) and Mesofila limnetica was previously referred to as Dimorpha-like sp. (AF411283).          63    64       Figure 2.5.  Bayesian phylogeny deduced from 923 bp of SSU rDNA sequences of 67 cercozoan taxa including several sequences derived from environmental studies; the phylogenetic positions of marine benthic cercozoan examined in this study are highlighted in black boxes.  The tree (mean ln L = -7530.23) is a consensus of 36,002 trees with the GTR+I+G using 4 rate categories implemented.  Numbers of 0.50 or higher at the notes indicate Bayesian posterior probabilities and PhyML bootstrap percentages higher than 50%. Black circles represent Bayesian posterior probability of 1.00.  Black diamonds represent Bayesian posterior probability of 1.00 and phyML bootstrap value of 100%.  The scale bar corresponds to 0.02 substitutions per site.  * Limnofila borokensis was previously misidentified as Gymnophrys cometa (AF411284) and Mesofila limnetica was previously referred to as Dimorpha-like sp. (AF411283).          65          66 2.5 References Al Qassab S, Lee WJ, Murray S, Patterson DJ (2002) Flagellates from stromatolites and surrounding sediments in Shark Bay, Western Australia. Acta Protozool 41: 91-144  Auer B, Arndt H (2001) Taxonomic composition and biomass of heterotrophic flagellates in relation to lake trophy and season. Freshwater Biol 46: 959-972  Barth D, Krenek S, Fokin SI, Berendonk TU (2006) Intraspecific genetic variation in Paramecium revealed by mitochondrial cytochrome c oxidase I sequences. J Eukaryot Microbiol 53: 20-25  Bass D, Cavalier-Smith T (2004) Phylum-specific environmental DNA analysis reveals remarkably high global biodiversity of Cercozoa (Protozoa). 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Biodivers Conserv 17: 243-259   70 CHAPTER 3: ULTRASTRUCTURE, LIFE CYCLE AND MOLECULAR PHYLOGENETIC POSITION OF A NOVEL MARINE SAND-DWELLING CERCOZOAN: CLAUTRIAVIA BIFLAGELLATA N. SP.*  3.1 Introduction Massart originally established Clautriavia in 1900 for gliding phagotrophic flagellates in interstitial environments with a non-metabolic cell, a single recurrent flagellum and a mid- ventral groove.  Since that time, only three species of Clautriavia have been described with light microscopy: C. mobilis Massart 1900 (the type species), C. parva Schouteden 1907, and C. cavus Lee and Patterson 2000.  The cell morphology and behavior of these flagellates is essentially indistinguishable from species of Protaspis (Cercozoa), except that members of the latter group possess two heterodynamic flagella rather than only one prominent recurrent flagellum (Hoppenrath and Leander 2006a).  Accordingly, Clautriavia has been interpreted to be descendents of Protaspis-like ancestors that have subsequently lost the anterior flagellum (Larsen and Patterson 1990).  However, the general morphological features of Clautriavia and Protaspis are also shared by several other very distantly related groups of eukaryotes living in the same environments, such as phagotrophic euglenids, cercozoans, dinoflagellates, and katablepharids; in fact, Clautriavia was once closely affiliated with euglenids based on the presence of paramylon-like granules within the cytoplasm (Walton 1915).  Because of this phylogenetic uncertainty and the very poor state of knowledge about this group, Clautriavia is currently treated as “eukaryotes of uncertain taxonomic affinity”.  * A version of this chapter has been accepted for publication.  Chantangsi C, Leander BS Ultrastructure, life cycle and molecular phylogenetic position of a novel marine sand-dwelling cercozoan: Clautriavia biflagellata n. sp. Protist   71 Ultrastructural data and comparative analyses of DNA sequences will help us to better understand the basic cellular organization, phylogenetic position, and evolutionary history of Clautriavia and the multitude of other heterotrophic flagellates thriving in marine interstitial environments.  In this vein, I discovered, isolated, and successfully cultivated a novel species of Clautriavia living in marine sand samples collected from the eastern Pacific Ocean.  I was then able to characterize the general ultrastructure, life cycle, and molecular phylogenetic position of this novel lineage using small subunit (SSU) rDNA sequences, scanning and transmission electron microscopy, and high-resolution light microscopy.  3.2 Materials and methods 3.2.1 Sample collection Sand samples were collected from a benthic natural habitat at Brady’s Beach, Bamfield, Vancouver Island, BC, Canada on 18 June, 2007.  Flagellates were extracted from the sand samples through a 48 µm mesh using a melted seawater-ice method described by Uhlig (1964).  Two to three spoons of sand samples were placed into an extraction column wrapped with a 48 µm mesh.  Seawater ice cubes were then put on top of the sand samples and left to melt over several hours.  The organisms of interest were separated through the mesh and concentrated in a Petri dish that was filled with seawater and placed underneath the extraction column.  3.2.2 Light microscopy (LM) The Petri dish containing the flagellates was then observed using a Leica DMIL inverted microscope.  Cultivated cells at different life history stages were individually isolated and placed on a slide for light microscopy using phase contrast and differential   72 interference contrast (DIC) microscopy with a Zeiss Axioplan 2 imaging microscope connected to a Leica DC500 color digital camera.  3.2.3 Culture establishment The diatom Navicula sp., which was observed under a microscope and found to be a food source for C. biflagellata n. sp. in natural samples, was isolated from the same samples described above.  Several isolated cells of Navicula sp. were inoculated in a 96-well plate containing 200 µl of f/2 medium (Guillard 1975; Guillard and Ryther 1962) without silica (Si) (f/2-Si: f/2 medium but omit Na2SiO3·9H2O) and exposed to natural sunlight.  After significant cell growth, cells of Navicula sp. were transferred into a 24-well plate containing 2 ml of f/2-Si medium.  Individually isolated cells of C. biflagellata n. sp. were then washed in f/2-Si medium two times and added to the well-plates containing a lawn of Navicula in 2 ml of f/2-Si medium.  A stable culture of C. biflagellata n. sp. was established at 18 °C under a light:dark cycle of 6h:18h with Navicula as a food source.  The type strain of C. biflagellata n. sp. is being maintained in the Leander Laboratory, Departments of Zoology and Botany, University of British Columbia, Canada.  The Navicula sp. (i.e., diatom food) was deposited in the American Type Culture Collection (ATCC; Manassas, Virginia, USA) as ATCC PRA- 314.  The duplicate of the C. biflagellata will be deposited in the ATCC.  3.2.4 Scanning electron microscopy (SEM) Cells of Clautriavia biflagellata were isolated and placed into a small container covered on one side with a 5-µm polycarbonate membrane filter (Corning Separations Div., Acton, MA, USA).  The samples were pre-fixed for 30 min at room temperature in the container with a buffered mixture of 8% glutaraldehyde, 4% OsO4, and sucrose, giving a   73 final concentration of 0.1 M sucrose in 2% glutaraldehyde and 1% OsO4.  The samples were then post-fixed for 30 min at room temperature with a couple drops of 4% OsO4 and washed three times in filtered seawater to remove the fixative.  Cells were dehydrated through a graded series of ethanol and critical point dried with CO2 using a Tousimis Samdri 795 CPD (Rockville, MD, USA).  Dried filters containing the cells were mounted on aluminum stubs and then sputter coated with gold (5 nm thickness) using a Cressington high-resolution sputter coater (Cressington Scientific Instruments Ltd, Watford, UK).  The coated cells were viewed under a Hitachi S4700 scanning electron microscope.  3.2.5 Transmission electron microscopy (TEM) Cultured cells of C. biflagellata n. sp. were pre-fixed for 1 h at room temperature in a final concentration of 2.5% (v/v) glutaraldehyde and 0.1 M sucrose in 0.1 M sodium cacodylate buffer (SCB).  Cell pellets were then washed three times in 0.2 M SCB for 5 min each.  Post-fixation of the cell pellets consisted of a final concentration of 1% (v/v) OsO4 in 0.15 M SCB for 1 h at room temperature.  Fixed cells were then washed two times in 0.2 M SCB and dehydrated through a graded series of ethanol: 30% for 60 min; 50% for 30 min; 70%, 85%, 95% for 15 min each; and four times in 100% for 15 min each.  The cells were then exposed to two 10 min exchanges in a transition fluid of 1 part 100% ethanol and 1 part acetone; and two 10 min exchanges in 100% acetone.  Infiltration was performed with acetone-Epon resin mixtures (acetone, 2:1 for 1 h, 1:1 overnight, 1:2 for 5 h, Epon 812 resin overnight).  Cell pellets were embedded in Epon 812 resin and polymerized at 65 ºC for 30 hrs.  The embedded cells were sectioned with a diamond knife on a Leica EM-UC6 ultramicrotome; sections were collected on copper, formvar-coated slot grids and stained with uranyl acid and lead citrate (Sato’s lead method) (Hanaichi et al. 1986; Sato 1968).   74 TEM micrographs were taken with Hitachi H7600 and FEI G20 LaB6 200kV transmission electron microscopes.  3.2.6 DNA extraction and PCR amplification Cells of C. biflagellata were individually isolated and washed three times in autoclaved filtered seawater.  DNA was extracted using the protocol provided in the Total Nucleic Acid Purification kit by EPICENTRE (Madison, WI, USA).  Polymerase chain reaction (PCR) with the final reaction volume of 25 µl was performed in two rounds in a thermal cycler using puReTaq Ready-To-Go PCR beads (GE Healthcare Bio-Sciences, Inc., Québec, Canada).  The first round of PCR was conducted using forward PF1 (5'- GCGCTACCTGGTTGATCCTGCC-3') and reverse R4 primers (5'- GATCCTTCTGCAGGTTCACCTAC-3').  A PCR band of 1,850-bp was gel-purified with the UltraClean™ 15 DNA Purification Kit (MO BIO Laboratories, Inc., CA, USA) and used as a template for two subsequent nested PCR experiments.  (1) The 5'-half of the SSU rRNA gene was amplified using the forward PF1 primer and the reverse primer “nomet1134R” (5'- TTTAAGTTTCAGCCTTGCG-3'); (2) The 3' half of the SSU rRNA gene was amplified using the forward primer “917FD” (5'-GCCAGAGGTGAAATTCTNGG-3') and the reverse R4 primer.  The thermal cycler was programmed as follows: hold at 94 oC for 4 min; 5 cycles of denaturation at 94 oC for 30 sec, annealing at 45 oC for 1 min, and extension at 72 oC for 105 sec; 35 cycles of denaturation at 94 oC for 30 sec, annealing at 55 oC for 1 min, and extension at 72 oC for 105 sec; and hold at 72 oC for 10 min.  PCR products corresponding to the expected sizes were separated by agarose gel electrophoresis and gel-purified as described previously.  The cleaned DNA was cloned into pCR2.1 vector using the TOPO TA Cloning® kit (Invitrogen Corporation, CA, USA).  Plasmids with the correct insert size were   75 sequenced using BigDye 3.1 and the vector forward and reverse primers with an Applied Biosystems 3730S 48-capillary sequencer.  The DNA sequence of the partial SSU rRNA gene was deposited into GenBank (accession number FJ919772).  3.2.7 Sequence alignment The SSU rRNA gene sequences were assembled and edited using SequencherTM (version 4.5, Gene Codes Corporation, Ann Arbor, Michigan, USA).  Acquired sequences were initially identified by Basic Local Alignment and Search Tool (BLAST) analysis.  The SSU rDNA sequence derived from C. biflagellata n. sp. was aligned using ClustalW (Thompson et al. 1994) implemented in the MEGA (Molecular Evolutionary Genetics Analysis) program version 4 (Tamura et al. 2007) and further refined by eye.  Two multiple sequence alignments were created for phylogenetic analyses: (1) a 69-taxon global alignment comprising sequences of representatives from all major eukaryotic groups (1,134 unambiguous sites: data not shown), and (2) a 36-taxon cercozoan alignment covering representatives from different cercozoan subgroups (1,625 unambiguous sites).  All ambiguous sites were excluded from the alignments prior to phylogenetic analyses.  All alignment files are available upon request.  3.2.8 Phylogenetic analyses MrBayes version 3.1.2 was used to perform Bayesian analyses on the two datasets (Ronquist and Huelsenbeck 2003).  Four Markov Chain Monte Carlo (MCMC) chains — 1 cold chain and 3 heated chains — were run for 2,000,000 generations, sampling every 50th generation (tree).  The first 4,000 trees were discarded as burn-in (convergence was   76 confirmed by eye).  The remaining trees were used to compute the 50% majority-rule consensus tree.  Branch lengths of the trees were saved. Maximum likelihood analysis was performed on the 36-taxon cercozoan alignment using PhyML (Guindon and Gascuel 2003).  The input tree was generated by BIONJ with optimization of topology, branch lengths, and rate parameters selected.  The General Time Reversible (GTR) model with eight substitution rate categories was chosen, and the proportion of invariable sites and gamma distribution parameter were estimated from the original dataset.  PhyML bootstrap trees with 100 bootstrap datasets were constructed using the same parameters described above.  3.2.9 Sequence availability   The SSU rDNA sequences included in the molecular phylogenetic analyses are available from GenBank under the following accession numbers: Allapsa vibrans (AF411265), Allas diplophysa (AF411262), Auranticordis quadriverberis (EU484393), Bodomorpha minima (AF411276), Botuliforma benthica (FJ824126), Cercomonas sp. AZ6 (AF411268), Cryothecomonas aestivalis (AF290539), Discomonas retusa (FJ824131), Ebria tripartita (DQ303922), Euglypha rotunda (AJ418784), Neoheteromita globosa (U42447), Lecythium sp. (AJ514867), Limnofila borokensis (previously misidentified as Gymnophrys cometa) (AF411284), Massisteria marina (AF174372), un-named sp. SA-M (AF411278), Mesofila limnetica (previously referred to as Dimorpha-like sp.) (AF411283), Paulinella chromatophora (X81811), Proleptomonas faecicola (AF411275), Protaspis obaniformis (FJ824124), Protaspis grandis (DQ303924), Protaspis longipes [formerly Cryothecomonas longipes] (AF290540), Protaspis obliqua isolate 1 (FJ824121), P. obliqua isolate 2 (FJ824122), Protaspis oviformis (FJ824125), Protaspis rotunda (FJ824123), Pseudodifflugia   77 cf. gracilis (AJ418794), Pseudopirsonia mucosa (AJ561116), un-named sp. SA-R (AF411279), Neoheteromita sp. AZ3 (AF411280), Thaumatomastix sp. (AF411261), Thaumatomonas seravini (AF411259), Ventrifissura artocarpoidea (FJ824127), Ventrifissura foliiformis (FJ824128), Verrucomonas bifida (FJ824129), and Verrucomonas longifila (FJ824130).  3.3 Results 3.3.1 General morphology and life cycle The cell shape of the Clautriavia isolate was circular to broadly ovate and was slightly concave ventrally, particularly near the flagellar insertion point (Figures 3.1A-C, 3.1E-G). Two recurrent flagella of unequal length emerged from the same flagellar pit positioned on the anterior side of a shallow ventral depression (Figures 3.1B, 3.1D, 3.1F-G).  The shorter flagellum was thin, inactive and inconspicuous; the longer flagellum was thicker and involved in gliding motility along substrates.  The short flagellum could only be observed with careful examination (Figures 3.1D, 3.1F-G).  The longer flagellum was about 2X the cell length and was vigorously motile when the cells were pipetted into the water column and during cell division (Figures 3.1B, 3.1E).  The cell surface of the Clautriavia isolate was covered with an interspersed distribution of minute pores (Figures 3.1E-H). Although a permanent oral or feeding apparatus was not present, the Clautriavia isolate fed on small diatoms and coccoid “green” algae through the ventral side of the cell (Figures 3.2D-E, 3.2G).  The formation of a common food vacuole was observed in the plasmodium stage (Figures 3.2D-E, 3.4A).  The emergence of pseudopodia for locomotion and feeding was never observed in the culture condition.  Reproduction was achieved by one of two possible methods depending on the density of prey cells in the culture as illustrated in Figure   78 3.6: (1) binary division of a uninucleated parent cell, producing two uninucleated daughter cells (Figures 3.2A-C, 3.3); and (2) production of large plasmodia (i.e., multinucleated cells with upwards of 20 nuclei) that subsequently divide multiple times to form several uninucleated daughter cells (Figure 3.2D-G, 3.4).  Binary fission of uninucleated parent cells occurred when prey cells in the culture dish were relatively scarce; the cleavage furrow formed along the mid-sagittal plane and proceeded from the anterior end of the cell toward the posterior end (Figures 3.2A-C, 3.3B-C).  Large multinucleated plasmodia generated by multiple nuclear divisions formed when prey cells in the culture dish were abundant. Locomotion of the plasmodium stage varied depending on its shape.  Flat plasmodia (Figure 3.2D) were capable of gliding along the substratum by means of flagella; large spherical or irregular plasmodia (Figure 3.2F) usually did not glide although flagellar beating was noticeble.  3.3.2 Main cytoplasmic components The cytoplasm of the Clautriavia isolate was generally colorless except for food vacuoles containing pigmented prey cells and for a few pigmented granules (Figures 3.1A-D, 3.2, 3.3B, 3.4A, 3.5F).  The cells also contained large lipid globules and numerous mitochondria with well-defined tubular cristae (Figures 3.1D, 3.3A-C, 3.4A-B, 3.4D, 3.5A, 3.5E-F).  The Clautriavia isolate lacked a cell wall of any kind and possessed a uniform layer of muciferous bodies immediately beneath the plasma membrane (Figures 3.2E, 3.3A-D, 3.4A-B, 3.4D).  TEM sections showed the cell surface with minute pores (Figures 3.3E, 3.4A-B) and demonstrated a highly vacuolated cytoplasm containing a prominent nucleus that was surrounded by a distinct layer of vesicles (Figures 3.5A-B).  The nucleus was in a close association with microbody of irregular shapes and, in some sections, surrounded by   79 this microbody (Figures 3.3A-B, 3.4A, 3.4C).  The nucleus was also in close proximity to a Golgi body complex (Figure 3.4C).  Although the nucleus did not contain conspicuously condensed chromosomes, euchromatin could be distinguished from heterochromatin and the nucleolus (Figures 3.3A-C, 3.4A, 3.5A-B).  The nucleus in the gliding cells was positioned immediately adjacent to the anterior flagellar pit (Figures 3.1B-D, 3.5B) and was connected to the two basal bodies by a prominent microtubular root (Figures 3.5B-D).  3.3.3 Molecular phylogenetic position Phylogenetic analyses of 1,134 unambiguous aligned sites from 69 SSU rDNA sequences, covering representatives from all major eukaryotic supergroups, demonstrated that the new Clautriavia isolate nested within the Cercozoa with very strong statistical support; the 12 cercozoan sequences included in this alignment branched together 100% of the time in both ML bootstrap analyses and Bayesian analyses (data not shown – see Methods).  A more comprehensive analysis consisting of an alignment of 1,625 homologous positions and 36 cercozoan SSU rDNA sequences, including representatives from the most relevant cercozoan subgroups, placed the new Clautriavia isolate near the clade consisting of Auranticordis quadriverberis (a free-living marine benthic tetraflagellate) and Pseudopirsonia mucosa (a tiny parasitic biflagellate of diatoms).  This relationship was recovered in all analyses of the complete dataset and received a Bayesian posterior probability of 0.96; this relationship was only weakly supported with ML bootstrap analyses (Figure 3.7).      80 3.3.4 Taxonomic descriptions Clautriavia Massart 1900, emend. Chantangsi et Leander, 2009 Diagnosis: Free-living, gliding, phagotrophic flagellates with one prominent recurrent flagellum that extends past the length of the cell and, sometimes, one very short recurrent flagellum which is difficult to detect.  The short recurrent flagellum, if present, sits within a ventral depression beneath a pit from which both flagella emerge.  Cells round, oval, or slightly oblong in shape, dorsoventrally flattened and with a mid-ventral groove.  Cell contains a uniform distribution of muciferous bodies immediately beneath the plasma membrane.  Cells are static in shape and capable of ingesting prey cells through the ventral side. Longitudinal binary fission occurs along the mid-sagittal plane.  Large cellular plasmodia consisting of three or more nuclei may be present during the life cycle when food is abundant; the plasmodia divide to produce several uninucleated daughter cells.  Clautriavia biflagellata Chantangsi et Leander, 2009 Hapantotype: Both resin-embedded cells used for TEM and cells on gold sputter- coated SEM stubs have been deposited in the Beaty Biodiversity Research Centre (Marine Invertebrate Collection) at the University of British Columbia, Vancouver, Canada. Iconotypes: Figures 3.1B, 3.1E-F, 3.2F, and 3.6A. Diagnosis: Cell is round, oval, or broadly ovate in outline; about 12-20 µm wide and 15-20 µm long.  Gliding cells are rigid and dorsoventrally flattened.  Two unequal flagella are directed posteriorly and emerge from a ventral subapical pit that is surrounded by a shallow ventral depression.  The shorter flagellum is inconspicuous, about 3 µm long, relatively thin, and is confined to the ventral depression; the longer recurrent flagellum is about 2X the cell length, thicker, and extends beyond the cell.  This long flagellum makes   81 contact with the substratum and is involved in gliding motility.  The cell surface is porous and without a cell wall or test.  A uniform layer of muciferous bodies is positioned immediately beneath the plasma membrane.  The nucleus with nucleolus is located at the anterior end of the cell and connected to the basal bodies by a prominent microtubular root. The cytoplasm is colorless except for the presence of food vacuoles containing pigmented prey cells.  Neither pseudopodia nor extrusomes were observed. DNA sequence: Small subunit rRNA gene sequence [GenBank accession no. FJ919772]. Type locality: Tidal sand-flat at Brady’s Beach (48°49'40'' N, 125°09'10'' W), Vancouver Island, British Columbia, Canada.  The specimen was found on June 18, 2007. Habitat: Marine sand. Etymology: The etymology for the specific epithet, Latin bi, two; L. flagellum, whip. The specific epithet reflects the presence of two flagella.  3.4 Discussion Molecular phylogenetic studies have shown that several heterotrophic flagellates previously treated as eukaryotes of uncertain taxomonic affinity fall within the Cercozoa, such as Allantion, Allas, Bodomorpha and Spongomonas (Cavalier-Smith 2000); Cryothecomonas (Kühn et al. 2000); Ebria (Hoppenrath and Leander 2006b); Gymnophrys and Lecythium (Nikolaev et al. 2003); Massisteria (Atkins et al. 2000); Metopion and Metromonas (Bass and Cavalier-Smith 2004); Proleptomonas (Vickerman et al. 2002); and Protaspis (Hoppenrath and Leander 2006a).  My study has established a new member of the Cercozoa, namely Clautriavia biflagellata n. sp., and provides evidence showing that it is the   82 nearest sister lineage to the Auranticordida clade [i.e., Auranticordis and Pseudopirsonia] rather than a close relative of Protaspis species.  3.4.1 Comparison of Clautriavia and Auranticordis Even though the general morphology of Auranticordis, Pseudopirsonia, and Clautriavia are very different from one another – i.e., A. quadriverberis is a large, multi- lobed, bright orange tetraflagellate that thrives within marine sand (Chantangsi et al. 2008), and P. mucosa is a tiny flagellate that parasitizes planktonic diatoms (Kühn et al. 1996), there are some significant similarities in these lineages at the ultrastructural level.  However, because ultrastructrual data are not known for P. mucosa, I must limit my comparisons to A. quadriverberis and C. biflagellata n. sp.  Both Clautriavia and Auranticordis possess an interspersed distribution of pores on the cell surface that are associated with a uniform layer of muciferous bodies positioned immediately underneath the plasma membrane.  This distinctive feature is most obvious in transmission electron micrographs and is also among the most conspicuous features of A. quadriverberis when viewed with light microscopy (Chantangsi et al. 2008); although the uniform distribution of muciferous bodies is not immediately obvious in light micrographs of C. biflagellata n. sp., they are detectable with careful examination of images taken at focal planes through the cell surface (e.g., Figures 3.2E, 3.3C).  Both lineages also possess a highly vacuolated cytoplasm and a prominent microtubular root that connects the anterior end of the nucleus with the flagellar basal bodies (see Figures 3.5B-D and Chantangsi et al. 2008).  In addition, my C. biflagellata possesses the nucleus surrounded by a unique microbody.  The microbody has been reported in several cercozoans, such as Bodomorpha (Mylnikov 1984), Cercomonas (Karpov et al. 2006), Cholamonas (Flavin et al. 2000), Heteromita (MacDonald et al. 1977), Katabia (Karpov et   83 al. 2003), and Massisteria (Patterson and Fenchel 1990).  The microbody in these taxa is elongated, irregular or sometime reticulated and is found around the nucleus and other positions in the cytoplasm (Myl'nikov and Karpov 2004).  By contrast, the microbody in Clautriavia is appressed to the nucleus (see Figures 3.4A, 3.4C) and was never observed elsewhere in the cytoplasm.  3.4.2 Comparison of Clautriavia and Protaspis Clautriavia and Protaspis are both benthic flagellates that possess a mid-ventral groove and glide along substrates with a prominent recurrent flagellum (Lee and Patterson 2000). The former is generally considered a uniflagellated eukaryote whereas the latter is obviously biflagellated (Massart 1900; Skuja 1939).  Clautriavia has been interpreted to be a Protaspis species with an anterior flagellum that has either been damaged, evolutionarily lost or simply overlooked (Lee and Patterson 2000).  Because two of the three species of Clautriavia were described over 100 years ago, namely C. mobilis Massart 1900 and C. parva Schouteden 1907, it is possible, and perhaps even expected, that an inconspicuous flagellum, if present in these Clautriavia species, was not detected with the microscopes utilized at that time. However, the third species of Clautriavia, namely C. cavus Lee and Patterson 2000, was described less than a decade ago as a flagellate with only one trailing flagellum; it is less likely that these authors would have overlooked a second shorter flagellum, if present. There are several other dissimilarities between Protaspis and C. biflagellata n. sp. For instance, although food vacuoles containing diatoms and coccoid “green” algae are obvious in C. biflagellata n. sp., the mechanism of phagocytosis does not appear to involve pseudopods emerging from a ventral slit like that found in Protaspis.  Moreover, C. biflagellata n. sp. performs asexual reproduction by longitudinal binary fission along the   84 mid-sagittal plane; in contrast, asexual division in Protaspis occurs along the frontal plane (Skuja 1948).  Clautriavia biflagellata n. sp. can also form large cellular plasmodia containing three or more nuclei that subsequently divide into individual uninucleated daughter cells; these plasmodia form when prey cells are profuse and presumably function to optimize asexual cell proliferation in favorable conditions.  Nonetheless, this life cycle stage has not been observed in any Protaspis species.  At the ultrastructural level, cells of Protaspis are surrounded by a thick multilayered cell wall that does not contain pores (Hoppenrath and Leander 2006a); by contrast, the cell surface of C. biflagellata n. sp. lacks a cell wall, and contains an interspersed distribution of pores.  Moreover, unlike that of Protaspis, C. biflagellata n. sp. does not contain nuclei with conspicuously condensed chromosomes nor a cytoplasm containing batteries of extrusomes (Hoppenrath and Leander 2006a). For over a century, both Clautriavia and Protaspis were treated as eukaryotes of uncertain phylogenetic position or tentatively lumped with other existing groups like euglenids or dinoflagellates.  Hoppenrath and Leander (2006a) recently demonstrated that Protaspis is a member of the Cercozoa, specifically within the Cryomonadida, using ultrastructural and molecular phylogenetic data.  My molecular phylogenetic data from C. biflagellata n. sp. suggest that this lineage is only distantly related to Protaspis and is instead more closely related to an emerging lineage of cercozoans consisting of Auranticordis, Pseudopirsonia, and relatives.  This molecular phylogenetic position is congruent with the ultrastructural characters of C. biflagellata n. sp. described above.      85 3.4.3 Emended diagnosis of Clautriavia Clautriavia was originally described as a gliding cell with one prominent recurrent flagellum.  Although this is generally consistent with the features of the isolate described here, this isolate actually possessed two recurrent flagella: a long prominent one used in gliding and a very short one that is difficult to detect with light microscopy.  The overall morphology and behavior of this isolate otherwise closely conforms to Clautriavia (e.g., cell shape and cell size, gliding motility, and a mid-ventral groove).  I anticipate that several close relatives of C. biflagellata n. sp. also possess an inconspicuous second flagellum of various lengths ranging from 0 to only a few microns.  Given this information, erecting a new genus name based on the relative length of an inconspicuous flagellum would be both impractical and uninformative.  Therefore, I have chosen to classify this isolate within Clautriavia rather than to erect a new genus.  Among the three previously described species of Clautriavia, C. biflagellata n. sp. is most similar to C. cavus.  I chose not to assign my novel isolate to C. cavus because C. biflagellata n. sp. was relatively larger in cell size and is the first member of the genus to be shown to have two recurrent flagella, rather than one.  Accordingly, I have emended the original description of this genus to include gliding flagellates with two flagella: one prominent recurrent flagellum that extends past the length of the cell and, if present, one very short recurrent flagellum that may be difficult to detect.        86      Figure 3.1.  Light and scanning electron micrographs (LMs and SEMs, respectively) of Clautriavia biflagellata n. sp. A-C. LMs showing the prominent recurrent flagellum (arrows), the shorter inactive flagellum (arrowhead), the flagellar pit (double arrowheads), the nucleus (N), and nucleolus (quadruple arrowhead).  D. LM of a flattened cell that more clearly demonstrates the prominent recurrent flagellum (arrow), the shorter inactive flagellum (arrowhead), the nucleus (N), and lipid globules (asterisks).  E. SEM showing a dorsal view of the cell and the prominent recurrent flagellum (arrow).  F-G. SEMs showing ventral views of the cell demonstrating the flagellar pit (double arrowheads), the prominent recurrent flagellum (arrows), and the shorter inactive flagellum (arrowheads) within a ventral depression (vd).  G. A close-up SEM showing two flagella (arrow and arrowhead) emerging from flagellar pit (double arrowhead) and its surrounding ventral depression area (vd).  H. A high magnification SEM showing an interspersed distribution of pores (arrowheads) on the cell surface.  (A-D, bar = 10 µm; E-F, bar = 5 µm; G-H, bar = 1 µm)         87          88       Figure 3.2.  Light micrographs (LMs) of Clautriavia biflagellata n. sp. showing different stages in the life cycle.  A-C. A series of LMs showing binary cell division (nucleus, N; flagellar pit, double arrowhead; prominent recurrent flagellum, arrows).  D-E. LMs showing the large multinucleated cell plasmodia that form when prey cells are abundant.  E. LM with the focal plane near the cell surface showing the uniform layer of muciferous bodies; note the granular appearance near the cell surface.  (nucleus, N; prominent recurrent flagellum, arrows; shorter inactive flagellum, arrowheads; flagellar pit, double arrowhead).  F. LM showing a large multinucleated and multilobed cell plasmodium.  G. LM of a flattened cell plasmodium that more clearly demonstrates multiple nuclei (arrowheads) and nucleoli (double arrowheads); the prominent recurrent flagella are indicated with arrows.  (A-G, bar = 10 µm)          89          90      Figure 3.3.  Light and transmission electron micrographs (LMs and TEMs, respectively) showing general ultrastructural features of Clautriavia biflagellata n. sp. during interphase and division.  A. TEM showing the general ultrastructural organization of an interphase cell (prominent recurrent flagellum, double arrowhead; microbody, triple arrowhead; the anterior nucleus, N; nucleolus, n; mitochondria, arrowheads; lipid droplets, L; and a uniform superficial layer of muciferous bodies, arrows).  B. TEM of a cell showing an elongated nucleus (N) surrounded by microbody (triple arrowhead), lipid droplets (L), an ingested diatom (d) within a food vacuole, and a uniform superficial layer of muciferous bodies, (arrows).  C. LM showing two nuclei (N) following mitosis and two prominent recurrent flagella (double arrowheads) that will segregate with each daughter cell.  Several lipid droplets (L) and muciferous bodies (arrows) were found distributed within the cytoplasm.  D. High magnification TEM of the cell surface showing the uniform layer of muciferous bodies (arrows) immediately beneath the plasma membrane.  E. High magnification TEM showing pores (arrows) on the cell surface.  (A-B, bar = 2 µm; C, bar = 10 µm; D-E, bar = 0.5 µm)        91               92      Figure 3.4.  Transmission electron micrographs (TEMs) of Clautriavia biflagellata n. sp. showing general ultrastructural features of the uninucleated gliding cells and the large multinucleated plasmodia.  A. Low magnification TEM through a large multinucleated plasmodium showing a highly vacuolated cytoplasm, several nuclei (N) surrounded by microbodies (quadruple arrowheads), mitochondria (m), duplicated flagellar pits (double arrowheads), flagella (arrowheads), lipid droplet (L), food vacuoles (fv) containing diatom prey cells (d), a superficial layer of muciferous bodies (arrows), and a pore (triple arrowhead).  B. A sagittal TEM through a gliding cell showing a highly vacuolated cytoplasm, lipid globules (L), a uniform layer of muciferous bodies below the cell surface (arrows), and a pore (triple arrowhead).  C. A high magnification TEM showing Golgi apparatus (arrow) located near the nucleus (N) and microbody (quadruple arrowhead) .  D. A high magnification TEM showing lipid globules (L), mitochondrion (m) with well-defined tubular cristae and a uniform layer of muciferous bodies below the cell surface (arrows).  (A, bar = 5 µm; B, bar = 2 µm; C-D, bar = 1 µm)        93          94      Figure 3.5.  High magnification transmission electron micrographs (TEMs) of Clautriavia biflagellata n. sp.  A-B. TEMs through the nucleus (N) showing lipid globules (L), a layer of vesicles around the nuclear envelope (arrows), nucleoli (n), intranuclear euchromatin (lighter) and heterochromatin (darker).  B. TEM showing two basal bodies (arrowheads) that are closely associated with the nucleus (N) via an electron dense zone.  The nucleus was surrounded by a distinct layer of vesicles (arrows).  C. TEM showing two flagella located within a flagellar pit (fp) and a prominent row of nine microtubules (arrowhead) positioned near the anterior end of the nucleus (N).  D. High magnification TEM of a dividing cell showing a duplicated flagellar apparatus consisting of a flagellar pit (fp) containing two flagella and prominent row of microtubules (arrowheads).  E. TEM showing a highly vacuolated cytoplasm and several mitochondria (arrowheads) with tubular cristae.  F. TEM showing lipid globules (L) and a food vacuole (fv) containing three degraded diatoms (d) positioned near the nucleus (N).  (A-B, bar = 2 µm; C, bar = 0.2 µm; D, bar = 0.5 µm; E, bar = 1 µm; F, bar = 2 µm)        95      96     Figure 3.6.  Illustration showing the main life cycle stages of Clautriavia biflagellata n. sp. A. Uninucleated (nucleus, N; nucleolus, n) cell with two unequal flagella that emerge from the same flagellar pit (fp); the flagellar pit is positioned on the anterior margin of a ventral depression (vd).  The prominent recurrent flagellum (rf) is used for gliding, while the shorter inconspicuous flagellum (sf) is confined to the ventral depression (vd). The cytoplasm contains lipid globules (l) and food vacuoles (fv) containing diatoms (d).  A layer of muciferous bodies (mb) is located immediately underneath the cell surface and is illustrated only in Figure 3.6A.  This structure is left out from other illustrations for clarity.  B. A pre- divisional cell with a duplicated flagellar apparatus and ventral depression.  C. A binucleated cell following mitosis.  D. A cell just before the complete cytokinesis along the mid-sagittal plane and the generation of two daughter cells.  The cycle represented by the arrows connecting A-D occurs when prey cells are scarce.  E-F. When prey cells are abundant, there is repeated replication of the nucleus and the flagellar apparatus resulting in trinucleated plasmodia (E) or multinucleated plasmodia (i.e., 4 or more nuclei) (F).  Uninucleated daughter cells are separated from the multinucleated plasmodia one at time and freely glide away along the substrate using their prominent recurrent flagellum (rf).       97          98     Figure 3.7.  A Bayesian phylogenetic tree topology inferred from 1,625 bp of SSU rDNA sequences from 36 cercozoan taxa.  The tree (mean ln L = -12383.82) is a consensus of 36,002 trees using a GTR+I+G+4 model.  Numbers on the branches indicate Bayesian posterior probabilities and PhyML bootstrap percentages higher than 0.50 or 50%, respectively.  Black circles represent Bayesian posterior probabilities of 1.00.  Black diamonds represent Bayesian posterior probabilities of 1.00 and ML bootstrap values of 100%.  The scale bar corresponds to 0.02 substitutions per site.  The dark box indicates the sequence of Clautriavia biflagellata n. sp. produced in this study.  * Allapsa vibrans was previously referred to as Allantion sp. (AF411265); Cercomonas sp. AZ6 was previously referred to as Cercomonas plasmodialis (AF411268); Limnofila borokensis was previously misidentified as Gymnophrys cometa (AF411284); Mesofila limnetica was previously referred to as Dimorpha-like sp. (AF411283); Neoheteromita globosa was previously known as Heteromita globosa (U42447); Neoheteromita sp. AZ3 was previously referred to as Spongomonas minima (AF411280); un-named sp. SA-M was previously referred to as Metopion-like sp. (AF411278); un-named sp. SA-R was previously referred to as Rigidomastix-like sp. (AF411279).       99         100 3.5 References Atkins MS, Teske AP, Anderson OR (2000) A survey of flagellate diversity at four deep- sea hydrothermal vents in the Eastern Pacific Ocean using structural and molecular approaches. 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Plenum Press, New York, USA, pp 26-60  Guillard RRL, Ryther JH (1962) Studies of marine planktonic diatoms. I. Cyclotella nana Hustedt and Detonula confervacea Cleve. Can J Microbiol 8: 229-239  Guindon S, Gascuel O (2003) PhyML - A simple, fast, and accurate algorithm to estimate large phylogenies by maximum likelihood. Syst Biol 52: 696-704  Hanaichi T, Sato T, Hoshino M, Mizuno N (1986) A stable lead stain by modification of Sato’s method. Proceedings of the XIth International Congress on Electron Microscopy, Japanese Society for Electron Microscopy, Kyoto, Japan, pp 2181-2182  Hoppenrath M, Leander BS (2006a) Dinoflagellate, euglenid or cercomonad? The ultrastructure and molecular phylogenetic position of Protaspis grandis n. sp. J Eukaryot Microbiol 53: 327-342  Hoppenrath M, Leander BS (2006b) Ebriid phylogeny and the expansion of the Cercozoa. 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Nucleic Acids Res 22: 4673-4680  Uhlig G (1964) Eine einfach Methode zur Extraktion der vagilen, mesopsammalen Mikrofauna. Helgol Wiss Meeresunters 11: 178-185  Vickerman K, Le Ray D, Hoef-Emden K, De Jonckheere J (2002) The soil flagellate Proleptomonas faecicola: cell organization and phylogeny suggest that the only described free-living trypanosomatid is not a kinetoplastid but has cercomonad affinities. Protist 153: 9- 24  Walton LB (1915) A review of the described species of the order Euglenoidina Bloch. class Flagellata (Protozoa) with particular refrence to those found in the city water supplies in other localities of Ohio. Ohio Biological Survey, Bulletin 4. Ohio State University Bulletin 19: 343-459   103 CHAPTER 4: MORPHOLOGY AND MOLECULAR PHYLOGENY OF A MARINE INTERSTITIAL TETRAFLAGELLATE WITH PUTATIVE ENDOSYMBIONTS: AURANTICORDIS QUADRIVERBERIS N. GEN. ET SP. (CERCOZOA)*  4.1 Introduction Marine benthic environments contain a diverse assortment of microorganisms that are still just beginning to be explored and characterized (Fenchel 1987; Hondeveld et al. 1992). The challenges associated with extracting and enumerating benthic microorganisms and the extreme variation of physical and chemical factors associated with the benthos have limited our understanding of these ecosystems (Hondeveld et al. 1992).  Nonetheless, both comparative morphological studies and environmental sequencing surveys have revealed a great deal of microeukaryotic diversity within the interstitial spaces of marine sediments (Al Qassab et al. 2002; Bass and Cavalier-Smith 2004; Bernard et al. 2000; Berney et al. 2004; Edgcomb et al. 2002; Hoppenrath and Leander 2006a; Larsen and Patterson 1990; Lee and Patterson 2000; Lee et al. 2003, 2005; López-García et al. 2003; Šlapeta et al. 2005; Stoeck et al. 2003; Vickerman et al. 2005).  The most conspicuous predatory flagellates in these habitats range from about 20-150 μm in size and fall into three major groups of eukaryotes that are very distantly related to one another: dinoflagellates, euglenids, and cercozoans. The Cercozoa is a large and diverse group of amoeboflagellates, with tubular mitochondrial cristae, that cluster together in molecular phylogenies inferred mainly from ribosomal gene sequences (small and large subunit rDNA) (Bass and Cavalier-Smith 2004; Bass et al. 2005; Cavalier-Smith 1998a, b; Cavalier-Smith and Chao 2003).  Although a  * A version of this chapter has been published.  Chantangsi C, Esson HJ, Leander BS (2008) Morphology and molecular phylogeny of a marine interstitial tetraflagellate with putative endosymbionts: Auranticordis quadriverberis n. gen. et sp. (Cercozoa). BMC Microbiol 8: 123   104 robust morphological synapomorphy is currently lacking for the group, members of the Cercozoa do share novel molecular traits (i.e., molecular synapomorphies), such as the insertion of one or two amino acid residues between the monomer tracks of highly conserved polyubiquitin genes (Bass et al. 2005).  Nonetheless, molecular phylogenetic studies have demonstrated that several enigmatic taxa, previously treated as Eukaryota insertae sedis, fall within the Cercozoa, such as Allantion, Allas, Bodomorpha and Spongomonas (Cavalier- Smith 2000); Cryothecomonas (Kühn et al. 2000); Ebria (Hoppenrath and Leander 2006b); Gymnophrys and Lecythium (Nikolaev et al. 2003); Massisteria (Atkins et al. 2000); Metopion and Metromonas (Bass and Cavalier-Smith 2004); Proleptomonas (Vickerman et al. 2002); and Protaspis (Hoppenrath and Leander 2006a).  Moreover, environmental sequencing surveys have demonstrated several cercozoan subclades without clear cellular identities, suggesting that the actual diversity of this group is composed of thousands of uncharacterized lineages (Bass and Cavalier-Smith 2004).  It must also be emphasized that morphological information from cercozoans, especially at the ultrastructural level, is largely absent from the literature.  Accordingly, I characterized the ultrastructure and molecular phylogeny of a highly unusual and rarely encountered tetraflagellate, Auranticordis quadriverberis n. gen. et sp. (Cercozoa), isolated from sand samples collected in a marine tidal flat.  Uncultured cells were individually isolated and prepared for DNA extraction (performed twice on different days, n = 5 and n = 1), transmission electron microscopy (TEM, n = 2) and scanning electron microscopy (SEM, n = 25).  This approach enabled us to describe the ultrastructure of intracellular pigmented bodies within A. quadriverberis that are most likely photosynthetic endosymbionts derived from cyanobacterial prey.     105 4.2 Materials and methods 4.2.1 Sampling and light microscopy (LM) Sand samples were collected from Spanish Banks, Vancouver, BC, Canada in March 2007.  Organisms were extracted from the sand samples through a 48 μm mesh using a melted seawater-ice method described by Uhlig (1964).  Briefly, 2-3 spoons of sand samples were placed into an extraction column wrapped with a 48 μm mesh.  Two to three seawater ice cubes were then put on top of the sand samples and left to melt over several hours.  The organisms of interest were separated through the mesh and concentrated in a Petri dish that was filled with seawater and placed underneath the extraction column.  The Petri dish containing the organisms was then screened using a Leica DMIL inverted microscope.  Cells were individually isolated and placed on a slide for light microscopy using phase contrast and differential interference contrast (DIC) microscopy with a Zeiss Axioplan 2 imaging microscope connected to a Leica DC500 color digital camera.  4.2.2 Scanning electron microscopy (SEM) Twenty-five cells of Auranticordis quadriverberis were individually isolated and placed into a small container covered on one side with a 10-μm polycarbonate membrane filter (Corning Separations Div., Acton, MA, USA).  The samples were pre-fixed in the container with OsO4 vapor for 30 min at room temperature and subsequently post-fixed for 30 min with a mixture of 8% glutaraldehyde and 4% OsO4, giving a final concentration of 2.5% glutaraldehyde and 1% OsO4.  The organisms were then washed three times in filtered seawater to remove the fixative and dehydrated through a graded series of ethanol. Dehydrated samples were critical point dried with CO2 using a Tousimis Samdri 795 CPD (Rockville, MD, USA).  Dried filters containing the cells were mounted on aluminum stubs   106 and then sputter coated with gold (5 nm thickness) using a Cressington high resolution sputter coater (Cressington Scientific Instruments Ltd, Watford, UK).  The coated cells were viewed under a Hitachi S4700 scanning electron microscope.  4.2.3 Transmission electron microscopy (TEM) Two individual cells of Auranticordis quadriverberis were prepared separately.  Each cell was pre-fixed with 2% (v/v) glutaraldehyde (in unbuffered seawater) at room temperature for 1 h.  Cells were then washed three times in filtered seawater and post-fixed with 1% (v/v) OsO4 (in unbuffered seawater) for another 1 h at room temperature.  Fixed cells were then washed three times in filtered seawater and were dehydrated through a graded series of ethanol.  Infiltration was performed with acetone-resin mixtures (acetone, 2:1, 1:1, 1:2, Epon 812 resin) and individually flat embedded in Epon 812 resin.  The resin containing the cell(s) was polymerized at 65°C for one day and sectioned with a diamond knife on a Leica EM-UC6 ultramicrotome.  The sections were collected on copper, formvar-coated slot grids and stained with uranyl acid and lead citrate (Sato's lead method) (Hanaichi et al. 1986; Sato 1968).  TEM micrographs were taken with a Hitachi H7600 transmission electron microscope.  4.2.4 DNA extraction and PCR amplification Five cells were individually isolated and washed three times in autoclaved seawater. DNA was extracted using the protocol provided in the Total Nucleic Acid Purification kit by EPICENTRE (Madison, WI, USA).  Polymerase chain reaction (PCR) was performed in a thermal cycler using puReTaq Ready-To-Go PCR beads (GE Healthcare Bio-Sciences, Inc., Québec, Canada).  The forward (PF1: 5'-GCGCTACCTGGTTGATCCTGCC-3') and reverse   107 (R4: 5'-GATCCTTCTGCAGGTTCACCTAC-3') primers for amplifying SSU rDNA were added into the tube with the final reaction volume of 25 μl.  The thermal cycler was programmed as follows: hold at 94 °C for 4 min; 5 cycles of denaturation at 94 °C for 30 sec, annealing at 45 °C for 1 min, and extension at 72 °C for 105 sec; 35 cycles of denaturation at 94°C for 30 sec, annealing at 55 °C for 1 min, and extension at 72 °C for 105 sec; and hold at 72 °C for 10 min.  PCR products corresponding to the expected size were separated by agarose gel electrophoresis and cleaned using the UltraClean™ 15 DNA Purification Kit (MO BIO Laboratories, Inc., CA, USA).  The cleaned DNA was cloned into pCR2.1 vector using the TOPO TA Cloning® kits (Invitrogen Corporation, CA, USA).  Plasmids with the correct insert size were sequenced using BigDye 3.1 and the vector forward and reverse primers, and an internal primer (525F: 5'-AAGTCTGGTGCCAGCAGCC-3') with an Applied Biosystems 3730S 48-capillary sequencer. The above processes were repeated on one additional cell of Auranticordis quadriverberis that was sampled and isolated at a different time, in order to assure authenticity of the obtained sequences. A total of two complete sequences of the SSU rDNA from two different isolates were deposited into GenBank [GenBank:EU484393 and GenBank:EU484394].  4.2.5 Sequence alignment and phylogenetic analyses Sequences were assembled and edited using Sequencher™ (version 4.5, Gene Codes Corporation, Ann Arbor, Michigan, USA).  Acquired sequences were initially identified by BLAST analysis.  New SSU rDNA sequences derived from two different isolates of Auranticordis quadriverberis were aligned with ClustalW (Thompson et al. 1994) using the MEGA (Molecular Evolutionary Genetics Analysis) program version 4 (Tamura et al. 2007)   108 and further refined by eye using MacClade (Maddison and Maddison 2000).  Three multiple sequence alignments were created: (1) a 69-taxon global alignment comprising sequences of representatives from all major eukaryotic groups (1,134 unambiguous sites: data not shown); (2) a 126-taxon cercozoan alignment consisting of cercozoan representatives and extensive environmental sequences (981 unambiguous sites: data not shown); and (3) a 32-taxon cercozoan alignment excluding the shorter and unrelated environmental sequences (1,526 unambiguous sites).  All gaps were excluded from the alignments prior to phylogenetic analyses.  The alignment files are available upon request. MrBayes version 3.1.2 was used to perform Bayesian analyses on all three datasets (Huelsenbeck and Ronquist 2001; Ronquist and Huelsenbeck 2003).  Two parallel runs were carried out on 2,000,000 generations with the four Markov Chain Monte Carlo (MCMC) chains – 1 cold chain and 3 heated chains – and sampling every 50th generation (tree).  The first 2,000 trees in each run were discarded as burn-in.  Branch lengths of the trees were saved. Maximum likelihood analyses were performed on all three datasets using PhyML (Guindon and Gascuel 2003).  Input trees for each dataset were generated by BIONJ with optimisation of topology, branch lengths, and rate parameters selected.  The General Time Reversible (GTR) model of nucleotide substitution was chosen.  The proportion of invariable sites and gamma distribution parameter were estimated from the original dataset.  Eight categories of substitution rates were selected. PhyML bootstrap trees with 100 bootstrap datasets were constructed using the same parameters as the individual ML trees.    109 4.2.6 Sequence availability The SSU rDNA nucleotide sequences included in 32-taxon analyses for this paper are available from the GenBank database under the following accession numbers: Allas diplophysa [GenBank:AF411262], Auranticordis quadriverberis [GenBank:EU484393 and GenBank:EU484394], Bodomorpha minima [GenBank:AF411276], Bodomorpha sp. [GenBank:DQ211596], Cercomonas plasmodialis [GenBank:AF411268], Cryothecomonas aestivalis [GenBank:AF290539], Dimorpha-like sp. [GenBank:AF411283], Ebria tripartita [GenBank:DQ303922], Euglypha rotunda [GenBank:AJ418784], Exuviaella pusilla [GenBank:DQ388459], Gymnophrys cometa [GenBank:AF411284], Heteromita globosa [GenBank:U42447], Lecythium sp. [GenBank:AJ514867], Massisteria marina [GenBank:AF174372], Metopion-like sp. [GenBank:AF411278], Paulinella chromatophora [GenBank:X81811], Proleptomonas faecicola [GenBank:AF411275], Protaspis grandis [GenBank:DQ303924], Pseudodifflugia cf. gracilis [GenBank:AJ418794], Pseudopirsonia mucosa [GenBank:AJ561116], Rigidomastix-like sp. [GenBank:AF411279], Spongomonas minima [GenBank:AF411280], Thaumatomastix sp. [GenBank:AF411261], thaumatomonadida environmental sample [GenBank:EF023494], Thaumatomonas coloniensis [GenBank:DQ211591], Thaumatomonas seravini [GenBank:AF411259], uncultured eukaryote [GenBank:AB252750], uncultured eukaryote [GenBank:AB252755], uncultured eukaryote [GenBank:AB252756], uncultured eukaryote [GenBank:AB275058], and uncultured marine eukaryote [GenBank:DQ369017].       110 4.3 Results 4.3.1 General morphology and behaviour Auranticordis quadriverberis was able to glide slowly using four tightly bundled flagella that were oriented posteriorly.  The cells of A. quadriverberis were also able to change shape, albeit only slightly, and could be prominently lobed, heart-shaped or ovoid (Figures 4.1A-F, 4.1H).  In general, the cells had a narrower anterior apex and an expanded posterior end and were composed of four major lobes (L): L1, L2, L3, and L4 (Figure 4.1A). L1 was smaller than other three lobes and was separated from L2, to the right, by a ventral depression (vd) and separated from L4, to the left, by a ventral groove (gr) that contained the four recurrent flagella (Figure 4.1A).  Apart from differences in cell shape and the effects of cell plasticity, there was also variation in the size of different individuals, ranging from 35-75 μm in diam. (n = 65).  The cells were conspicuously orange in color, caused mostly by the presence of linear arrays of tiny orange muciferous bodies that were distributed over the entire surface of the cell (Figures 4.1A-B).  Microscopic observations indicated that these bodies secrete sticky mucilage when the cells are disturbed, suggesting that the bodies function for adhesion to the substratum.  TEM micrographs showed that the muciferous bodies were small compartments (780 nm in diam.) positioned underneath the cell membrane and filled with amorphous material that was secreted as mucilaginous strands (Figures 4.2D, 4.3B-C).  The surface of A. quadriverberis was also corrugated and consisted of over 80 longitudinal ridges that spanned from the anterior apex to the posterior end (Figures 4.2A-C). The grooves between the ridges contained numerous tiny pores through which the mucilage from the muciferous bodies was secreted (Figure 4.2C).  TEM sections through the cell surface also demonstrated a single row of microtubules positioned beneath each ridge (Figure 4.3E).  No test or cell wall was present.   111 The four flagella of A. quadriverberis originated from an anterior flagellar pocket and nested tightly within the ventral groove, making them nearly invisible under the light microscope (Figures 4.1B, 4.1D-E, 4.1G, 4.2A-B, 4.2E, 4.3F, 4.4F).  Electron microscopy demonstrated that the flagella were arranged in two pairs and covered with flagellar hairs or mastigomenes (Figure 4.2E).  Except for very slight differences in length, all four flagella were morphologically identical and slightly longer than the cell (Figures 4.1B, 4.1D-E).  The flagella were also homodynamic and associated with gliding motility along the substratum. Pseudopodia were not observed.  4.3.2 Main cytoplasmic components Auranticordis quadriverberis contained a large nucleus (15-20 μm in diam.) situated in the anterior region of the cell (Figures 4.3A, 4.3D).  Although the position of the nucleus in living specimens cannot be readily seen under the light microscope, the nucleus is visible in compressed cells as a comparatively clear area (Figures 4.1G, 4.1I).  TEM sections demonstrated the nuclear envelope and a few prominent nucleoli (Figures 4.3A, 4.3D, 4.4D- E).  The nucleus was pointed at the anterior end and was connected to a striated band near the basal bodies and microtubular roots (Figures 4.3D, 4.4D-E).  Moreover, bundles of (non- microtubular) fibrous material were also observed within the cytoplasm near the cell periphery (Figure 4.4A). The cells of A. quadriverberis also contained an accumulation of black material near the anterior part of the cell, lipid globules and Golgi bodies (Figures 4.1C, 4.1F, 4.1I, 4.4B, 4.4H-I).  Although mitochondria with tubular cristae were not definitively observed, several elongated bodies that were highly reminiscent of acristate mitochondria were found near the periphery of the cell (Figures 4.3A, 4.4G).  The cells also contained 2-30 pale orange bodies   112 that were variable in shape and usually about 4-5 μm in diam.; however, some of these bodies were 14 μm long (Figures 4.1C, 4.1F, 4.1I, 4.3A, 4.5A-G, 4.6).  The pale orange bodies were distributed throughout the cell, but were most abundant in the anterior region of the cell. Each pale orange body was enveloped by two tightly pressed inner membranes and surrounded by sac-like vesicles (Figures 4.5A, 4.5C, 4.5F).  The innermost membrane invaginated into the lumen of the body and formed several unstacked thylakoids around the periphery (Figures 4.5A-C, 4.5E).  The sac-like vesicles occasionally butted together to form perpendicular partitions outside of the two inner membranes (Figure 4.5F).  The central core of the pale orange bodies was devoid of membranes and contained a central electron dense region containing tailed viral particles (Figures 4.5D, 4.5G).  4.3.3 Molecular phylogenetic position of Auranticordis Phylogenetic analyses of a 69-taxon dataset representing all major groups of eukaryotes showed A. quadriverberis branching within the Cercozoa with very strong statistical support (data not shown).  This cercozoan clade, comprised of Chlorarachnion reptans, Cryothecomonas aestivalis, C. longipes, Ebria tripartita, Euglypha rotunda, Heteromita globosa, and A. quadriverberis, was strongly supported in both maximum likelihood (ML) and Bayesian analyses (ML boostrap = 100 and Bayesian posterior probabilities = 1.00; data not shown).  A more comprehensive analysis of 981 homologous positions in 126 cercozoan SSU rDNA sequences, including several shorter environmental sequences, placed A. quadriverberis near Pseudopirsonia mucosa (a parasitic nanoflagellate of diatoms) and two unidentified cercozoans with 1.00 Bayesian posterior probabilities (data not shown). Accordingly, I performed phylogenetic analyses of 1,571 positions in 32 cercozoan taxa that   113 excluded the shortest environmental sequences and included the closest relatives of A. quadriverberis in the 126-taxon alignment. Figure 4.7 illustrates the phylogenetic analyses of the 32-taxon dataset.  Like in the analyses of 126 taxa, the two different isolates of A. quadriverberis clustered with two uncultured eukaryotes and P. mucosa (Figure 4.7).  A subclade consisting of A. quadriverberis, P. mucosa, and environmental sequence AB252755 was recovered with a posterior probability of 1.00 and 73% PhyML bootstrap value.  A more inclusive clade consisting of A. quadriverberis, P. mucosa and environmental sequences AB252755 and AB275058 received high statistical support (posterior probability of 1.00 and PhyML bootstrap value of 97%) (Figure 4.7).  Members of this clade also shared a derived molecular character within the context of 160 cercozoan sequences covering representatives from all known cercozoan subclades: namely, the substitution of cytosine (C) for thymine (T) at position 324 (with reference to the complete SSU rDNA sequence of Cercomonas sp.; GenBank accession no. AF411266, culture ATCC PRA-21) in Helix 12, based on the predicted secondary structure of the SSU rRNA gene in Palmaria palmata (Wuyts et al. 2000).  4.3.4 Taxonomic descriptions Taxonomic treatment for Auranticordis quadriverberis Phylum Cercozoa Cavalier-Smith 1998 Genus Auranticordis gen. nov. Chantangsi, Esson and Leander 2008 Diagnosis: Uninucleate tetraflagellates; four recurrent flagella inserted subapically and bundled together within a ventral longitudinal groove; all flagella about one cell length; cell shapes are prominently lobed, ovoid or heart-shaped; nucleus at anterior end of cell, with   114 nucleoli; no cell wall or test; minute orange muciferous bodies distributed in linear arrays over the entire cell; cytoplasm with pale orange pigmented bodies, usually concentrated at the anterior end; corrugated cell surface; black inclusions usually present at anterior part of the cell; locomotion by slow gliding; cell deformations possible; marine habitat. Type species: Auranticordis quadriverberis. Etymology: Latin aurantium, n. orange; L. cordis, n. heart.  The generic name reflects two characteristic features of this taxon: orange cell coloration and inverted heart-shaped cells.  Species Auranticordis quadriverberis spec. nov. Chantangsi, Esson and Leander 2008 Description: Cell shape ovoid, prominently lobed or inverted heart-shaped; cell size 35-75 μm long, 25-70 μm wide; four homodynamic flagella, inserted subapically and bundled within a ventral longitudinal groove; anterior nucleus with nucleoli; bright orange coloration caused by linear rows of minute orange muciferous bodies; corrugated cell surface with about 80 longitudinal ridges; no cell wall or test; cytoplasm with 2-30 pale orange pigmented bodies; black inclusions usually present at anterior part of the cell; locomotion by slow gliding. Small subunit rRNA gene sequences [GenBank:EU484393 and GenBank:EU484394]. Type locality: Tidal sand-flat at Spanish Banks, Vancouver, British Columbia, Canada. The specimen was found during March and May, 2007. Hapantotype: Both resin-embedded cells used for TEM and cells on gold sputter- coated SEM stubs have been deposited in the Beaty Biodiversity Research Centre (Marine Invertebrate Collection) at the University of British Columbia, Vancouver, Canada. Iconotype: Figures 4.1B, 4.1E-F, 4.1H, and 4.6.   115 Type locality: Spanish Banks, Vancouver, BC, Canada (39°28' N, 74°15' W). Habitat: Marine sand. Etymology: The etymology for the specific epithet, Latin quattuor, four; L. verberis, n. whip.  The specific epithet reflects the presence of four flagella.  4.4 Discussion 4.4.1 Comparative morphology The distinctly orange color of A. quadriverberis sets these flagellates apart from other organisms living in the same benthic environment.  To our knowledge, similar organisms have not been recorded previously (Al Qassab et al. 2002; Larsen and Patterson 1990; Lee and Patterson 2000; Lee et al. 2003, 2005; Vørs 1993); however, the orange color of A. quadriverberis is most reminiscent of the anoxic euglenozoan Calkinsia aureus (Lackey 1960). The presence of four recurrent flagella in A. quadriverberis is another distinctive feature.  Most cercozoans possess two flagella, although Cholamonas cyrtodiopsidis also has four flagella that are inserted subapically (Flavin et al. 2000; Myl'nikov and Karpov 2004). The flagella of C. cyrtodiopsidis form two symmetrical pairs comprising one long and one stubby flagellum (Flavin et al. 2000; Myl'nikov and Karpov 2004).  This flagellar organization differs from A. quadriverberis, which has two pairs of tightly bundled flagella originating from the same flagellar reservoir.  Cholamonas cyrtodiopsidis was assigned to the Cercomonadida due to possession of a microbody and kinetid architecture that is similar to some species of Cercomonas (Flavin et al. 2000; Myl'nikov and Karpov 2004).  Although both A. quadriverberis and C. cyrtodiopsidis possess four flagella, this character state is unlikely to be synapomorphic for these species: A. quadriverberis inhabits marine sand,   116 whereas C. cyrtodiopsidis inhabits the intestines of diopsid flies (Flavin et al. 2000). Moreover, the distinctive features present in one species tend not to be shared by the other (e.g., the paranuclear bodies found in C. cyrtodiopsidis are not present in A. quadriverberis). Because the phylogenetic position of C. cyrtodiopsidis has not yet been evaluated with molecular phylogenetic data, our ability to infer the evolution of the tetraflagellated state within the Cercozoa is limited. The flagella of A. quadriverberis are covered by hairs, and although this stands in contrast to the smooth flagella described in most other cercozoans, such as Cercomonas and Proleptomonas (Myl'nikov and Karpov 2004), the hairs could be homologous to those described in the predatory soil-dwelling flagellate Aurigamonas solis (Karpov 2000; Vickerman et al. 2005).  The four flagella of A. quadriverberis were also recurrent and homodynamic during gliding motility, which is unlike the heterodynamic flagella of most other interstitial cercozoans (e.g., Cercomonas, Heteromita, Katabia, Proleptomonas, and Protaspis) (Hoppenrath and Leander 2006a; Myl'nikov and Karpov 2004).  The gliding cells of A. quadriverberis were plastic and capable of slow changes in shape that was somewhat similar to that found in euglenids (Leander et al. 2007).  This plasticity is probably generated by the row of microtubules locating underneath the cell membrane (Figure 4.3E). The nucleus of A. quadriverberis is difficult to see in living cells, which is also unlike most other cercozoans (e.g., Aurigamonas, Cercomonas, Ebria, Euglypha, Heteromita, Protaspis, Thaumatomastix, and Thaumatomonas) (Ekelund et al. 2004; Hoppenrath and Leander 2006a, b; Lee and Patterson 2000; Vickerman et al. 2005).  The bloated shape of the cell and the dense distribution of minute orange muciferous bodies that subtend the entire surface of the cell obscured the nucleus.  The ultrastructure of the nucleus is similar to that of other cercozoans (e.g., contained several nucleoli) (Drebes et al. 1996; Hoppenrath and   117 Leander 2006a; Karpov et al. 2006; Thomsen et al. 1991; Vickerman et al. 2005); however, A. quadriverberis lacked permanently condensed chromosomes like those found in Cryothecomonas, Ebria, and Protaspis (Drebes et al. 1996; Hargraves 2002; Hoppenrath and Leander 2006a, b; Thomsen et al. 1991; Vickerman et al. 2005).  The shape of the nucleus in A. quadriverberis was indented at one side, a feature also noticed in the nucleus of Protaspis grandis (Hoppenrath and Leander 2006a), and had a prominent anterior projection oriented towards the flagellar pocket.  An anterior projection was also observed in the nucleus of Cercomonas; in both genera, the anterior projection was associated with a broad striated band and the ventral (posterior) roots of the anterior and posterior flagella (VP) (Karpov et al. 2006; Myl'nikov and Karpov 2004).  However, the characteristic microtubular cone present in Cercomonas (Karpov et al. 2006; Myl'nikov and Karpov 2004) was not observed in A. quadriverberis. The cytoplasm of A. quadriverberis contained lipid globules, Golgi bodies, and muciferous bodies.  The muciferous bodies were compartments organized in linear arrays and filled with an amorphous matrix that appeared bright orange under the light microscope. Extrusomes like these have also been reported in C. armigera as a minute peripheral concavities filled with a homogeneous matrix (Thomsen et al. 1991).  Other types of extrusomes that have been found in different cercozoan species, such as trichocysts, microtoxicysts, kinetocysts, and osmiophilic bodies (Hoppenrath and Leander 2006a; Karpov et al. 2006; Myl'nikov and Karpov 2004), were absent in A. quadriverberis.  The lipid globules varied considerably in size and were most abundant in the posterior region of A. quadriverberis.  These globules were reminiscent of those described in Protaspis (Hoppenrath and Leander 2006a).  Although the mode of feeding in A. quadriverberis was   118 not clearly observed, evidence of ingested bacteria was observed within its cytoplasm (Figure 4.4C). The cytoplasm of A. quadriverberis was highly vacuolated and looked similar to the cytoplasm described in Cryothecomonas armigera and Protaspis grandis (Hoppenrath and Leander 2006a; Thomsen et al. 1991).  The anterior part of the cell, however, contained black bodies similar to those that have been observed in other distantly related eukaryotes, such as some semi-anoxic euglenids and ciliates.  Moreover, distinct mitochondria with tubular cristae, which are characteristic of other cercozoans, were not found in A. quadriverberis. Putative mitochondria were, however, observed around the cell periphery (Figure 4.4G), and the lack of cristae in these organelles reflects either degenerate mitochondria associated with a low-oxygen environment or fixation artifact (Hackstein 2001).  The size of the putative mitochondria ranged between 135-185 nm long, which is smaller than the mitochondria described in most cercozoans.  For example, the mitochondria of Aurigamonas solis are about 630 nm (Vickerman et al. 2005), the mitochondria of Cercomonas are about 485 nm (Karpov et al. 2006), the mitochondria of Cryothecomonas longipes are about 280 nm (Schnepf and Kühn 2000), and the mitochondria of P. grandis are about 500 nm (Hoppenrath and Leander 2006a).  Although the implementation of fluorescent stains, like Mitotracker, could help establish the identity of these structures (Esseiva et al. 2004), this approach is limited by the scarcity of these organisms in natural environments and the unpredictability of finding them in the samples.  4.4.2 Putative primary endosymbionts Several light orange bodies about 4-14 μm in diam. were distributed within the cell and were especially abundant towards the anterior end of the cell.  Although the ultrastructure of   119 these pigmented bodies is novel, the presence of thylakoid-like membranes and a central space containing a densely stained inclusion is consistent with three possible identities that differ by the degree of integration with the host cell: (1) the bodies are ingested (photosynthetic) prey cells that are in the earliest stages of being degraded, (2) the bodies are transient photosynthetic endosymbionts that are continuously replenished by kleptoplasty, or (3) the bodies are permanently integrated photosynthetic endosymbionts (i.e., plastids).  The plausibility of each of these hypotheses is addressed below. The orange color of these bodies is reminiscent of the plastids in some microalgae, such as dinoflagellates and diatoms that occupy the same habitats as A. quadriverberis. However, neither dinoflagellate theca nor diatom frustules were found associated with these bodies in any TEM sections, and the ultrastructure of the bodies was very different from the known ultrastructural diversity in the plastids of diatoms and dinoflagellates.  Some cyanobacteria are known to have pale orange coloration that is similar to the orange bodies within A. quadriverberis (Graham and Wilcox 2000).  These orange bodies were surrounded by two tightly compressed inner membranes and sac-like vesicles.  Whereas typical food bodies show degrees of being digested by cellular enzymes, nearly all of the pigmented bodies observed were completely intact in all of the cells I observed (n = 70), suggesting that they are constant fixtures of the host cell cytoplasm. Primary endosymbiosis, involving a photosynthetic prokaryote within a eukaryotic cell, results in three surrounding membranes: two cyanobacterial inner membranes and a third, outer phagosomal membrane.  Green algae/land plants, red algae, and glaucophytes possess primary plastids (Archibald 2005; McFadden 2001; Palmer 2003).  Two membranes surround the plastids of green algae and red algae, and the third outer phagosomal membrane is inferred to have been lost (Archibald 2005; Keeling 2004; McFadden 2001; Palmer 2003).   120 Secondary endosymbiosis occurs through the engulfment, integration and maintenance of either a green or red alga by a predatory eukaryote.  This process produced the plastids of cryptomonads, haptophytes, stramenopiles, dinoflagellates, apicomplexans, and euglenids (Archibald 2005; McFadden 2001; Palmer 2003).  Two different lineages of cercozoans have independently acquired plastids through endosymbiosis: (1) chlorarachniophytes have secondary plastids derived from green algae (Gilson and McFadden 1995; Palmer 2003) and (2) Paulinella chromatophora has primary plastids derived from cyanobacterial prey (Kies 1974; Kies and Kremer 1979; Yoon et al. 2006). Like in Paulinella and the cyanelles of glaucophytes, the ultrastructure of the pigmented bodies within A. quadriverberis is most consistent with the ultrastructure of free- living cyanobacteria, suggesting an independent primary endosymbiotic origin (Bhattacharya et al. 1995, 2007; Kies 1974; Kies and Kremer 1979; McFadden 2001; Yoon et al. 2006). For instance, TEM sections through the pigmented bodies demonstrated a mode of division that is similar to division described in the cyanelles of Cyanophora paradoxa (Hall and Claus 1963) (Figure 4.5C).  Moreover, the thylakoids in the endosymbionts of P. chromatophora, the cyanelles of glaucophytes, and coccoid photosynthetic cyanobacteria are unstacked and arranged concentrically around the periphery of the cell (Jensen 1993; Kies 1974; Kies and Kremer 1990).  A similar arrangement was observed in the pigmented bodies of A. quadriverberis (Figures 4.5A-C), although the majority of the thylakoids projected inward towards the core of the body.  The central area within the pigmented bodies of A. quadriverberis resembled the pyrenoids in the cyanelles of Glaucocystis nostochinearum (Kies and Kremer 1990). The thylakoid-free core of the pigmented bodies also contained polygonal viral particles.  TEM sections through these particles demonstrated complete tailed phages similar   121 to those known to infect cyanobacteria (Clokie and Mann 2006; Dodge 1973; Padan and Shilo 1973) (Figure 4.5G).  Viral particles similar to those described in the pigmented bodies of A. quadriverberis have also been described in the same region in the plastids of other eukaryotes, such as the "polyhedral bodies" in the primary endosymbionts of P. chromatophora (Kies 1974) and the cyanelles of the glaucophyte Gloeochaete wittrockiana (Kies and Kremer 1990), and have also been found in the free-living photosynthetic cyanobacterium Nostoc punctiforme (Jensen 1993).  Two other important characters that have been used to infer a cyanobacterial origin for primary plastids are: (1) the presence of phycobilisomes and (2) the presence of a peptidoglycan wall (Bhattacharya et al. 1995; Kies 1974; Kies and Kremer 1979).  However, as previously mentioned, neither phycobilisomes nor a peptidoglycan layer was present in the orange bodies in A. quadriverberis.  4.5 Conclusion My characterization of A. quadriverberis n. gen. et sp. demonstrates several novel features within the Cercozoa, such as four homodynamic flagella, densely distributed linear rows of orange muciferous bodies, and putative endosymbionts with an enigmatic overall structure.  The discovery of this highly distinctive lineage underscores how poorly we understand the actual cellular diversity of cercozoans and, potentially, represents one of the few independent cases of primary endosymbiosis within the Cercozoa and beyond.  Although endosymbioses are known to have occurred many different times independently, the transformation of endosymbionts into organelles is considered to be much less common (Cavalier-Smith and Lee 1985).  In order to more confidently infer the origin of the pigmented bodies in A. quadriverberis, experiments involving autofluorescence and the amplification of plastid molecular markers (e.g., 16S rDNA and psb genes) could be   122 performed (Yoon et al. 2006).  These studies will be hampered mainly by the scarcity and unpredictability of finding these cells in natural samples.  Nonetheless, additional studies on A. quadriverberis and its putative endosymbionts will enable us to better understand the extent of endosymbiosis across the tree of eukaryotes and the convergent processes associated with the establishment and integration of endosymbionts within eukaryotic cells.                      123     Figure 4.1.  Light micrographs (LMs) of Auranticordis quadriverberis n. gen. et sp. showing cell color, main cytoplasmic components, and variation in cell shape.  A. Differential interference contrast (DIC) image focused on rows of longitudinally arranged orange muciferous bodies (arrowhead), the ventral groove (double arrowhead), lobe 1 (L1), a ventral depression (vd), L2, L3, and L4.  B. An inverted heart-shaped cell with visible flagella (arrow) emerging from the posterior region of the ventral groove.  C. A flattened cell showing larger pale orange bodies (putative primary endosymbionts, arrowheads) distributed in the anterior end of the cell.  D. DIC image showing the position of the ventral groove (double arrowhead) with flagella (arrow) relative to a prominent L1 and L4.  E. Phase contrast micrograph demonstrating the distal end of the flagella (arrow) emerging from the ventral groove.  F. DIC micrograph showing black bodies (asterisk) accumulated at the anterior end of the cell and two pale orange bodies (putative primary endosymbionts, arrowheads).  G. A squashed cell showing the anterior nucleus (N) and flagella (arrow).  H. DIC micrograph showing a cell with prominent lobes.  I. A squashed cell showing variation in the shape and size of the pale orange bodies (putative primary endosymbionts, arrowheads).  (A-I, Bar = 10 μm).       124          125        Figure 4.2.  Scanning electron micrographs (SEMs) of Auranticordis quadriverberis n. gen. et sp.  A. An anterior view of the cell showing the anterior apex (arrowhead), ventral groove (double arrowhead) and flagella (arrow) (Bar = 10 μm).  B. A higher magnification view of the anterior end of the cell (arrowhead) showing the flagella (arrow) within the ventral groove (double arrowhead) (Bar = 2 μm).  C. High magnification view of the ridges showing several tiny pores (arrowheads) in the grooves (Bar = 1 μm).  D. High magnification view of secreted mucus (arrowheads) (Bar = 0.5 μm).  E. High magnification view of the ventral groove showing all four flagella (arrows) bundled together and covered in hairs (Bar = 0.5 μm).           126           127     Figure 4.3.  Transmission electron micrographs (TEMs) of Auranticordis quadriverberis n. gen. et sp.  A. Low magnification view showing the main cellular components: black bodies (b), nucleus (N), pale orange bodies (putative primary endosymbionts, PE), a degraded PE (double arrowhead) surrounded by sac-like vesicles (asterisk), surface ridges (arrows), and the ventral depression (vd) (Bar = 10 μm).  B. Section through the surface showing a row of muciferous bodies (arrowheads) containing (orange) amorphous material.  Each muciferous body is about 500-900 nm in diameter (Bar = 0.5 μm).  C. High magnification view of muciferous bodies (arrowheads) and secreted mucus (arrow) (Bar = 0.5 μm).  D. Section through the anterior region of the cell showing black bodies (b), the flagellar pocket (double arrowhead), four flagella (arrows), a nucleolus (n), a pointed nucleus (N), and the ventral groove (gr) (Bar = 5 μm).  E. High magnification section through the surface ridges (arrow) showing underlying microtubules (arrowhead) and muciferous bodies (double arrowheads) (Bar = 0.5 μm).  An inset showing a magnified view of a surface ridge (arrow) with a row of microtubules underneath (arrowhead) (Bar = 0.5 μm).  F. Transverse section showing all four flagella within a flagellar pocket (arrowhead) near the nuclear anterior projection (N) (Bar = 1 μm).       128          129     Figure 4.4.  Transmission electron micrographs (TEMs) of Auranticordis quadriverberis n. gen. et sp., showing different cytoplasmic components.  A. High magnification TEM showing a vacuolated cytoplasm (arrowheads) and fibrous material (fs) distributed beneath the cell periphery (Bar = 0.5 μm).  B. High magnification view of the black inclusions (arrowheads) (Bar = 2 μm).  C. An ingested bacterium found within cytoplasm of A. quadriverberis (Bar = 0.25 μm).  D. A section through the nucleus (N) showing nucleoli (arrowheads) and an invaginated area (double arrowhead) (Bar = 2 μm).  E. High magnification TEM showing the nuclear envelope (arrow), the nucleus (N), and a striated band (double arrowhead) positioned between the nuclear tip and a microtubular root (arrowhead) (Bar = 0.5 μm).  F. Tangential section through the flagella (arrowheads) lying within the ventral groove (gr) (Bar = 1 μm).  G. A putative mitochondrion (arrowhead) positioned near the cell periphery (Bar = 0.2 μm).  An inset showing two putative mitochondria (Bar = 0.5 μm).  H. TEM showing lipid globules (lg) near the posterior part of the cell (Bar = 1 μm).  I. High magnification view of a Golgi apparatus (arrowhead) (Bar = 0.5 μm).        130          131     Figure 4.5.  Transmission electron micrographs (TEMs) showing the ultrastructure of putative primary endosymbionts in Auranticordis quadriverberis n. gen. et sp.  A. Low magnification TEM showing four putative endosymbionts, each surrounded by sac-like vesicles (sc) defined by an outer membrane (Bar = 2 μm).  B. High magnification TEM showing two enveloping inner membranes (arrowheads) and thylakoids (arrows) that are continuous with the innermost enveloping membrane (Bar = 0.2 μm).  C. TEM showing the thylakoids, the sac-like vesicle (sc), and a cleavage furrow indicative of division (arrowheads) (Bar = 0.5 μm).  D. High magnification TEM showing the central core of an endosymbiont containing viral particles (arrowheads) (Bar = 0.5 μm).  E. High magnification TEM showing a pronounced invagination of the innermost enveloping membrane (arrowhead) (Bar = 0.5 μm).  F. High magnification TEM showing the membrane (arrowheads) that defines the sac-like vesicle (sc) and the two innermost enveloping membranes (double arrowheads) (Bar = 0.2 μm).  G. TEM showing viral particles (arrowhead) consisting of a polygonal head and tail, and positioned within the core of an endosymbiont (Bar = 0.5 μm).  An inset showing a complete tailed viral particle (Bar = 0.2 μm).       132          133 Figure 4.6.  A schematic line drawing of Auranticordis quadriverberis n. gen. et sp.  The line drawing was constructed from light micrographs and showing a lobed cell, rows of tiny orange muciferous bodies (small circles), four flagella within ventral groove, a ventral depression (lightly stippled area to the left of the flagella), and four putative primary endosymbionts (large shaded circles).     134        Figure 4.7.  Maximum likelihood (ML) tree (-ln L = 10139.70214) inferred from 32 SSU rDNA sequences, 1,571 unambiguously aligned sites and a GTR+I+G+8 model of nucleotide substitutions.  Numbers above the branches denote PhyML bootstrap percentages, and numbers below the branches denote Bayesian posterior probabilities.  Black circles denote PhyML bootstrap percentages and posterior probabilities of 100% and 1.00, respectively. Line drawings were modified from the following sources: Auranticordis quadriverberis (this study) and Pseudopirsonia sp. (Kühn et al. 2004, © to and reproduced with the permission of Elsevier).  The asterisk next to sequence [GenBank:DQ388459] was derived from an environmental sequencing survey and was listed in GenBank as the dinoflagellate Exuviaella pusilla by Lin et al. (2006).          135         136 4.6 References Al Qassab S, Lee WJ, Murray S, Patterson DJ (2002) Flagellates from stromatolites and surrounding sediments in Shark Bay, Western Australia. 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Nonetheless, phylogenetic analyses of concatenated gene datasets have subsequently demonstrated that along with the Foraminifera and the Radiozoa (i.e., most of the traditional radiolarians), the Cercozoa is strongly nested within the supergroup Rhizaria (Adl et al. 2005; Burki et al. 2007, 2008; Keeling et al. 2005; Nikolaev et al. 2004; Polet et al. 2004). Cercozoans, along with euglenids and dinoflagellates, are among the most commonly encountered predatory flagellates in marine benthic environments, and environmental surveys of this diversity have demonstrated a huge number of cercozoan lineages that have yet to be adequately characterized (Bass and Cavalier-Smith 2004; Chantangsi et al. 2008; Hoppenrath and Leander 2006a, b).  Marine benthic cercozoans, especially members of the Cryomonadida (e.g., Cryothecomonas and Protaspis) and their benthic and planktonic relatives (e.g., Botuliforma, Clautriavia, Ebria, Thaumatomastix, Ventrifissura, and Verrucomonas), have so far been relatively neglected by the protistological community (Chantangsi et al. 2008).  Improved knowledge of these particular lineages is important for  * A version of this chapter has been submitted for publication.  Chantangsi C, Hoppenrath M, Leander BS Evolutionary relationships among marine cercozoans as inferred from combined SSU and LSU rDNA sequences and polyubiquitin insertions.   143 gaining a more comprehensive understanding of marine benthic ecosystems and the overall evolutionary history of cercozoans. The Cercozoa and the Foraminifera share a significant molecular character involving the presence of novel amino acid insertion(s) at the junctions between monomers of polyubiquitin genes (Archibald et al. 2003).  Ubiquitin is a small regulatory protein composed of 76 amino acids and functions to mark other proteins for destruction (Archibald et al. 2003; Bass et al. 2005).  This protein is found only in eukaryotes and plays essential roles in many biological processes, such as cell cycle regulation, DNA repair, transcriptional regulation, signal transduction, endocytosis, embryogenesis, and apoptosis (programmed cell death) (Hershko and Ciechanover 1998).  Ubiquitin genes are also highly conserved, as indicated by only three residue differences between humans and yeast, and can be configured in three main ways: (1) individual genes with single open reading frames; (2) genes fused to ribosomal protein genes; and (3) genes organized as linear head-to-tail ubiquitin coding region repeats, called polyubiquitin genes (Bass et al. 2005).  Although the amino acid sequences of polyubiquitin genes cannot be used directly to construct phylogenetic trees, the specific amino acids inserted between the monomers (e.g., serine or theonine) might provide molecular signatures for specific subclades within the Cercozoa and the Foraminifera (Archibald and Keeling 2004; Archibald et al. 2003; Bass et al. 2005). Phylogenetic analyses of multi-gene datasets have proven to be a powerful approach to better resolving relationships among eukaryotes (e.g., Baldauf et al. 2000; Bapteste et al. 2002; Burki and Pawlowski 2006; Burki et al. 2007, 2008; Harper et al. 2005; Kim et al. 2006; Rokas et al. 2003).  However, large-scale data (i.e., hundreds of genes or whole genomes) are only available for a very limited number of eukaryotes, and phylogenetic analyses of these datasets are computationally challenging, time consuming (Moreira et al.   144 2007), and currently impractical for studying uncultivated lineages.  Phylogenetic analyses of combined SSU and LSU rDNA sequences, on the other hand, have been shown to be a pragmatic and effective way to significantly improve the resolution of distant relationships within the tree of eukaryotes (e.g., Moreira et al. 2007).  These molecular markers are, therefore, expected to provide a promising avenue for evaluating the genealogical relationships within the Cercozoa, where most lineages need to be isolated from the ocean one cell at a time.  Accordingly, I amplified and sequenced LSU rDNA, SSU rRNA, and polyubiquitin genes from several uncultivated cercozoans collected from marine benthic habitats in order to better understand the biodiversity and evolutionary history of this group.  5.2 Materials and methods 5.2.1 Source of samples and light microscopy (LM) Fourteen cercozoan taxa were examined in this study (Table 5.1, Figure 5.1); only three of these have been cultivated: Cryothecomonas sp. (strain APCC MC5-1Cryo), Gymnochlora stellata (strain CCMP 2057), and Lotharella vacuolata (strain CCMP 240).  Placocista sp. was isolated from freshwater aquatic moss, and individual cells representing the remaining ten taxa were isolated from marine sand samples collected near Vancouver, British Columbia (details in Table 5.1).  Cercozoan cells were extracted from the sand samples through a 48 µm mesh using a melted seawater-ice method described by Uhlig (1964).  Briefly, 2-3 spoons of sand samples were placed into an extraction column wrapped with the mesh, and two to three seawater ice cubes were then put on top of the sand samples and left to melt over several hours.  The cells of interest passed through the mesh and were concentrated in a seawater-filled Petri dish that was placed underneath the extraction column.  The Petri dish containing the cells was then observed using a Leica DMIL inverted microscope.  Cells were   145 individually isolated and placed on a slide for imaging and identification using phase contrast and differential interference contrast (DIC) microscopy with a Zeiss Axioplan 2 imaging microscope (Figure 5.1).  5.2.2 DNA extraction and PCR amplification Cells were individually isolated and washed three times in either autoclaved distilled water or autoclaved filtered seawater, depending on the species.  DNA was extracted using the protocol provided in the Total Nucleic Acid Purification kit by EPICENTRE (Madison, WI, USA).  Semi-nested polymerase chain reaction (PCR) with a final reaction volume of 25 µl was performed in a thermal cycler using puReTaq Ready-To-Go PCR beads (GE Healthcare Bio-Sciences, Inc., Québec, Canada).  The first PCR amplification was conducted using the outermost forward and reverse primers which were NPF1 and R4 for SSU rDNA, 28S-1F and 28S-4R for LSU rDNA, and UBIQ1 and IUB2 for ubiquitin gene (Table 5.2). The first PCR product was used as a template for a second round of PCR involving the internal primers provided in Table 5.2.  For the amplification of SSU rDNA, the primers NPF1 and either 1242RD or CercR were used to obtain a 5' fragment of the gene; the overlapping 3' fragment was amplified with either 525F or Eugl_b and R4.  For the amplification of LSU rDNA, the 5' fragment of the gene was obtained using 28S-1F and either 28S-2R, 28S-1638R, 28S-1611R, or 28S-3R; the 3' fragment of the LSU rDNA was obtained using 28S-1426F and either 28S-3093R, 28S-3116R, or 28S-4R.  For some taxa, specific primers were designed in order to amplify the LSU rDNA: 28S-1499F for Ebria tripartita; 28S-1903R for Lotharella vacuolata; and 28S-PlacoR, 28S-PlacoR2, and 28S- PlacoR3 for Placocista sp.  Moreover, the specific sequencing primers used are also listed in Table 5.2.  In all cases, the primers UBIQ1 and IUB2 were used to amplify sections of the   146 polyubiquitin tract.  This primer pair generated a ladder of ubiquitin gene products ranging from a small fragment representing one half of a monomer (~120 bp) to increasing numbers of tandem repeats of the polyubiquitin tract that spanned the monomer-monomer junctions (e.g., ~340 bp representing 1.5 monomers and 570 bp representing 2.5 monomers). The thermal cycler was programmed as follows: hold at 94 oC for 4 min; 5 cycles of denaturation at 94 oC for 30 sec, annealing at 45 oC for 1 min, and extension at 72 oC for 105 sec; 35 cycles of denaturation at 94 oC for 30 sec, annealing at 50 oC for 1 min, and extension at 72 oC for 105 sec; and hold at 72 oC for 10 min.  PCR products corresponding to the expected size were separated by agarose gel electrophoresis and cleaned using the UltraClean™ 15 DNA Purification Kit (MO BIO Laboratories, Inc., CA, USA).  The cleaned DNA was cloned into pCR2.1 vector using the TOPO TA Cloning® kits (Invitrogen Corporation, CA, USA).  Plasmids with the correct insert size were sequenced using BigDye 3.1 and the vector primers, and appropriate internal primers, with an Applied Biosystems 3730S 48-capillary sequencer (Table 5.2).  5.2.3 Sequence alignment Sequences were assembled and edited using SequencherTM (version 4.5, Gene Codes Corporation, Ann Arbor, Michigan, USA).  Acquired sequences were initially identified by Basic Local Alignment and Search Tool (BLAST) analysis.  New sequences and other additional sequences retrieved from GenBank database were aligned using online MAFFT version 6 (Katoh et al. 2002).  The aligned sequences were then imported to the MEGA (Molecular Evolutionary Genetics Analysis) program version 4 (Tamura et al. 2007) and further refined by eye.  Three multiple sequence alignments were created for phylogenetic analyses: (1) an 85-taxon cercozoan SSU rDNA alignment covering representatives from   147 different cercozoan subgroups (1,443 sites); (2) a 25-taxon SSU rDNA alignment consisting of 23 cercozoan taxa and 2 radiozoans (1,415 sites); and (3) a 25-taxon combined SSU and LSU rDNA alignment consisting of 23 cercozoan taxa (ingroup) and two radiozoans (outgroup) (3,829 sites: 1,415 SSU and 2,414 LSU).  The last two alignments were each composed of the same composition of taxa.  SSU rDNA sequences from foraminiferans were excluded from the analyses because of their extremely high nucleotide substitution rates. The alignment files are available upon request.  5.2.4 Phylogenetic analyses MrBayes version 3.1.2 was used to perform Bayesian analyses on three datasets (Huelsenbeck and Ronquist 2001; Ronquist and Huelsenbeck 2003).  Four Markov Chain Monte Carlo (MCMC) chains — 1 cold chain and 3 heated chains — were run for 5,000,000 generations, sampling every 50th generation (tree).  The first 5,000 trees were discarded as burn-in (trees sampled before the likelihood plots reached a plateau).  The remaining trees were used to compute the 50% majority-rule consensus tree.  Branch lengths of the trees were saved. Maximum likelihood analyses were performed on the three datasets using PhyML (Guindon and Gascuel 2003).  The General Time Reversible (GTR) model of nucleotide substitution and optimization of equilibrium frequencies were chosen.  The proportion of invariable sites and gamma shape parameter were estimated from the original dataset.  Eight categories of substitution rates were selected.  PhyML bootstrap trees with 100 replicates for the 85-taxon and 25-taxon SSU rDNA datasets and 1,000 replicates for the 25-taxon combined SSU and LSU rDNA dataset were constructed using the same parameters as the individual ML trees.   148 5.2.5 Sequence availability The nucleotide sequences generated in this study were submitted to the GenBank database and listed in Table 5.1.  5.3 Results and discussion 5.3.1 Preliminary phylogeny of cercozoans as inferred from SSU rDNA sequences Phylogenetic analyses of the 85-taxon dataset (SSU rDNA) containing representatives of all major cercozoan lineages recovered three main clades: (1) a well supported clade containing parasitic cercozoans (i.e., Ascetosporea, Phagomyxida, and Plasmodiophorida) and free-living gromiids; (2) a clade consisting of chlorarachniophytes and Metromonas simplex, a free-living gliding flagellate; and (3) a large clade consisting of all other cercozoans in the analysis (Figure 5.2).  ML analyses grouped the large cercozoan clade and the chlorarachniophyte + Metromonas clade together with very strong statistical support (bootstrap value = 96% data not shown), to the exclusion of the clade consisting of the parasitic cercozoans and gromiids.  The SSU rDNA phylogenies also showed several well supported subclades, namely the Ascetosporea, Auranticordida, Chlorarachnea, Cryomonadida, Euglyphida, Glissomonadida, Gromiidea, Pansomonadida, Phaeodarea, Phagomyxida, Plasmodiophorida, Thaumatomonadida, Ventrifissuridae, Verrucomonadidae, some lineages of Cercomonadida, and a clade consisting of the Botuliformidae and the Ebriida (Figure 5.2).  These results are consistent with several previous studies (Bass and Cavalier-Smith 2004; Bass et al. 2005, 2009; Burki et al. 2002; Cavalier-Smith and Chao 2003; Chantangsi et al. 2008; Hoppenrath and Leander 2006a, b; Karpov et al. 2006; Polet et al. 2004; Vickerman et al. 2005; Wylezich et al. 2002, 2007).  However, several of the earliest cercozoan branches were only weakly or modestly supported by the data, and in some   149 cases, the statistical values for these branches were lower than those recovered in previous analyses (Bass and Cavalier-Smith 2004; Bass et al. 2005, 2009; Cavalier-Smith and Chao 2003).  My primary aim in this study was to assess and bolster the robustness of these relationships with the inclusion of additional taxa (i.e., several uncultivated marine and freshwater lineages shown in Table 5.1, Figure 5.1) and additional comparative molecular data for these taxa, namely LSU rDNA sequences and polyubiquitin insertions.  5.3.2 Phylogeny of mostly marine cercozoans as inferred from SSU+LSU rDNA sequences Phylogenetic analyses of the two 25-taxon datasets – one of only SSU rDNA sequences and one of combined SSU and LSU rDNA sequences – demonstrated two major clades within the ingroup (outgroup = radiozoans): a chlorarachniophyte clade and a clade consisting of all other cercozoans in the analysis (Figure 5.3).  The trees resulting from the analyses of the SSU rDNA alignment and the SSU+LSU rDNA alignment showed different topologies for some lineages, but these differences reflected very weak statistical support. For example, Clautriavia biflagellata was grouped with Paracercomonas marina in the 25- taxon SSU rDNA tree, whereas in the SSU+LSU rDNA tree, C. biflagellata branched as a sister taxon to Placocista sp. CC (Figure 5.3); the latter result is more consistent with the more comprehensive analyses of the 85-taxon alignment of SSU rDNA sequences (Figure 5.2).  Likewise, Ventrifissura foliiformis branched as the sister lineage to verrucomonads in the SSU+LSU rDNA tree but to Cercomonas sp. in the SSU rDNA tree (Figure 5.3). Although the precise phylogenetic positions of V. foliiformis and C. biflagellata remain unresolved, my analyses of both alignments did recover several well-supported subclades within the non-chlorarachniophyte cercozoans, including the Cryomonadida,   150 Thaumatomonadida, Verrucomonadidae, and the clade consisting of the Botuliformidae and Ebriida. Moreover, some relatively deep branches that received only modest statistical support in the SSU rDNA tree received stronger support in the analyses of the SSU+LSU rDNA alignment.  For instance, the large clade consisting of non-chlorarachniophyte cercozoans was recovered with a posterior probability (PP) of 0.87 and a bootstrap percentage (BP) of 86% in the SSU rDNA tree, but received PP = 1.00 and BP = 100% in the analyses of SSU+LSU rDNA (Figure 5.3).  I was also able to determine a more reliable phylogenetic position for some lineages in the combined analyses (SSU+LSU rDNA).  Chlorarachnion reptans, for example, clustered within the chlorarachniophytes without a clear internal position in the SSU rDNA trees, but in the combined analyses, this taxon was positioned as the sister lineage to the clade consisting of Bigelowiella and Norrisiella with modest to strong support (PP  1.00 and BP of 77) (Figure 5.3).  This relationship is consistent with the very recent study of chlorarachniophytes by Ota et al. (2009). These results help demonstrate the utility of combined SSU and LSU rDNA sequence alignments for inferring the evolutionary relationship within the Cercozoa.  It is widely appreciated that increased taxonomic sampling improves phylogenetic resolution (Graybeal 1998; Hillis 1996), and the relatively small number of ingroup taxa included in my combined analyses is a direct reflection of the limited number of LSU rDNA sequences currently available for cercozoans.  Accordingly, my study has almost tripled the taxonomic sampling of LSU rDNA sequences from non-chlorarachniophyte cercozoans, by providing 12 new sequences from mostly poorly understood lineages collected from marine habitats.  The high copy number of SSU and LSU rRNA genes in nuclear genomes facilitates the acquisition of these sequences via PCR from a very small amount of starting material.  The SSU and LSU   151 rDNA sequences reported here were derived mostly from uncultivated, albeit identified, marine microeukaryotes.  Fewer than five uncultured cells were isolated per DNA extraction, and in some cases, I acquired SSU and LSU rDNA sequences from only one uncultured cell that was manually isolated from heterogenous sand samples (Table 5.1, Figure 5.1).  This overall approach offers a pragmatic and effective way to evaluate the cellular and molecular biodiversity of cercozoans within a robust phylogenetic context.  5.3.3 Patterns of polyubiquitin insertions within the Cercozoa An insertion of one or two amino acid residues at the intermonomeric junction of polyubiquitin molecules has been found in both the Cercozoan and the Foraminifera, and this feature is, so far, absent in all other eukaryotes (Archibald et al. 2003; Bass et al. 2005) (Figure 5.4).  Because ubiquitin genes are so highly conserved across eukaryotes, the novel insertion is potentially a robust molecular marker for identifying members of the Cercozoa and specific subclades within the group.  This is significant, because the Cercozoa currently lacks unifying features at the morphological or behavioral levels.  Nonetheless, cercozoans clustered into three major clades in my phylogenetic analyses, and these clades reflect different patterns of polyubiquitin insertions.  For instance, chlorarachniophytes and members of the clade consisting of parasitic cercozoans and free-living gromiids possess one amino acid insertion (i.e., Alanine = A, Serine = S or Threonine = T); Metromonas possessed an insertion of two amino acids (i.e., serine plus glycine = SG); and members of the clade consisting of the remaining cercozoans possessed an insertion of either one amino acid (if one, all with serine = S) or two amino acids (most with SG) (Figure 5.2).  Paracercomonas marina and Massisteria marina possessed two uncommon amino acid insertions, namely SA (or SG) in the former and asparagine plus glycine = NG in the latter (Figure 5.2).   152 It has been suggested that the ancestral state for cercozoans possesses one amino acid insertion, like that in Foraminifera and chlorarachniophytes, and more derived cercozoans possess two residues between monomers of ubiquitin (Bass et al. 2005).  Archibald et al. (2003) and Bass et al. (2005) demonstrated polyubiquitin sequences for three foraminiferan genera — Bathysiphon, Haynesina, and Reticulomyxa — and showed that all of them possess only one amino acid insertion between the monomers, namely either A or T. Subsequent investigations have demonstrated that the following cercozoans also possess one amino acid insertion between the monomers of the polyubiquitin tract: Aurigamonas, Bigelowiella, Cercobodo, Gromia, Helkesimastix, Lotharella, Metopion, Plasmodiophora, and Spongospora (Archibald and Keeling 2004; Archibald et al. 2003; Bass et al. 2005; Vickerman et al. 2005).  My study expanded this dataset by demonstrating seven additional polyubiquitin sequences from mainly uncultivated marine cercozoans: Cryothecomonas sp. APCC, Ebria tripartita, Protaspis oviformis, and Protaspis grandis have an insertion consisting of serine; Clautriavia biflagellata, Ventrifissura sp., and Verrucomonas bifida have an insertion consisting of serine plus glycine (Figure 5.4).  The distribution of different states for the polyubiquitin insertions on my phylogenetic tree inferred from SSU rDNA sequences (Figures 5.2 and 5.3) shows that some lineages with one amino acid insertion are nested within lineages possessing two amino acid insertions.  For example, Aurigamonas solis (insertion = S) and Cercobodo agilis (insertion = S) formed a clade that was nested within a more inclusive, albeit weakly supported, clade consisting of glissomonads (insertion = SG) and cercomonads (insertion = SG) (Figure 5.2).  Moreover, members of the clade consisting of the Ebriida and the Cryomonadida were united by a consistent pattern of polyubiquitin insertions (i.e., one [S]) and were nested within a paraphyletic assembage of cercozoans (i.e., Cercomonas, Clautriavia, Mesofila,   153 Paracercomonas, Placocista, Thaumatomastix, Thaumatomonas, Ventrifissura, and Verrucomonas) that possessed a two-residue insertion of SG (Figures 5.3 and 5.4). My data indicate that the possession of one amino acid insertion between the monomers of the polyubiquitin tract is an ancestral state for some lineages (e.g., chlorarachniophytes and foraminiferans) and a derived state (a reversal) for other lineages (e.g., ebriids and cryomonads) that had ancestors with a two-residue insertion.  In other words, the gain and loss of amino acids within the polyubiquitin insertion is inferred to have happened multiple times independently within the Cercozoa.  If so, then the degree of evolutionary conservation of the polyubiquitin gene, specifically the insertions, might be lower than previously appreciated.  Although the overall heterogeneity in the size and states of the polyubiquitin insertions are becoming clearer with more data, these data also help reinforce our understanding of the universality of insertions within the Cercozoa; the complete absence of insertions has never been definitively observed in any cercozoan examined so far.  This suggests that the presence of the insertion is more functionally significant, and therefore more evolutionarily conserved, than the specific number and identity of the amino acids inserted (Archibald and Keeling 2004).  However, current data demonstrate that serine is a particularly common polyubiquitin insertion within the Cercozoa. Additional polyubiquitin sequences from both foraminiferans and cercozoans will continue to help establish the universality of the insertions and the utility of this molecular character for recognizing members of this extremely diverse assemblage of microeukaryotic diversity.  5.3.4 Patterns of polyubiquitin insertions within the Rhizaria The Rhizarian supergroup is comprised of not only the Cercozoa and the Foraminifera but also the Radiozoa (i.e., acanthareans and polycystines).  A putative subclade consisting of   154 the latter two groups is referred to as the Retaria and is based primarily on phylogenetic analyses of combined SSU and LSU rDNA datasets (Cavalier-Smith 1999, 2002; Moreira et al. 2007).  However, the sister relationships between the Foraminifera and the Radiozoa can be a result of long branch attraction artefact.  In addition, radiozoans appear to lack the novel polyubiquitin insertions found in cercozoans and foraminiferans (Archibald and Keeling 2004; Bass et al. 2005).  Because polyubiquitin gene sequences are extremely conserved across the full breadth of eukaryotic diversity, the presence of polyubiquitin insertions have been used as a robust molecular synapomorphy that, instead, unites the Cercozoa with the Foraminifera to the exclusion of the Radiozoa (Archibald and Keeling 2004; Archibald et al. 2003; Bass et al. 2005).  The Retaria hypothesis and the hypothesis that the Cercozoa and the Foraminifera are sister lineages are, thus, mutually exclusive.  Moreira et al. (2007) reconciled these conflicting data by proposing that the most recent ancestor of the Radiozoa subsequently lost the polyubiquitin insertions after their origin in the most recent ancestor of all rhizarians; in other words, the presence of polyubiquitin insertions in cercozoans and foraminiferans is inferred to be symplesiomorphic in both groups.  Although Moreira et al. (2007) provided good support for the Retaria hypothesis, the sister relationship between the Foraminifera and the Cercozoa cannot be ruled out at this time, especially in light of the high degree of evolutionary conservation known for polyubiquitin gene sequences.  Therefore, more comprehensive phylogenetic analyses involving more radiozoan taxa and more gene sequences will be required to more definitively establish the deepest relationships within the Rhizaria and the intriguing evolutionary history of the polyubiquitin gene.   155 Table 5.1.  Cercozoan protists whose genes were amplified and sequenced in this study.  Sequences derived from my study are underlined.  GenBank Accession Number Taxa Designations/Source of Organisms SSU LSU UBI Botuliforma benthica -/Pachena Beach, Vancouver Island, BC, Canada FJ824126 GQ144685 - Clautriavia biflagellata CC001/Brady Beach, Vancouver Island, BC, Canada FJ919772 GQ144682 GQ144694 Cryothecomonas sp. MC5-1Cryo/Antarctic Protist Culture Collection GQ144679 GQ144683 GQ144695 Ebria tripartita -/Bamfield Inlet, Vancouver Island, BC, Canada DQ303922 DQ303923 GQ144684 GQ144696 Gymnochlora stellata 2057/Center for Culture of Marine Phytoplankton AF076171 GQ144686 - Lotharella vacuolata 240/ Center for Culture of Marine Phytoplankton AF076168 GQ144687 - Placocista sp. CC/Grouse Mountain, Vancouver, BC, Canada GQ144680 GQ144688 - Protaspis grandis -/Boundary Bay, Vancouver, BC, Canada DQ303924 GQ144689 GQ144697 Protaspis oviformis -/Spanish Banks, Vancouver, BC, Canada FJ824125 - GQ144698 Thaumatomastix sp. CC002/Boundary Bay, Vancouver, BC, Canada GQ144681 GQ144693 - Ventrifissura foliiformis -/Boundary Bay, Vancouver, BC, Canada FJ824128 GQ144690 - Ventrifissura sp. CC005/Boundary Bay, Vancouver, BC, Canada - - GQ144699 Verrucomonas bifida -/Boundary Bay, Vancouver, BC, Canada FJ824129 GQ144691 GQ144700 Verrucomonas longifila -/Spanish Banks, Vancouver, BC, Canada FJ824130 GQ144692 -  155   156 Table 5.2.  Oligonucleotide primers used for amplification and sequencing in this study.  Primers Direction Sequence 5'-3' Annealing region1 SSU rDNA NPF1 Forward 5'-TGCGCTACCTGGTTGATCC-3' 1-19 525F Forward 5'-AAGTCTGGTGCCAGCAGCC-3' 568-586 Eugl_b Forward 5'-ACGACTCCATTGGCA-3' 1099-1085 1242RD Reverse 5'-GTCYGGACCTGGTAAGTTTTC-3' 1243-1223 CercR Reverse 5'-TCGAGGTCTCGTTCGTTAACGG-3' 1359-1338 R4 Reverse 5'-GATCCTTCTGCAGGTTCACCTAC-3' 1826-1804 LSU rDNA 28S-1F Forward 5'-ACCCGCTGAATTTAAGCAT-3' 1-19 28S-568F Forward 5'-TTGAAACACGGACCAAGGAG-3' 762-781 28S-713F2 Forward 5'-CTAACATATRTGCGAGTATTTG-3' 783-804 28S-1426F2 Forward 5'-AAYTAGCCCTGAAAATGGATGG-3' 1417-1438 28S-1499F2 Forward 5'-ATGAGTASGHGGGCGTG-3' 1490-1506 28S-2F Forward 5'-GCAGATCTTGGTGGTAG-3' 1557-1573 28S-2R2 Reverse 5'-CTMCCACCAAGATCYGC-3' 1573-1557 28S-1527F2 Forward 5'-CAAATGAGAACTTTGAAGACT-3' 1585-1605 28S-1638R2 Reverse 5'-CACRHGRAACCTTTCTCCACTTCA-3' 1628-1605 28S-PlacoR22 Reverse 5'-YTTCCCTATCTCTTAGGAY-3' 1719-1701 28S-1903R2  Reverse 5'-ACGTGARGTGCTTTACCAGC-3' 1759-1740 28S-PlacoR2 Reverse 5'-TCATCCGAAGACAACCTGC-3' 1760-1742 28S-1759F2 Forward 5'-GCCYGRGAAGAGTTA-3' 1795-1809 28S-1611R Reverse 5'-CTTGGASACCTGMTGCGG-3' 1986-1969 28S-3R Reverse 5'-CACCTTGGAGACCTGCT-3' 1989-1973 28S-PlacoR32 Reverse 5'-CATTGCGTCAACATCYTTTC-3' 2425-2406 28S-3093R2 Reverse 5'-CAATCCKACACTTGGCCYC-3' 3071-3053 28S-3116R2 Reverse 5'-CGTTCCCTGTTGGWGGA-3' 3092-3076 28S-4R Reverse 5'-TTCTGACTTAGAGGCGTTCAG-3' 3081-3061 Ubiquitin UBIQ1 Forward 5'-GGCCATGCARATHTTYGTNAARAC-3'  -4-20 IUB2 Reverse 5'-GATGCCYTCYTTRTCYTGDATYTT-3'  108-85  1 Annealing region was provided with reference to SSU rDNA sequence of Thaumatomastix sp. CC002 [GenBank accession no. GQ144681]; LSU rDNA sequence of Thaumatomastix sp. CC002 [GenBank accession no. GQ144693]; and ubiquitin gene sequence of Cryothecomonas sp. APCCMC5-1Cryo [GenBank accession no. GQ144695].  Annealing region of 28S-PlacoR, 28S-PlacoR2, and 28S-PlacoR3 was provided with reference to LSU rDNA sequence of Placocista sp. CC [GenBank accession no. GQ144688]; that of 28S- 1903R to LSU rDNA sequence of Lotharella vacuolata [GenBank accession no. GQ144687]; and that of 28S-4R to LSU rDNA sequence of Gymnochlora stellata [GenBank accession no. GQ144686]. 2 Primers that were newly designed in this study.   157         Figure 5.1.  Light micrographs (LMs) of the cercozoans examined in this study.  A. Botuliforma benthica.  B. Clautriavia biflagellata.  C. Cryothecomonas sp. (strain APCC MC5-1Cryo).  D. Ebria tripartita.  E. Gymnochlora stellata (strain CCMP 2057).  F. Lotharella vacuolata (strain CCMP 240).  G. Placocista sp. (CC-Grouse Mountain).  H. Protaspis grandis.  I. Protaspis oviformis.  J. Thaumatomastix sp. (CC002-Boundary Bay). K. Ventrifissura foliiformis.  L. Ventrifissura sp. (CC005-Boundary Bay).  M. Verrucomonas bifida.  N. Verrucomonas longifila.  (Bars = 10 µm)            158      159      Figure 5.2.  Phylogenetic tree inferred from Bayesian analysis of 1,443 bp of SSU rDNA sequences from 85 cercozoan taxa; the phylogenetic positions of the cercozoan sequences derived from this study are highlighted in black boxes.  The tree (mean ln L = -20900.61) is a consensus of 95,001 trees with the GTR+I+G using 8 rate categories implemented.  Numbers of 0.50 or higher at the notes indicate Bayesian posterior probabilities and PhyML bootstrap percentages higher than 50%.  Black circles represent Bayesian posterior probability of 1.00. Black diamonds represent Bayesian posterior probability of 1.00 and phyML bootstrap value of 100%.  Several long branches were shortened to one half (labeled 1/2) or one fourth (labeled 1/4) of their actual length.  The scale bar corresponds to 0.1 substitutions per site. Capital letters in square brackets indicate amino acid insertions of polyubiquitin molecules. Polyubiquitin-amino-acid insertions generated from this study are highlighted in black boxes. Black squares to the right of the tree represent the number of amino acid residues in the polyubiquitin insertion in the representative member(s) of the taxa indicated.         160    161       Figure 5.3.  Comparison of phylogenetic trees inferred from Bayesian analysis of a 25-taxon alignment consisting of 23 cercozoan taxa and 2 radiozoans; the upper tree was inferred from 1,415-bp of SSU rDNA sequences (mean ln L = -9998.43), and the lower tree was inferred from 3,829-bp of SSU + LSU rDNA sequences (mean ln L = -27661.69).  The phylogenetic positions of the cercozoan sequences derived from this study are highlighted in black boxes. Each tree is a consensus of 95,001 trees with the GTR+I+G using 8 rate categories implemented.  Numbers at the nodes indicate Bayesian posterior probabilities and PhyML bootstrap percentages.  Diamonds represent Bayesian posterior probability of 1.00 and PhyML bootstrap value of 100%.  Several long branches were shortened to one half (labeled 1/2) of their actual length.  The scale bar corresponds to 0.1 substitutions per site.  Black squares to the right of the tree represent the number of amino acid residues in the polyubiquitin insertion in the representative member(s) of the taxa indicated.         162     163          Figure 5.4.  Illustration showing the junction between two ubiquitin monomers (UBIQ) within the polyubiquitin tract of rhizarians (i.e., cercozoans, foraminiferans, and radiozoans) and an array of other eukaryotes.  Insertions of one or two amino acids between the ubiquitin monomers are marked in bold.  The seven cercozoan sequences generated by this study are highlighted in black boxes.             164    165 5.4 References Adl SM, Simpson AG, Farmer MA, Andersen RA, Anderson OR, Barta JR, Bowser SS, Brugerolle G, Fensome RA, Fredericq S, James TY, Karpov S, Kugrens P, Krug J, Lane CE, Lewis LA, Lodge J, Lynn DH, Mann DG, McCourt RM, Mendoza L, Moestrup O, Mozley-Standridge SE, Nerad TA, Shearer CA, Smirnov AV, Spiegel FW, Taylor MF (2005) The new higher level classification of eukaryotes with emphasis on the taxonomy of protists. J Eukaryot Microbiol 52: 399-451  Archibald JM, Keeling PJ (2004) Actin and ubiquitin protein sequences support a cercozoan/foraminieran ancestry for the plasmodiophorid plant pathogens. J Eukaryot Microbiol 51: 113-118  Archibald JM, Longet D, Pawlowski J, Keeling PJ (2003) A novel polyubiquitin structure in Cercozoa and Foraminifera: evidence for a new eukaryotic supergroup. Mol Biol Evol 20: 62-66  Baldauf SL, Roger AJ, Wenk-Siefert I, Doolittle WF (2000) “A kingdom level phylogeny of eukayotes based on combined protein data”. Science 290: 972-977  Bapteste E, Brinkmann H, Lee JA, Moore DV, Sensen CW, Gordon P, Duruflé L, Gaasterland T, Lopez P, Müller M, Philippe H (2002) The analysis of 100 genes supports the grouping of three highly divergent amoebae: Dictyostelium, Entamoeba, and Mastigamoeba. Proc Natl Acad Sci U S A 99: 1414-1419  Bass D, Cavalier-Smith T (2004) Phylum-specific environmental DNA analysis reveals remarkably high global biodiversity of Cercozoa (Protozoa). Int J Syst Evol Microbiol 54: 2393-2404  Bass D, Moreira D, López-García P, Polet S, Chao EE, von der Heyden S, Pawlowski J, Cavalier-Smith T (2005) Polyubiquitin insertions and the phylogeny of Cercozoa and Rhizaria. Protist 156: 149-161  Bass D, Chao EE, Nikolaev S, Yabuki A, Ishida K, Berney C, Pakzad U, Wylezich C, Cavalier-Smith T (2009) Phylogeny of novel naked Filose and Reticulose Cercozoa: Granofilosea cl. n. and Proteomyxidea revised. Protist 160: 75-109  Burki F, Berney C, Pawlowski J (2002) Phylogenetic position of Gromia oviformis Dujardin inferred from nuclear-encoded small subunit ribosomal DNA. Protist 153: 251-260  Burki F, Pawlowski J (2006) Monophyly of Rhizaria and multigene phylogeny of unicellular bikonts. Mol Biol Evol 23: 1922-1930  Burki F, Shalchian-Tabrizi K, Pawlowski J (2008) Phylogenomics reveals a new 'megagroup' including most photosynthetic eukaryotes. Biol Lett 4: 366-369   166 Burki F, Shalchian-Tabrizi K, Minge M, Skjaeveland A, Nikolaev SI, Jakobsen KS, Pawlowski J (2007) Phylogenomics reshuffles the eukaryotic supergroups. PLoS ONE 2: e790  Cavalier-Smith T (1998a) A revised six-kingdom system of life. Biol Rev Camb Philos Soc 73: 203-266  Cavalier-Smith T (1998b) Neomonada and the origin of animals and fungi. In Coombs GH, Vickerman K, Sleigh MA, Warren A (eds) Evolutionary relationships among protozoa. Kluwer Academic Publishers, London, pp 375-407  Cavalier-Smith T (1999) Principles of protein and lipid targeting in secondary symbiogenesis: euglenoid, dinoflagellate, and sporozoan plastid origins and the eukaryote family tree. J Eukaryot Microbiol 46: 347-366  Cavalier-Smith T (2002) The phagotrophic origin of eukaryotes and phylogenetic classification of Protozoa. Int J Syst Evol Microbiol 52: 297-354  Cavalier-Smith T, Chao EE (2003) Phylogeny and classification of Phylum Cercozoa (Protozoa). Protist 154: 341-358  Chantangsi C, Esson HJ, Leander BS (2008) Morphology and molecular phylogeny of a marine interstitial tetraflagellate with putative endosymbionts: Auranticordis quadriverberis n. gen. et sp. (Cercozoa). BMC Microbiol 8: 123  Graybeal A (1998) Is it better to add taxa or characters to a difficult phylogenetic problem? Syst Biol 47: 9-17  Guindon S, Gascuel O (2003) PhyML - A simple, fast, and accurate algorithm to estimate large phylogenies by maximum likelihood. Syst Biol 52: 696-704  Harper JT, Waanders E, Keeling PJ (2005) On the monophyly of chromalveolates using a six-protein phylogeny of eukaryotes. Int J Syst Evol Microbiol 55: 487-496  Hershko A, Ciechanover A (1998) The ubiquitin system. Annu Rev Biochem 67: 425-479  Hillis DM (1996) Inferring complex phylogenies. Nature 383: 130-131  Hoppenrath M, Leander BS (2006a) Dinoflagellate, euglenid or cercomonad? The ultrastructure and molecular phylogenetic position of Protaspis grandis n. sp. 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Mol Biol Evol 23: 2455-2466  Moreira D, von der Heyden S, Bass D, Lopez-Garcia P, Chao E, Cavalier-Smith T (2007) Global eukaryote phylogeny: combined small- and large-subunit ribosomal DNA trees support monophyly of Rhizaria, Retaria and Excavata. Mol Phylogenet Evol 44: 255- 266  Nikolaev SI, Berney C, Fahrni JF, Bolivar I, Polet S, Mylnikov AP, Aleshin VV, Petrov NB, Pawlowski J (2004) The twilight of Heliozoa and rise of Rhizaria, an emerging supergroup of amoeboid eukaryotes. Proc Natl Acad Sci U S A 101: 8066-8071  Ota S, Vaulot D, Le Gall F, Yabuki A, Ishida K (2009) Partenskyella glossopodia gen. et sp. nov., the first report of a Chlorarachniophyte that lacks a pyrenoid. Protist 160: 137-150  Polet S, Berney C, Fahrni J, Pawlowski J (2004) Small-subunit ribosomal RNA gene sequences of Phaeodarea challenge the monophyly of Haeckel's Radiolaria. 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Protist 156: 335-354    168 Wylezich C, Meisterfeld R, Meisterfeld S, Schlegel M (2002) Phylogenetic analyses of small subunit ribosomal RNA coding regions reveal a monophyletic lineage of euglyphid testate amoebae (Order Euglyphida). J Eukaryot Microbiol 49: 108-118  Wylezich C, Mylnikov AP, Weitere M, Arndt H (2007) Distribution and phylogenetic relationships of freshwater Thaumatomonads with a description of the new species Thaumatomonas coloniensis n. sp. J Eukaryot Microbiol 54: 347-357                       169 CHAPTER 6: CONCLUSION  6.1 Cryptic biodiversity of marine interstitial benthic cercozoans Environmental DNA surveys have shown a great deal of hidden diversity within the Cercozoa (Bass and Cavalier-Smith 2004; Bass et al. 2009; Brad et al. 2008; Chen et al. 2008; Massana and Pedrós-Alió 2008; Park et al. 2008; Piquet et al. 2008; Šlapeta et al. 2005; Tian et al. 2009).  My investigations into the biodiversity of heterotrophic flagellates in marine benthic habitats in British Columbia, Canada have demonstrated several undescribed taxa with morphological features that resemble the cercozoan genera Cryothecomonas and Protaspis.  Phylogenetic analyses of small subunit rDNA sequences derived from two uncultured isolates of Protaspis obliqua and nine novel cercozoan species (within four novel genera) provided organismal anchors that helped establish the cellular identities of several different environmental sequence clades (Chapter 2).  Although this investigation underscored the immense diversity of marine benthic cercozoans awaiting our discovery, these data also showed that the rarity of distinctive morphological features in cryomonads, and other groups of cercozoans, makes the identification and systematics of the group very difficult.  Therefore, an improved understanding of cercozoan biodiversity will benefit from an alternative approach for species identification — the DNA barcode.  6.2 Application of DNA barcoding to species identification of marine interstitial cercozoans    One of my thesis goals was to explore the biodiversity of cercozoans living in marine benthic habitats (Chapter 2).  However, such an investigation could potentially underestimate the actual diversity, if high similarities in morphological and behavioral features among the   170 examined organisms obscure the identities of separate lineages.  The marine cercozoans found during my study demonstrated this scenario and therefore served as an ideal system to to test the applicability of a DNA barcoding approach. The universal presence and ease of amplification of SSU rRNA gene help make this gene a suitable molecular marker for studying uncultivated lineages.  I examined SSU rDNA sequences of 17 marine cercozoans, whose morphology and behavior are very similar, and evaluated the potential of these sequences as DNA barcodes.  My investigation demonstrated that a 618-bp region at the 5' fragment of the SSU rDNA sequences can serve as a diagnostic tool for species delimitation.  Nucleotide sequence analysis of this region showed high intergeneric sequence divergences of about 7% and very low intraspecific sequence divergences of 0-0.5%.  Furthermore, phylogenetic analyses inferred from this barcoding region showed very similar tree topologies to those inferred from the full length of the gene. Overall, my study indicates that the 618-bp barcoding region of SSU rDNA sequences is a useful molecular signature for understanding the biodiversity and interrelationships of marine benthic cercozoans.  6.3 Morphostasis within the Cercozoa: a case study of Clautriavia vs. Protaspis Several taxa of benthic cercozoans had dorsoventrally flattened cell shapes, feeding by means of pseudopodia that emerge from a ventral slit or aperture, and gliding motility via heterodynamic flagella.  These features are well suited for living in interstitial environments and are shared by several distantly related cercozoan subclades, namely Cryomonadida, Discomonadidae, Ventrifissuridae, Verrucomonadidae, and a lineage of Clautriavia (Chapter 2).  One of the best (or most obvious) examples of morphostasis reported in my thesis was found between Clautriavia and Protaspis (Chapters 2 and 3).   171 Clautriavia is a genus of uncertain taxonomic affinity that was initially described as comprised of gliding cells with one prominent trailing flagellum and a mid-ventral groove (Massart 1900).  The genus was classified either with euglenids on the basis of similar paramylon-like granules or with cercozoans, specifically Protaspis spp., on the basis of general similarities in cell morphology and behavior (Larsen and Patterson 1990; Walton 1915).  In my study, I isolated and cultivated a novel species of Clautriavia, namely C. biflagellata n. sp., from marine sand samples collected from the west coast of Vancouver Island, Canada and characterized this isolate with high resolution microscopy (LM, SEM, and TEM) and small subunit (SSU) rDNA sequences.  The gliding cells of C. biflagellata were round to oval in outline, dorsoventrally flattened, and capable of engulfing other eukaryotic cells (e.g., diatoms).  The cells possessed two recurrent flagella of unequal length that emerged from a subapical pit within a ventral depression. Although Protaspis and Clautriavia are strikingly similar to one another in general morphology and behaviour, molecular phylogenetic analyses demonstrated that C. biflagellata was only distantly related to Protaspis spp.  This phylogeny is consistent with reproductive behavior and ultrastructural details.  For example, C. biflagellata performs asexual reproduction by longitudinal binary fission along the mid-sagittal plane; in contrast, asexual division in Protaspis occurs along the frontal plane (Skuja 1948).  In addition, at the ultrastructural level, cells of Protaspis are surrounded by a thick multilayered cell wall that does not contain pores (Hoppenrath and Leander 2006); by contrast, the cell surface of C. biflagellata lacks a cell wall, and contains an interspersed distribution of pores.  Moreover, unlike that of Protaspis, C. biflagellata does not contain nuclei with conspicuously condensed chromosomes or a cytoplasm containing batteries of extrusomes (Hoppenrath and Leander 2006).  Overall, my phylogenetic analysis and comparative ultrastructural data   172 showed that C. biflagellata is closely related to the recently established Auranticordida clade, consisting of Pseudopirsonia mucosa and Auranticordis quadriverberis.  6.4 Ultrastructural characterization and molecular phylogeny of Auranticordis quadriverberis n. gen. et sp. I discovered and characterized a rarely encountered, marine tetraflagellate, Auranticordis quadriverberis n. gen. et sp., isolated from sand samples collected from a tidal flat near Vancouver, Canada (Chapter 4).  These flagellates possessed several novel cellular features, such as (1) inverted heart-shaped cell appearance with some degree of cell plasticity, (2) gliding motility associated with four tightly bundled homodynamic recurrent flagella (the second known tetraflagellate cercozoan), (3) bright orange color due to possession of linear arrays of muciferous bodies distributed over the entire cell surface, and (4) inconspicuous mitochondria that lacked cristae.  Each cell also possessed about 2-30 pale orange bodies that were enveloped by two membranes and also surrounded by sac-like vesicles.  To our knowledge, these structures have never been described in any other lineage of eukaryotes.  The innermost membrane surrounding the pigmented bodies invaginated to form a dense array of unstacked thylakoids that extended toward a central pyrenoid-like inclusion containing tailed viral particles (known to infect cyanobacteria).  The general ultrastructure of these pigmented bodies was most consistent with putative photosynthetic endosymbionts of cyanobacterial origin (Kies 1974; Kies and Kremer 1979, 1990).  The phylogenetic analysis based on SSU rDNA sequences of A. quadriverberis demonstrated that this flagellate is a cercozoan and is most closely related to nanoflagellate parasites of diatoms, namely Pseudopirsonia mucosa, and several anoxic cercozoans known only from environmental sequencing surveys (Chantangsi et al. 2008).  The novel marine benthic   173 cercozoan C. biflagellata was also closely related to A. quadriverberis, which was in agreement with the ultrastructural data reported in Chapters 3 and 4.  Both lineages share: (1) porose cell surface; (2) a distinct layer of muciferous bodies immediately under the cell surface; (3) a robust microtubular root closely associated with the anterior end of the nucleus; (4) lack of a thick cell covering; and (5) the absence of conspicuously condensed chromosomes.  6.5 Phylogeny of the Cercozoa as inferred from polyubiquitin insertions and combined SSU and LSU rDNA data Most studies of phylogenetic relationships among the Cercozoa have been based largely on SSU rDNA sequences and those deduced phylogenies remain unresolved or gain very low support, especially at deep branching levels (Cavalier-Smith and Chao 2003). Concatenated datasets comprising both SSU and large subunit (LSU) rDNA sequences provide more robust phylogenies for relationships among several major groups of eukaryotes (Moreira et al. 2007).  I collected DNA sequences from both polyubiquitin and LSU rRNA genes from several different (mostly uncultivated) marine cercozoans, especially those dwelling in benthic environments (Chapter 5).  My investigation showed that the phylogenies constructed from combining both SSU and LSU rDNA sequences gained higher support values and improved relationships than those inferred solely from SSU rDNA sequences. A unique insertion of one or two amino acids at the intermonomeric junctions of the highly conserved polyubiquitin molecule is shared only by two very diverse protistan assemblages, the Cercozoa and the Foraminifera (Archibald and Keeling 2004; Archibald et al. 2003; Bass et al. 2005).  Possession of one amino acid residue at the polyubiquitin junctions, like that found in foraminiferans and some cercozoans, has been suggested to be an   174 ancestral state while possession of two residues represents a derived state.  Using the concatenated gene phylogeny as a framework, I demonstrated that the gain and loss of amino acid residues between polyubiquitin monomers happened several times independently.  I also helped demonstrate that the specific amino acid residue inserted (e.g., serine) reflects the origins of several cercozoans subclades.  6.6 Future directions My study underscores how poorly we understand the actual cellular diversity, ecology and phylogeny of the Cercozoa.  Several novel taxa were described in my investigation at both morphological and molecular levels.  Although most of the examined cercozoans were phylogenetically distant to one another, these lineages shared many features for living in benthic environments that can be interpreted as examples of either convergence or morphostasis.  However, some cercozoans possessed novel and unexpected features, such as the strikingly orange color and possession of possible primary endosymbionts in Auranticordis quadriverberis.  Nonetheless, I also demonstrated an alternative approach for the delimitation of species with similar morphologies by using a specific region of the SSU rRNA gene as DNA barcode.  My investigation also showed that ultrastructural homologies, which are not obvious at the light microscopic level, are evident by using transmission electron microscopy and molecular phylogenetics on relatively dissimilar looking cercozoans, like A. quadriverberis and C. biflagellata.  Lastly, the concatenation of SSU and LSU rDNAs with data on polyubiquitin insertions helped improve the overall phylogenetic framework for understanding cercozoan diversity. Estimations of cercozoan diversity weighed against current knowledge of the group indicate that the vast majority of cercozoan lineages remain to be discovered and   175 characterized.  As suggested by some studies, soil cercozoans might make up over half of the total protozoan biomass in these environments (Arndt et al. 2000); a similar ratio might also occur in marine benthic ecosystems.  Therefore, additional exploratory studies of cercozoan diversity are needed to more comprehensively understand the ecological and phylogenetic patterns in such environments.  These insights can be expedited by the utilization of a DNA barcoding approach.  In addition, improved knowledge of cercozoan diversity will require the ultrastructural characterization of representatives belonging to several different lineages; however, my study indicates that this line of research will be severely constrained by the rarity and uncultivability of many distinct cercozoans.  Moreover, my study also indicates that continued attempts to discover and cultivate novel cercozoans will almost certainly impact our overall understanding of endosymbiosis and eukaryotic cell evolution in general. Finally, the use of genome-scale analyses of some of the culturable cercozoan taxa will undoubtedly provide important insights into the evolutionary history of the Cercozoa and its relationship to other eukaryote supergroups.   176 6.7 References Archibald JM, Keeling PJ (2004) Actin and ubiquitin protein sequences support a cercozoan/foraminieran ancestry for the plasmodiophorid plant pathogens. J Eukaryot Microbiol 51: 113-118  Archibald JM, Longet D, Pawlowski J, Keeling PJ (2003) A novel polyubiquitin structure in Cercozoa and Foraminifera: evidence for a new eukaryotic supergroup. Mol Biol Evol 20: 62-66  Arndt H, Dietrich D, Auer B, Cleven E-J., Gräfenhan T, Weitere M, Myl'nikov AP (2000) Functional diversity of heterotrophic flagellates in aquatic ecosystems. In Leadbeater BSC, Green JC (eds) The flagellates. Taylor and Francis, London, pp 140-268  Bass D, Cavalier-Smith T (2004) Phylum-specific environmental DNA analysis reveals remarkably high global biodiversity of Cercozoa (Protozoa). Int J Syst Evol Microbiol 54: 2393-2404  Bass D, Moreira D, López-García P, Polet S, Chao EE, von der Heyden S, Pawlowski J, Cavalier-Smith T (2005) Polyubiquitin insertions and the phylogeny of Cercozoa and Rhizaria. Protist 156: 149-161  Bass D, Chao EE, Nikolaev S, Yabuki A, Ishida K, Berney C, Pakzad U, Wylezich C, Cavalier-Smith T (2009) Phylogeny of novel naked Filose and Reticulose Cercozoa: Granofilosea cl. n. and Proteomyxidea revised. Protist 160: 75-109  Brad T, Braster M, van Breukelen BM, van Straalen NM, Röling WF (2008) Eukaryotic diversity in an anaerobic aquifer polluted with landfill leachate. Appl Environ Microbiol 74: 3959-3968  Cavalier-Smith T, Chao EE (2003) Phylogeny and classification of Phylum Cercozoa (Protozoa). Protist 154: 341-358  Chantangsi C, Esson HJ, Leander BS (2008) Morphology and molecular phylogeny of a marine interstitial tetraflagellate with putative endosymbionts: Auranticordis quadriverberis n. gen. et sp. (Cercozoa). BMC Microbiol 8: 123  Chen M, Chen F, Yu Y, Ji J, Kong F (2008) Genetic diversity of eukaryotic microorganisms in Lake Taihu, a large shallow subtropical lake in China. Microb Ecol 56: 572-583  Hoppenrath M, Leander BS (2006) Dinoflagellate, euglenid or cercomonad? The ultrastructure and molecular phylogenetic position of Protaspis grandis n. sp. J Eukaryot Microbiol 53: 327-342    177 Kies L (1974) Elektronenmikroskopische Untersuchungen an Paulinella chromatophora Lauterborn, einer Thekamöbe mit blau-grünen Endosymbionten (Cyanellen). Protoplasma 80: 69-89  Kies L, Kremer BP (1979) Function of cyanelles in the thecamoeba Paulinella chromatophora. Naturwissenschaften 66: 578-579  Kies L, Kremer BP (1990) Phylum Glaucocystophyta. In Margulis L, Corliss JO, Melkonian M, Chapman DJ (eds) Handbook of Protoctista. Jones and Bartlett, Boston, pp 152-166  Larsen J, Patterson DJ (1990) Some flagellates (Protista) from tropical marine sediments. J Nat Hist 24: 801-937  Massana R, Pedrós-Alió C (2008) Unveiling new microbial eukaryotes in the surface ocean. Curr Opin Microbiol 11: 213-218  Massart J (1900) Clautriavia, un nouveau genre de flagellates. Bulletin des Séances, Societé royale des sciences médicales at naturelles de Bruxelles. 58: 133-134  Moreira D, von der Heyden S, Bass D, Lopez-Garcia P, Chao E, Cavalier-Smith T (2007) Global eukaryote phylogeny: combined small- and large-subunit ribosomal DNA trees support monophyly of Rhizaria, Retaria and Excavata. Mol Phylogenet Evol 44: 255- 266  Park SJ, Park BJ, Pham VH, Yoon DN, Kim SK, Rhee SK (2008) Microeukaryotic diversity in marine environments, an analysis of surface layer sediments from the East Sea. 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